Handbook of Neurochemistry and Molecular Neurobiology: Neurotransmitter Systems [3 ed.] 9780387303512, 9780387303826

Table of contents :
Front Matter....Pages i-viii
Back Matter....Pages 1-20
....Pages 21-73

Citation preview

1

Structural Organization of Monoamine and Acetylcholine Neuron Systems in the Rat CNS

L. Descarries . N. Mechawar

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

2 2.1 2.2 2.3

Dopamine (DA) Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mesocortical DA System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mesostriatal DA System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Diencephalospinal DA System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 3 3 4

3 Noradrenaline (NA) Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 3.1 Coeruleocortical NA System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 3.2 Myelencephalospinal NA System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 4

Adrenaline Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6

5 5.1 5.2 5.3

Serotonin (5-HT) Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rapheocortical 5‐HT System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rapheostriatal 5‐HT System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rapheospinal 5‐HT System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

6

Histamine Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7

7 7.1 7.2 7.3 7.4

Acetylcholine (ACH) Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Basalocortical ACh System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Septohippocampal ACh System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Neostriatal ACh Innervation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Spinal ACh Innervation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

8 8.1 8.2 8.3 8.4

Developmental Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 Dopamine Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 Noradrenaline Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12 Serotonin Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12 Acetylcholine Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

9

Concluding Remarks: A New Image of the Neuron . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

#

2008 Springer ScienceþBusiness Media, LLC.

6 6 7 7

2

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

Abstract: The anatomical, cytological, and ultrastructural features of monoamine (dopamine, noradrenaline, adrenaline, serotonin, histamine) and acetylcholine neuron systems have been examined in many regions of mammalian central nervous system, particularly in the rat. By considering these data with an emphasis on innervation densities and ultrastructural relationships, including results obtained during postnatal development, organizational principles and characteristics emerge for each of the modulatory systems. List of Abbreviations: ChAT, choline acetyltransferase; DA, dopamine; DBH, dopamine‐b‐hydroxylase; 5‐HT, 5‐hydroxytryptamine; NA, noradrenaline; PNMT, phenylethanolamine N‐methyltransferase; TH, tyrosine hydroxylase

1

Introduction

Nowadays, neuromodulation may be broadly defined as all actions of neuronally released compounds that produce more than a direct, short‐lived effect on neuronal firing. In this sense, all known neurotransmitters, including the neuropeptides, the purine nucleotides and nucleosides, as well as the gaseous compounds NO and CO, presumably qualify as neuromodulators. In this chapter, however, a more conventional definition is adopted, whereby neurotransmitters exert transient effects through receptors mostly confined to synaptic junctions, whereas neuromodulators may act for longer periods of time through receptors that are mostly located away from release sites. In this view, the major neuromodulatory systems are the catecholamine (dopamine (DA), noradrenaline (NA), and adrenaline), serotonin (5‐HT), histamine, acetylcholine (ACh), and neuropeptide systems, as opposed to the amino acid systems (glutamate, aspartate, glycine, and GABA). Purines, NO, and CO need not be considered as forming systems on their own, since these compounds appear to be always colocalized with modulators and transmitters. Such distinctions are merely operational, however, since it is becoming increasingly clear that one or more of the modulators and/or transmitters are most often coexistent in the same neurons. Owing to their progressive characterization by a variety of chemoanatomical techniques, and particularly fluorescence histochemistry, uptake autoradiography, and immunocytochemistry, the morphological features of the central monoamine and ACh systems can be currently described at three levels of morphological organization: (1) their overall anatomical distribution of constitutive cell bodies of origin and axonal projections throughout the CNS; (2) their regional and intraregional (subnuclear or laminar) distribution of axon terminals (or varicosities) in different territories of innervation; and (3) their ultrastructural characteristics, intrinsic and relational, of putative release sites in the various brain regions. This chapter examines the monoamine and ACh systems of rat brain from this triple standpoint, including data on their development. It focuses mainly on rat, as knowledge in this species is the most complete and detailed. The purpose is not to be exhaustive, but to illustrate some principles as well as organizational features prevailing within and between these systems. In conjunction with the increasing amount of data being currently accrued on the cellular and subcellular distribution of the multiple receptors for each of the neuromodulators, it is thus expected to gain insights into their complementary modes of action and functional properties.

2

Dopamine (DA) Neurons

The general organization of the dopamine (DA) system is rather compartmentalized compared with that of the other monoamines (for a detailed description and bibliographical listing, see Bjo¨rklund and Lindvall, 1984). In the rat CNS, at least six DA projection subsystems have been described—mesostriatal, mesolimbocortical, diencephalospinal, periventricular, incertohypothalamic, and tuberohypophyseal, in addition to DA interneurons in both the olfactory bulb and retina. The mesostriatal and mesolimbocortical DA systems originate from three mesencephalic groups of cell bodies located in the retrorubral field, substantia nigra, and ventral tegmental area, respectively designated as A8, A9, and A10 according to the nomenclature proposed by Dahlstro¨m and Fuxe (1964) at the time of their initial description. The other cell groups giving

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

1

rise to DA projections, A11 to A15, occupy various parts of the diencephalon. The most caudal, A11, is at the origin of a long descending projection to the spinal cord, whereas the other groups give rise to short projections to the diencephalon and circumscribed hypothalamic and hypophyseal areas. The DA neurons in the olfactory bulb (group A16) are a subset of the periglomerular interneuron population, and thus mostly found in the periglomerular area, with a few also scattered in the outer plexiform layer. In the retina, the A17 group consists of a relatively sparse subpopulation of amacrine cells interspersed in the inner portion of the inner nuclear layer. Thus, the DA systems are constituted by various cell types, from anaxonic to long axon neurons. For a detailed mapping of these cell groups, see Ho¨kfelt et al. (1984b). The number of DA cells in the mesencephalic tegmentum has been estimated at 15,000–20,000 on each side of rat brain (Hedreen and Chalmers, 1972; Guyenet and Crane, 1981; Swanson, 1982): some 9,000 in the ventral tegmental area (Swanson, 1982) and the remainder in the zona compacta of the substantia nigra and retrorubral field. These cells are at the origin of extensive mesotelencephalic projections. Neurons of the A10 group supply the mesolimbocortical system innervating structures such as the amygdala, septum, olfactory tubercle, nucleus accumbens and cerebral cortex, whereas those of the A8 and A9 groups are the main contributors of the nigrostriatal system.

2.1 Mesocortical DA System The DA projection to cerebral cortex is restricted in its distribution, at least in rat. This DA innervation was initially described as confined to the anteromedial or prefrontal, anterior cingulate or pre‐and supragenual, suprarhinal, perirhinal, piriform, and entorhinal cortex (for detailed references to the literature, see Descarries et al., 1987). Additional DA terminal fields were later identified in the dorsomedial frontal area, retrosplenial and adjacent occipital cortex, and in the deep layers of the frontal, parietal, temporal, and occipital neocortex (Descarries et al., 1987). In each of these areas, there is a strong predilection of the DA innervation for certain cortical layers. Moreover, axonal tracing studies have indicated that individual DA neurons innervating the cerebral cortex have fairly circumscribed intracortical territories of projection and do not collateralize extensively (Fallon and Loughlin, 1982; Swanson, 1982; Albanese and Minciacchi, 1983; Loughlin and Fallon, 1984). The heterogeneous distribution of the mesocortical DA subsystem is substantiated by the available data on the number of DA terminals in the different regions of rat cerebral cortex, ranging from 4
104 in layer VI of the occipital cortex to 3.1
106 in layers II–III of the supragenual cingulate cortex (Descarries et al., 1987). Basic neurochemical parameters have been deduced from these numbers. Assuming that cortical DA is mostly concentrated within the varicosities as opposed to intervaricose axon segments, it has been calculated that, depending on the cortical region, the average DA content of a single varicosity should range from 0.6 to 1.7
104 pg of DA for concentrations of 290–810 mg/g or 1.9–5.4
103 M. Interestingly, the figures for mediofrontal and cingulate cortex were very similar to those extrapolated for neostriatal DA varicosities (Doucet et al., 1986). All available observations suggest important regional and perhaps laminar differences in the frequency with which DA axon varicosities in adult rat cerebral cortex make synaptic specializations. Thus, in the anteromedial and the occipital cortex, these axon terminals appear to be mostly if not entirely synaptic (Se´gue´la et al., 1988; Papadopoulos et al., 1989), whereas in the suprarhinal cortex, only 56% display a junctional complex. Relatively low synaptic incidences of 39% and 20% have also been reported for DA terminals in monkey prefrontal and entorhinal cortex, respectively (Smiley and Goldman‐Rakic, 1993; Erickson et al., 2000). In terms of DA function, the significance of such differences between cortical regions has yet to be investigated.

2.2 Mesostriatal DA System Studies of anterogradely labeled axons after injection of biotin dextran in single nigrostriatal neurons have shown that most of these axons travel directly to the striatum, in which they branch abundantly, whereas others branch only sparsely in the striatum and arborize profusely in various extrastriatal structures,

3

4

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

including globus pallidus and entopeduncular and subthalamic nuclei (Gauthier et al., 1999; see also Prensa and Parent, 2001). In dorsal neostriatum, the density of DA varicosities or terminals, i.e., potential release sites of DA, has been estimated at 1
108/mm3 in the striatal matrix and 1.7
108/mm3 in the striosomes and subcallosal streak (Doucet et al., 1986), which is generally assumed to represent one‐tenth of all terminals in the striatum. Since the volume of rat neostriatum is approximately 45 mm3, the total number of DA varicosities in one striatum must be at least 4.5
109, which represents at least 1660 DA varicosities per neostriatal neuron (for unbiased estimates of neuron number in striatum, see Oorschot, 1996). Then, depending on whether the number of mesencephalic DA neurons projecting to neostriatum is considered to be 3,500 (Ande´n et al., 1966) or 7,000 (Bjo¨rklund and Lindvall, 1984), these individual neurons must be endowed, on average, with 1.3 or 0.6
106 axon varicosities in the striatum, emphasizing the bushy character of their arborization. Furthermore, because only 30%–40% of these DA varicosities are junctional, and 60%–70% do not form synaptic membrane specializations (Descarries et al., 1996), it may be inferred that this DA subsystem operates in large part by diffuse as well as synaptic transmission. Because of the extreme density of this DA innervation, it has also been hypothesized that the spontaneous and evoked release from such a multitude of asynaptic as well as synaptic varicosities might permanently maintain a basal extracellular level of DA throughout the striatum Descarries et al., 1996. This should allow, among other functions and in addition to the transsynaptic effects of DA, for a sustained regulation of widely distributed high‐affinity receptors on neurons, glia, and microvascular elements.

2.3 Diencephalospinal DA System The descending DA projection to the spinal cord originates from a few hundred cells (group A11) located in the dorsal and posterior hypothalamus, paraventricular hypothalamic nucleus, zona incerta, and caudal thalamus (Bjo¨rklund and Skagerberg, 1979; Ho¨kfelt et al., 1979; Swanson et al., 1981). Axons of these DA neurons have been described as bifurcating into an ascending branch to the diencephalon and a descending branch to the spinal cord (Lindvall and Bjo¨rklund, 1974a). In the cord, these descending axons travel partly within lamina I of the dorsal horn and adjoining parts of the dorsal funiculus and partly around the central canal, spreading scattered terminals to the spinal grey at all segmental levels, mainly in laminae III–IV of the dorsal horn, the intermediolateral cell column, the periependymal region, and the ventral horn (Skagerberg et al., 1982; Shirouzu et al., 1990; Ridet et al., 1992). In the intermediolateral cell column and ventral horn, these axon terminals appear to be all synaptic, making junctions with cell bodies and dendrites (Ridet et al., 1992). Around the central canal (at thoracic level), more than two thirds are synaptic, whereas in the dorsal horn, two thirds at the cervical level and one fourth at the thoracic level do not form conventional synapses (Ridet et al., 1992). Thus, as in the cerebral cortex and neostriatum, the diencephalospinal DA system might operate at least partly by diffuse transmission, depending on the region innervated. It is not yet known whether the synaptic and asynaptic DA terminals in the different regions and/or at different levels of the cord arise from collaterals of the same cells.

3

Noradrenaline (NA) Neurons

The entire neuraxis, except for neostriatum, receives a NA innervation, issued from two major clusters of brainstem neurons, one in the locus coeruleus (A6) and its dorsolateral extension (A4) and the other in a series of smaller cell groups occupying the ventrolateral aspect of the medulla and pons (A1, A5, A7) (for a detailed description, see Moore and Card, 1984; see also Ho¨kfelt et al., 1984b). After dopamine‐b‐hydroxylase (DBH) immunostaining, the total number of these cells has been estimated at about 5,000 on each side of the brainstem (Swanson and Hartman, 1975). Noradrenergic neurons are also present in the nucleus tractus solitarii‐dorsal vagal complex and the area postrema (A2). Both the locus coeruleus and lateral tegmental NA groups give rise to ascending and descending projections. The locus coeruleus projects principally to the cerebral cortex, thalamus, cerebellum, and spinal cord, whereas the lateral tegmental groups projects principally to the basal forebrain, hypothalamus, brainstem, and spinal cord (Ungerstedt, 1971; Lindvall and Bjo¨rklund, 1974a, b, 1978).

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

1

3.1 Coeruleocortical NA System The locus coeruleus (A6), comprising approximately 1,500 neuronal cell bodies on each side of the brainstem (Descarries and Saucier, 1972; Swanson, 1976), is at the origin of the entire NA innervation of the cerebral cortex, including hippocampus (Ungerstedt, 1971; for detailed references to the literature, see Audet et al., 1988). A single coeruleocortical axon probably collateralizes from front to back throughout the cortex, infiltrating its whole thickness (Morrison et al., 1981; Nagai et al., 1981; Loughlin et al., 1982). Moreover, studies with retrogradely transported fluorescent dyes have convincingly demonstrated that at least some of the NA neurons innervating the cerebral cortex can concomitantly innervate other distant CNS regions such as the cerebellum or spinal cord (Ade`r et al., 1980; Nagai et al., 1981; Room et al., 1981; Steindler 1981). The intracortical distribution of NA terminals has been studied in considerable detail in rat and monkey. In rat neocortex, the NA axons and their varicosities have been shown to be rather uniformly distributed between the various cytoarchitectonic areas, whereas in primates, there might be a greater degree of regional and laminar heterogeneity (Levitt et al., 1984; Morrison et al., 1984). The average density of NA innervation has been estimated at 1.2
106 varicosities/mm3 of tissue in rat neocortex, with no statistically significant difference between the seven cortical areas examined in the anterior half of the brain (Audet et al., 1988). In every region, the number of NA terminals was greatest in the molecular layer and decreased progressively in the underlying cortex, with a two‐to threefold difference between the upper and lower layers. These numerical data allowed to estimate the possible number of cortical NA varicosities per locus coeruleus cell body of origin (at least 300, 000), their average number per cortical neuron (30–50), their actual incidence among all terminals in the cortex (1/1,000), and the mean endogenous amine content per varicosity (0.22 fg) (Audet et al., 1988). A similar quantitative study in the dorsal hippocampus revealed a more heterogeneous regional and laminar distribution and a significantly higher density of NA innervation, averaging 2.1
106 varicosities/mm3 (Oleskevich et al., 1989). On the basis of a hippocampal volume of 56 mm3, and assuming an equal share of hippocampal NA terminals per neuron, this should represent a further load of 78,400 terminals per locus coeruleus neuron. The number of NA varicosities per hippocampal neuron should range from 20 to 40 per granule cell in the dentate gyrus to 180 per pyramidal cell in CA3, and could represent one varicosity per 880–1,500 synapses, in the DG and CA1, respectively. Similar to neocortex, the NA content per varicosity should be in the order of 0.16–0.21 fg, for concentrations in the 102 M range. In both neocortex and hippocampus, the synaptic incidence of NA varicosities has been shown to be very low, with reported frequencies of 17% or 26% in the parietal cortex (Smiley et al., 1992; Se´gue´la et al., 1990), 7% in the frontal cortex (Cohen et al., 1997), and 16% in the CA1 region of dorsal hippocampus (Umbriaco et al., 1995). A single study in monkey provided a value of 18% for prefrontal cortex (Aoki et al., 1998). Thus, there seems to be a principle of coherence at stake here, whereby a highly divergent projection system such as the coeruleocortical NA system establishes rather loose interrelationships at the ultrastructural level as well, whereas more compartmentalized and more focused projections, such as the mesocortical DA system, will establish more frequent, and perhaps more rigid, synaptic connections (Descarries et al., 1988). Because of its widespread distribution as well as largely asynaptic character, this NA system appears ideally built to act at a distance, on vast neuronal ensembles, and exert rather general, sustained and/or indirect or mediated effects, as expected from a neuromodulator.

3.2 Myelencephalospinal NA System The NA innervation of spinal cord provides further evidence of the extreme divergence of locus coeruleus neurons. Numerous studies have indicated that, although the medullary NA cell groups contribute to this innervation, its principal source is the locus coeruleus (A6) and the A5 and A7 lateral tegmental cell groups (Westlund et al., 1981, 1982, 1983; Fritschy and Grzanna, 1990). The locus coeruleus and adjacent subcoeruleus NA neurons supply input to both the dorsal and ventral horns at all segmental levels, whereas the majority of the NA innervation of the intermediolateral cell column appears to arise from the A5 and A7 groups. The density of this spinal innervation is much greater than that of its DA counterpart. Yet, as

5

6

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

in the case of the DA system, only 25%–29% of the NA terminals in the dorsal horn would display junctional specializations, whereas a much greater proportion (87% and 85%) is synaptic in the intermediolateral cell column and ventral horn, respectively (Rajaofetra et al., 1992; Ridet et al., 1993).

4

Adrenaline Neurons

Neurons that are immunopositive not only for the biosynthetic enzymes tyrosine hydroxylase (TH) and DBH but also for phenylethanolamine N‐methyltransferase (PNMT), and which presumably synthesize and release adrenaline (Ho¨kfelt et al., 1974), also give rise to long and widely collateralized, albeit less abundant, axonal projections distributed from the forebrain to the spinal cord (for detailed description, see Ho¨kfelt et al., 1984a). The cell bodies of these neurons form three small groups, C1, C2, and C3, confined to medulla oblongata and located near and within the A1 and A2 NA cell groups and on the midline, respectively, within and dorsal to the medial longitudinal fasciculus. PNMT‐immunopositive fibers and nerve endings have been described as concentrated along the ventricular system and most abundant in the bed nucleus of the stria terminalis, various nuclei of hypothalamus, periaqueductal grey matter, brainstem nuclei of visceral afferent and efferent systems, locus coeruleus, and intermediolateral cell column of the spinal cord. In most of these CNS regions, the fine structural features and relationships as well as functional properties of the adrenergic nerve terminals remain to be characterized. In view of its anatomical distribution, this system is assumed to play a significant role in neuroendocrine mechanisms and blood pressure control. In adult rat spinal cord, PNMT‐immunopositive axon terminals have been shown to establish axosomatic and axodendritic synaptic contacts with the preganglionic neurons of the intermediolateral cell column (Milner et al., 1988).

5

Serotonin (5-HT) Neurons

The serotonin (5‐hydroxytryptamine; 5‐HT) system is even more widespread than the NA system, as there is no single region of CNS without a 5‐HT innervation, including each of the circumventricular organs and the cerebroventricular cavity, which is lined by the so‐called supra‐ependymal plexus of varicose 5‐HT fibers (Steinbusch, 1981). Owing to the variety of methodological approaches applicable for the identification and examination of central 5‐HT neurons, this is undoubtedly the transmitter‐defined system about which the most is known in terms of distribution, cytological features, and particularly, ultrastructural relationships. The 5‐HT neuronal cell bodies are mainly found near or in the midline or raphe region of the medulla, pons, and mesencephalon, in nine groups designated B1–B9 according to Dahlstro¨m and Fuxe’s nomenclature (1964). The more caudal groups (B1, B2, and B3 in raphe pallidus, obscurus, and magnus) project mostly to the medulla and spinal cord, whereas the most rostral (B5–B9, in raphe medianus, dorsalis, and the supralemniscal region) provide extensive innervation to the diencephalon and telencephalon. 5‐HT cell bodies have also been found in the area postrema, caudal locus coeruleus, and nucleus interpeduncularis. The nucleus raphe dorsalis is the most prominent, with more than 11,000 5‐HT neurons, representing approximately one‐third of its entire neuron population (Descarries et al., 1982). It is also the best characterized in terms of the cytological features of its constituent neurons and afferent and efferent connectivity. Much as the locus coeruleus NA neurons, it is likely that some of the nucleus raphe dorsalis 5‐HT neurons are highly collateralized and simultaneously project to vast areas of forebrain distant from one another (Fallon and Loughlin, 1982).

5.1 Rapheocortical 5‐HT System The 5‐HT innervations of cerebral cortex, hippocampus, and neostriatum have been the most thoroughly examined. In seven cytoarchitectonic areas from the anterior half of the cerebral cortex, the density of regional and laminar 5‐HT innervation was quantified after uptake labeling of the 5‐HT varicosities in whole hemisphere sections incubated with tritiated 5‐HT in the presence of a monoamine oxidase inhibitor (Audet et al., 1989). The mean regional density of cortical 5‐HT innervation was thus estimated at 5.8
106

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

1

varicosities/mm3 of tissue, with significant variations between areas. The highest laminar density was always that of layer I, except in piriform cortex, but each region showed a distinct laminar pattern of 5‐HT innervation. On the basis of these figures, the average number of cortical 5‐HT varicosities per cell body of origin could be calculated to be at least 500,000, their average number per cortical neuron from 145 to 230, their incidence among all cortical axon terminals 1 in 200, and their mean endogenous amine content 0.045 fg for a concentration in the order of 3
103 M. A similar study carried out in the subiculum, Ammon’s horn, and dentate gyrus of the dorsal hippocampus yielded values of 2.7
106 varicosities/mm3 for the average density, but with the values in subiculum > Ammon’s horn > dentate gyrus, and a marked heterogeneity in laminar distribution (Oleskevich and Descarries, 1990). The average number of hippocampal 5‐HT varicosities per cell body of origin could thus be evaluated at 150,000, the number of 5‐HT varicosities per target neuron at 20–130, and the mean endogenous amine content per hippocampal 5‐HT varicosity at 0.05–0.07 fg, a value similar to that in cerebral cortex. In rat neocortex, a detailed study by Se´gue´la and coworkers (1989) has estimated the synaptic incidence of 5‐HT axon varicosities at 36% and 28% in the superficial and deep layers of the frontal cortex, 46% in the parietal cortex, and 37% in the occipital cortex. Even lower frequencies of synaptic specialization were reported for the sensorimotor and prefrontal cortex of monkey (DeFelipe and Jones, 1988; Smiley and Goldman‐Rakic, 1996) and for the cat auditory cortex (DeFelipe et al., 1991). Frequencies of synaptic 5‐HT varicosities ranging between 12% and 24% have also been reported for CA1, CA3, and the dentate gyrus of hippocampus (Oleskevich et al., 1991; Cohen et al., 1995; Umbriaco et al., 1995).

5.2 Rapheostriatal 5‐HT System In neostriatum, the 5‐HT innervation appears rather uniformly distributed, without any suggestion of a patch and matrix pattern. Its density increases from rostral to caudal, however, and is always higher ventrally than dorsally. It ranges from 4.8
106 varicosities/mm3 rostrally to 6.3
106 caudally, for an average of 5.6
106 (Mrini et al., 1995), almost equal to that in cerebral cortex. Such a density corresponds to approximately 90 5‐HT varicosities per neostriatal neuron, thus roughly 18 times less the number for DA terminals. It predicts a value in the order of 0.09 fg for the mean 5‐HT content per neostriatal 5‐HT varicosity, compared to 0.045 fg and 0.06 fg in the cerebral cortex and hippocampus, respectively. The proportion of these 5‐HT varicosities engaged in synaptic contact has been estimated at 10%–13%. In the rostral striatum, for example, this small proportion should correspond to some 4.8
105 synapses/mm3, i.e., approximately 1 in every 2000 striatal synapses.

5.3 Rapheospinal 5‐HT System The 5‐HT innervation of spinal cord is relatively dense, with particular regions of the spinal grey standing out because of their strong 5‐HT innervation: the dorsal horn, particularly lamina I and to a lesser extent lamina II, the ventral horn motor nuclei (laminae VIII and IX), and the intermediolateral cell column in the thoracic cord (Bowker et al., 1982; Skagerberg and Bjo¨rklund, 1985). As in the case of the DA and the NA innervations of the dorsal horn, the 5‐HT innervation in this area has been shown to be only partly synaptic (37%) (Ridet et al., 1993), with little variation between the different laminae of the dorsal horn or at different spinal cord levels (Marlier et al., 1991). In the intermediolateral cell column and anterior horn, however, the 5‐HT varicosities are presumably mostly if not entirely synaptic (Poulat et al., 1992; Ridet et al., 1993), as also reported for DA and NA axon varicosities.

6

Histamine Neurons

Immunocytochemical studies with antibodies against histidine decarboxylase or histamine itself have revealed the existence in the rat brain of a widely distributed network of fine, varicose, unmyelinated axons, originating from small subgroups of nerve cell bodies located in the tuberomammillary nucleus

7

8

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

(Watanabe et al., 1984; Inagaki et al., 1988; Panula et al., 1989). This system comprises long projections to regions such as the olfactory bulb, cerebral cortex, caudate‐putamen, and thalamus, a relatively dense innervation of numerous areas of hypothalamus, and descending projections to the upper and lower brainstem, cerebellum, and spinal cord. To date, the only light and electron microscopic immunocytochemical description of these nerve fibers and axon varicosities has shown that in cerebral cortex and neostriatum the majority of histamine varicosities do not form synaptic specializations (Takagi et al., 1986).

7

Acetylcholine (ACH) Neurons

Our present knowledge on the structural basis of ACh transmission in the CNS has been largely acquired from the detailed examination of neurons immunostained for choline acetyltransferase (ChAT), the rate‐ limiting enzyme for ACh synthesis. All regions of the CNS are pervaded by dense networks of ChAT‐ immunostained axons originating either from projection neurons located in the basal forebrain or midbrain, and/or from local interneurons (Armstrong et al., 1983; Woolf, 1991). The latter have been shown to contribute either a small fraction (neocortex), almost all (neostriatum), or the totality (spinal cord) of respective regional ACh innervations. Groups of ACh projection neurons and their targets have been extensively described from investigations combining tract‐tracing methods and ChAT immunocytochemistry (Rye et al., 1984; Saper, 1984) and are now commonly referred to as Ch1–Ch8 on the basis of nuclear localization (Mesulam et al., 1983; Mesulam, 1988). The basal forebrain groups Ch1–Ch4 provide for the rich and widespread innervations of neocortex, hippocampus, olfactory bulb, and amygdala. The medial septum and nucleus of the vertical limb of the diagonal band of Broca, Ch1 and Ch2, respectively, send dense ACh projections to the hippocampal formation, while the lateral portion of the horizontal limb of the diagonal band of Broca (group Ch3) innervates mainly the olfactory bulb. Together, groups Ch1–Ch3 also contribute a limited fraction of the total cortical ACh innervation, with axons restricted to limbic areas (cingulate, entorhinal, orbitofrontal and piriform cortex). In contrast, the whole cortical mantle (including these limbic areas) and amygdala receive a rich axon network stemming from the Ch4 neurons in the nucleus basalis magnocellularis of Meynert, which is spread over the substantia innominata and globus pallidus. The pontomesencephalic ACh system (Ch5–Ch6) is the principal projection pathway outside the basal forebrain and has most of its efferents reaching the thalamus and basal forebrain. ACh neurons in the habenula constitute the Ch7 system, which extends projections exclusively to the interpeduncular nucleus via the fasciculus retroflexus pathway. Finally, the parabigeminal nucleus group Ch8, also limited in its scope of innervation, targets the majority of its axons to the superior colliculus (tectum). The availability of a highly sensitive monoclonal antibody with high affinity for whole rat brain ChAT (Cozzari et al., 1990) has allowed the development of experimental conditions leading to the integral staining of ACh axon networks through the full thickness of perfusion‐fixed brain sections. This made it possible to conduct thorough and unbiased electron microscopic descriptions of ChAT‐immunostained axon varicosities (Umbriaco et al., 1994), and led to the development of a semicomputerized light microscopic method to quantify the length of axons and related number of varicosities from ChAT‐ immunostained axon networks (Mechawar et al., 2000). In consequence, our group has produced quantitative descriptions of the cortical, hippocampal, and neostriatal ACh innervations, both in terms of quantified distribution and ultrastructural features (Umbriaco et al., 1994, 1995; Contant et al., 1996; Mechawar et al., 2000; Aznavour et al., 2002). In this section, we mainly discuss these systems (basalocortical, septohippocampal, and neostriatal), as they have been the most thoroughly scrutinized of all central ACh innervations. The ACh innervation intrinsic to the spinal cord will also be described, as it allows for meaningful comparisons with the monoaminergic innervations of this region.

7.1 Basalocortical ACh System It is currently estimated that, in the rat brain, there are 7,000–9,000 Ch4 neurons projecting to the cortex (Rye et al., 1984; Gritti et al., 1993). Although the great majority of ACh axons in cerebral cortex

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

1

originate from the basal forebrain, a 20%–30% portion is contributed by intrinsic bipolar interneurons scattered throughout layers II–VI (Johnston et al., 1981; Eckenstein and Thoenen, 1983; Eckenstein and Baughman, 1984; Levey et al., 1984). In a detailed light microscopic description of ChAT‐immunostained axons distributed in the rat cerebral cortex, 13 different patterns of ACh innervation were identified that generally corresponded to functionally similar cortical areas (Lysakowski et al., 1989). This patterning has recently been associated with modality‐and region‐specific ACh release in neocortex (Fournier et al., 2004), indicating at least a regional control in Ch4 output activity. The laminar and regional densities of the cortical ChAT immunoreactive axon network have been estimated in transverse sections from the frontal (Fr1), parietal (Par1), and occipital (Oc1) cortical areas (Mechawar et al., 2000). The number of varicosities per unit length of axon was counted directly at the microscope and found to be constant throughout these areas (average of 4 varicosities/10 mm of axon). In consequence, the actual number of varicosities in the network could be directly derived from measurements of its length, and the laminar and regional densities of ACh innervation expressed in meters of axon and millions of varicosities per mm3 of tissue. The densest ACh innervation was thus measured in the frontal, followed by the occipital and the parietal cortex, with respective values of 5.4, 4.6, and 3.8
106 varicosities/mm3. In the three areas, layers I and V were the most densely innervated, with respective interareal means of 5.3 and 5.0
106 varicosities/mm3. The least densely innervated were layers IV and VI of the primary sensory areas, with interareal means (Par1 and Oc1) of 3.4 and 3.8
106 varicosities/mm3, respectively. As expected, the laminar distributions were area specific, and characterized by uniformly high densities throughout the frontal cortex and lower densities in layers II/III, IV, and VI of the parietal cortex, as well as in layers IV and VI of the occipital cortex. Compared to monoaminergic innervation densities previously measured in the same areas, the mean density of 4.6
106 ACh varicosities/mm3 for these regions represented the densest of all neuromodulatory inputs to the neocortex. When broken down to individual cells, these figures allow to calculate that, on average, each ACh neuron projecting to the cerebral cortex must be endowed with an axonal arborization totalling at least 0.5 m in length and bearing more than 200,000 varicosities. Moreover, this is likely to be an underestimate, since Ch4 neurons have also been shown to extend several axon collaterals within the basal forebrain, which make synapse onto dendrites of surrounding (unspecified) neurons (Zaborszky and Duque, 2000). This latter observation leads to the conclusion that the activity of basalocortical ACh neurons may itself be subjected to a dual cholinergic modulation: one intrinsic, from its own recurrent axon collaterals; and one extrinsic, from its ACh afferents of the mesopontine tegmentum. The relational features of cortical ACh axon terminals (varicosities) were first described in the primary somatosensory cortex of adult rat (Umbriaco et al., 1994). In all layers of Par1, only a small fraction of these ChAT immunoreactive varicosities were found to form a synaptic contact (junctional complex), i.e., 10%, 14%, 11%, 21%, and 14% in layers I, II/III, IV, V, and VI, respectively, for an interlayer mean of 14%. In general, cortical ACh varicosities were relatively small, and those bearing a synaptic junction were slightly but significantly larger than their nonsynaptic counterparts, i.e., 0.67 versus 0.57 mm in diameter, respectively. The junctional complexes formed by these terminals were single, occupied a small fraction of the total surface of varicosities (3%), and were almost always symmetrical (99%). The relatively few synapses made by ACh varicosities were always axodendritic, either on branches (75%) or on spines (25%). Subsequent investigations in other laboratories have confirmed that the vast majority of ACh varicosities in rat cortex are asynaptic, with reported estimates of 14% and 9% for the frontoparietal and entorhinal region, respectively (Che´dotal et al., 1994; Vaucher and Hamel, 1995). The value of 66% recently reported by Turrini and coworkers (2001) for layer V of rat parietal cortex after labeling with the vesicular ACh transporter was presumably the result of a sampling bias, as suggested by a significantly larger size of the profiles examined in that particular study. In the prefrontal cortex of rhesus monkey, Mrzljak et al. (1995) reported that 44 of 100 serially sectioned ChAT-immunoreactive axon varicosities at the border of layers II and III made synaptic contact, mostly onto small dendritic shafts. In a similar study of two samples of human anterior temporal lobe removed at surgery for epilepsy, Smiley et al. (1997) found 28 of 42 ACh varicosities from layer I and II endowed with small but identifiable synaptic specializations. Whether such variations of synaptic incidence reflect sampling biases, regional differences or species differences remains to be determined. In any event, these

9

10

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

data allow the conclusion that the actions of ACh depend on both diffuse and synaptic transmission in the neocortex of primates as well as of rat (further discussion in Descarries et al., 2002).

7.2 Septohippocampal ACh System The vast majority of ACh axons in the hippocampus originate from the estimated 7,150 ACh Ch1–Ch2 neurons in each septum (Cadete‐Leite et al., 2003). Indeed, very few bipolar ACh interneurons have been described in the hippocampus, most of which were observed in the stratum lacunosum moleculare of CA1 (Aznavour et al., 2002). As previously found for the neocortical innervation, the periodicity of varicosities along ACh axons remained constant at 4 per 10 mm throughout the dorsal hippocampus, suggesting that this is an intrinsic feature of ACh innervations. The densest laminar ACh innervations were measured in the stratum lacunosum moleculare of CA3, stratum pyramidale of CA1 and CA3, and stratum moleculare of the dentate gyrus, with respective values of 7.5, 8.2, 6.8, and 7.8 million varicosities/mm3 (Aznavour et al., 2002). Regional values were similarly high, ranging from 4.9
106 varicosities/mm3 in CA1 to 6.2
106 varicosities/mm3 in CA3, for an average of 5.9
106 varicosities/mm3 in hippocampus, a value 28% greater than for neocortex and higher than any neuromodulatory input to cortex described so far. Ultrastructural analysis of this innervation in the stratum radiatum of CA1 revealed additional common features with the basalocortical ACh system (Umbriaco et al., 1995). The ACh varicosities in this hippocampal region measured 0.6 mm on average, and only 7% were synaptic. Their few synaptic contacts were symmetrical and made either with dendritic branches or spines. Together with immunoelectron microscopic data demonstrating a predominant extrasynaptic localization of muscarinic ACh receptors in hippocampus or cerebral cortex (reviewed in Volpicelli and Levey, 2004), the above results strongly suggest that the modulatory effects of ACh on cortical function, so well documented in hippocampus, are largely conveyed by diffuse (or volume) transmission in addition to synaptic transmission. In both regions in which this innervation is relatively dense, many of these effects could also depend on the existence of a low ambient level of ACh, permanently maintained in the extracellular space in spite of the presence of acetylcholinesterase (for further discussion, see Descarries et al., 1997).

7.3 Neostriatal ACh Innervation The neostriatum receives by far the densest ACh innervation in the mammalian brain, as manifested by the high measures of cholinergic markers expressed throughout this region. Among others, values of ACh content (Cheney et al., 1975; Hoover et al., 1978), ChAT activity (Hoover et al., 1978), and choline uptake (Rea and Simon, 1981) are particularly elevated. These parameters reflect the profuse ACh axon network originating from cholinergic aspiny interneurons, which are estimated to account for less than 2% of all neostriatal neurons (Woolf and Butcher, 1981; Phelps et al., 1985). ACh interneurons in the caudate and putamen are large cells resembling their basal forebrain counterparts (Armstrong et al., 1983), and also give rise to fine unmyelinated axons periodically adorned with small, round, or ovoid varicosities. Although this local cell population provides the neostriatum with most of its cholinergic innervation, a minor fraction of ACh axons has been found to originate from the Ch5–Ch6 system (Woolf and Butcher, 1981, 1986). The ultrastructural features of these putative release sites were first described by Contant and coworkers (1996), after ChAT‐immunostaining in single thin sections for electron microscopy. As previously found in cerebral cortex and hippocampus, neostriatal ACh varicosities were seldom engaged in synaptic contact. Their frequency of junction of 3% in single sections amounted to 9% when extrapolated to the whole volume of varicosities. The very few ACh synapses were made with synaptic branches (6/10) or spines (4/10). Other axon terminals, unlabeled, were often directly apposed to neostriatal ACh varicosities. Occasional juxtaposition of ChAT‐immunostained varicosities was also observed. From this and a subsequent study in the developing brain (Aznavour et al., 2003), it was concluded that the diffuse mode of transmission was an inherent characteristic of ACh neurons, both as interneurons (neostriatum) and projection neurons (cerebral cortex).

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

1

7.4 Spinal ACh Innervation Unlike the DA, NA, and 5‐HT systems, there are no descending ACh projections from the brain to the spinal cord (Sherriff et al., 1991). Therefore, the spinal ACh innervation originates exclusively from the different categories of spinal ACh neurons, which have been thoroughly described by Barber and collaborators (1984). The preponderant type of ACh neuron in this region is the ventral horn somatic motor neuron, regarded as the canonical ACh neuron since Langley’s seminal work on neuromuscular activation one century ago. The only other ACh neurons with projections leaving the spinal cord are the preganglionic autonomic neurons, located in the intermediate grey matter at thoracolumbar and lumbosacral levels. The three other types of ACh cells in the spinal cord are the small laminae III–V neurons of the dorsal horn, lamina VII partition neurons at the border between the dorsal and ventral horns, and lamina X neurons in the central grey surrounding the central canal. Transverse sections of adult rat spinal cord immunostained for ChAT reveal that somatic motor neurons are organized in central, medial, and lateral motor columns, and that the medial and lateral columns can be further divided into five subcolumns (Barber et al., 1984). There are apparently widespread intra‐ and intercolumnar interactions, as suggested by extent of intermingling longitudinal and transverse motor dendrite bundles. Likewise, at autonomic levels, the somata and dendritic arborizations of partition cells and central canal ACh neurons are heavily mixed with those of autonomic neurons. These extensive interconnections between morphologically diverse ACh neurons have been described as the basis of a cholinergic propriospinal system (Sherriff and Henderson, 1994; Huang et al., 2000). A rich ACh axon network pervades all layers and regions of the spinal cord, including the ependymal cell layer (Phelps et al., 1984; Scha¨fer et al., 1998). These varicose axons originate in various (undetermined) proportions from the different types of ACh neurons mentioned above. Initial reports have shown ChAT‐ immunoreactive terminals to contact both ACh and non‐ACh elements in the spinal cord. The current prevalent view is that these terminals are mostly, if not exclusively, synaptic in nature. This view is likely biased by the fact that most studies on the subject have focused specifically on contacts made on the cell bodies and proximal dendrites of motoneurons, Renshaw cells, and sympathetic preganglionic neurons (Markham and Vaughn, 1990; Alvarez et al., 1999). Of these descriptions, the ones concerning the ACh innervation on motoneuron somatodendrites are of particular interest. This innervation, which is thought to arise from the canal cluster cells, consists of C‐type cholinergic terminals characterized by periodic postsynaptic specializations called subsurface cisterns (Nagy et al., 1993).

8

Developmental Aspects

The early development of monoamine and ACh neurons in mammalian CNS has led to the notion that these modulatory systems play significant roles in the installment and refinement of neuronal connectivity during brain maturation. In rat, for example, dopaminergic fibers enter the cortical plate just before birth (Kalsbeek et al., 1988); noradrenergic fibers reach the anterior parietal cortex around E20, and occipital cortex at birth (Verney et al., 1982); serotoninergic fibers and terminals (Seiger and Olson, 1973), as well as pioneering cholinergic fibers (Mechawar and Descarries, 2001), are already seen in the cortical plate at birth (for reviews, see Semba, 1992, 2004). Yet, only two laboratories have examined the morphological features of these systems at electron as well as light microscopic levels during development: the University of Thessaloniki group, in Greece, which has focused on the monoamine systems, and our own laboratory, in Montreal, which has mainly studied the developing cholinergic system.

8.1 Dopamine Neurons The ultrastructural features of DA neurons have been examined in two brain regions during development: lateral septum and striatum. The DA innervation of the lateral septum was reported to undergo a marked reorganization during the first two postnatal weeks, when it acquired features comparable to the adult (Antonopoulos et al., 1997). The ultrastructural analysis suggested that there might be two different DA inputs to this region: the first developing earlier in life, affecting remote parts of neurons through

11

12

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

symmetrical axodendritic synapses, and the second developing later and affecting neuronal somata through asymmetrical axosomatic synapses. In the striatum, the same authors described the DA innervation of caudate‐putamen and nucleus accumbens as exhibiting a similar type of synaptic connectivity throughout development, but evolving from symmetrical synapses mostly located on dendritic shafts at an early stage to a later stage where symmetrical axospinous synapses also became a prominent feature (Antonopoulos et al., 2002). Interestingly, this study also indicated that after initial rises to values >90% and 80% in the islands and matrix of caudate‐putamen, respectively, and almost 100% and 75% in the shell and core of the nucleus accumbens, the proportion of DA varicosities making synapses declined after P7 in the caudate‐putamen and after P14 in the nucleus accumbens to respective values of 63% and 35% in the islands and matrix of caudate‐ putamen, and 71% and 47% in the shell and core of nucleus accumbens.

8.2 Noradrenaline Neurons The noradrenergic innervations of the septum, motor and visual cortex, and dorsal lateral geniculate nucleus were described in developing rat brain, by means of light and electron microscopic immunocytochemistry with DBH antibodies (Latsari et al., 2002, 2004; Antonopoulos et al., 2004). In all four regions, few, relatively thick NA fibers were present at birth, which arborized gradually into an adult pattern of thinner varicose fibers by the second postnatal week, and reached the adult density of innervation one week later. Irrespective of the postnatal age examined, only a minority of these NA varicosities displayed synaptic specializations (10%–15%), which were usually symmetrical and found on dendritic branches. It was concluded from these data that, in these brain regions at least, transmission by diffusion is the major mode of NA action in the developing as well as adult brain.

8.3 Serotonin Neurons The growth of 5‐HT innervations was examined in numerous subcortical regions of postnatal rat brain: lateral septum (Dinopoulos et al., 1993), lateral ventricles (Dinopoulos and Dori, 1995), lateral geniculate (Dinopoulos et al., 1995), basal forebrain (Dinopoulos et al., 1997), superior colliculus, and ventrolateral thalamic nucleus (Dori et al., 1998). Acquisition of such data from the developing cortex (Dori et al., 1996) was complicated by the fact that, within the first weeks after birth, thalamocortical neurons transiently express 5‐HT plasma membrane transporter and vesicular monoamine transporter (Lebrand et al., 1996) and are thus immunoreactive for 5‐HT and indistinguishable from bona fide 5‐HT neurons. A constant feature of the 5‐HT innervations in the postnatal period, albeit parenchymal or intraventricular, was their progressive change from a few, thick and smooth unmyelinated axonal fibers at birth to a ramified and relatively dense network of fine varicose axons, infiltrating the whole region and reaching its adult pattern of distribution and density within the first three weeks. At the ultrastructural level, however, much more diversity was apparent. In the lateral septum, 5‐HT varicosities were described as almost always synaptic and showing symmetrical synapses, whether on somata, dendritic shafts, or spines. In the dorsal portion of the lateral septum, they formed characteristic pericellular basket‐like arrangements around cell somata and their primary dendrites, as previously described for DA terminals (Descarries and Beaudet, 1983). In the lateral ventricles, the 5‐HT varicosities were located close to the ventricular surface of the ependymal lining, but never made synapses on ependymal cells, even when morphologically mature (shape, size, and content). In the lateral geniculate and basal forebrain, the synaptic frequency of the 5‐HT varicosities displayed a biphasic temporal profile. The proportion of varicosities forming synapses was reported to increase from birth to the end of the second postnatal week, and then decline markedly in the third week before increasing again to adult values of about 40% in both regions. A similar biphasic pattern was also described in the superficial layers of the superior colliculus (involved in visual functions); whereas in its deep layers and in the ventrolateral thalamic nucleus (areas involved in motor functions), the proportion of 5‐HT varicosities engaged in synaptic contact showed a continuous increase from birth to adulthood, to become a fully synaptic innervation making mostly symmetrical synapses onto dendritic shafts.

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

1

8.4 Acetylcholine Neurons A recent description of the ACh innervation in the frontal, parietal, and occipital neocortex of the postnatal rat has revealed that this ACh system develops rapidly and much earlier than previously suspected (Mechawar and Descarries, 2001). At birth, a few ChAT‐immunostained fibers capped with growth cones are already seen throughout the cortical plate and marginal zone. By P4, faintly immunoreactive interneurons are first detected. Rapid ingrowth and proliferation ensues resulting in an adult‐like distribution of a highly elaborate network of fine varicose ACh axons by P8. In the parietal area, adult densities of innervation are reached by P16, while development continues until the end of the first month in the frontal area and even later in the occipital area. Two parameters of this ingrowth have been quantified (Mechawar and Descarries, 2001): the elongation (and branching) of ACh axons and their number of axon varicosities per unit length. This latter value was shown to increase steadily in all cortical layers and areas, doubling from 2 varicosities per 10 mm of axon at P4 to the adult ratio of 4 per 10 mm at P16. It was thus possible to calculate that, within the first two weeks after birth, a single basalocortical neuron produced an average of 2 cm of axon and 9,000 varicosities/day, i.e., almost 1 mm of axon and 400 varicosities/hour. A similar study has also been carried out in CA1, CA3, and the dentate gyrus of dorsal hippocampus (Aznavour et al., 2005). At P8, an elaborate network of varicose ChAT‐immunostained axons was already present in all three hippocampal regions. As in neocortex, the number of axon varicosities per unit length of ACh axon increased during the first two weeks after birth to reach the adult value of 4 per 10 mm at P16. At this age, the laminar distribution of this network resembled that of maturity, but adult densities of axons and axon varicosities were reached only by P32. Between P8 and P32, the mean densities in the three regions increased from 8.4 to 14 m of axons, and 2.3 to 5.7 million varicosities/mm3 of tissue. This suggested that, on average, the septohippocampal ACh neurons are capable of generating 7.5 cm of axon and 27,000 varicosities/day, i.e., more than 3 mm of axon and 1,120 varicosities/hour. Such growth rates are even higher than previously estimated for nucleus basalis ACh neurons innervating the neocortex, and emphasize the remarkable growth capacities of ACh neurons. These studies have also examined the intrinsic and relational features of ACh varicosities in the developing neocortex and hippocampus (Mechawar et al., 2002; Aznavour et al., 2005). In both regions, these varicosities were of similar size throughout development and only slightly smaller than in the adult. They were endowed with aggregated synaptic vesicles, and the frequency with which they showed a mitochondrion increased gradually with age, from about 20% at P8 to >40% at P32. As in the adult, the vast majority of these varicosities were asynaptic throughout the postnatal period. Since the proportion of synaptic ACh varicosities was stable during development, it could be inferred that the number of ACh synapses had already reached its adult value by the end of the second week, at least in the parietal cortex (0.55
106/mm3 at P16, compared to 0.53
106 at >P60). The postnatal growth and ultrastructural characteristics of a developing ACh innervation have also been examined in the neostriatum, where this innervation arises almost completely from a limited number of large interneurons. As in cortex and hippocampus, the invasion of neostriatum by ACh axons mostly takes place during the first two weeks after birth, but continues at a slower rate until the fourth postnatal week (Aznavour et al., 2003), in keeping with biochemical measurements of ChAT activity (Coyle and Yamamura, 1976). As in neocortex and hippocampus, the intrinsic and relational ultrastructural features of these rapidly growing axons were examined at three developmental time points, i.e., after the first, second, and fourth postnatal weeks (P8, P16, and P32) (Aznavour et al., 2003). Again, a low synaptic incidence was measured at all three ages, indicating that this structural feature is an intrinsic determinant of the functioning of this system during development as well as in the adult.

9

Concluding Remarks: A New Image of the Neuron

The detailed comparison of the structural attributes of neuromodulatory systems, whether at the level of their general organization, regional distribution, or fine structure, underscores some consistent features of the major modulatory systems. First and foremost, it requires a drastic revision of what may be called the

13

14

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

traditional (textbook) image of the neuron, mostly inherited from the study of the neuromuscular junction. Thus, relatively dense and widespread innervations by fine, unmyelinated axons whose multiple branches bear innumerable varicosities (terminals) appears to be the rule for most if not all modulatory neurons in the adult and developing CNS. It may also be assumed that these axonal arborizations, endowed with varicosities that often lack a junctional specialization, are highly plastic, not only in a functional sense but also in their structural configuration. As initially postulated by Descarries and coworkers (1975, 1977; Beaudet and Descarries, 1978), it is likely that such small varicosities, lying free in the neuropil, undergo incessant movements of remodeling and translocation along their parent fibers, resulting in release of their transmitter or modulator in different microlocations of the tissue at different moments in time. Several lines of evidence suggest that many of the peptidergic systems might also share such properties. It is also apparent that these structural determinants apply to both extrinsic and intrinsic neuromodulatory systems, as defined by Katz and Frost (1996). By definition, extrinsic neuromodulation refers to the capacity of a system to cause global changes, affecting many functional circuits simultaneously. The best examples of extrinsic modulatory systems are the DA, NA, 5‐HT, and almost all ACh systems. To varying degrees, these systems pervade most CNS regions and originate from relatively small numbers of neuronal cell bodies, grouped in discrete nuclei. Thus, through their innervation of numerous cortical and subcortical regions, the DA subsystems contribute to various aspects of motor function, neuroendocrine control, motivation, and behavioral learning. The NA, 5‐HT, and ACh systems are even more widespread, innervating most if not all regions of the CNS. They seem to be involved in capacities of a rather global nature and rely on the simultaneous functioning of widely distributed numerous neuronal circuits, such as waking and sleep, arousal, attention, emotional states, learning, memory, and ultimately consciousness. In this perspective, state‐ and context‐dependent responsiveness of the nervous system may be viewed as resulting from the joint activity of multiple, spatially and temporally overlapping extrinsic neuromodulatory systems, as well as that of more specialized, function‐specific systems. Intrinsic neuromodulation arises from neurons entirely contained within a given circuitry (i.e., interneurons). The massive ACh innervation of neostriatum issued from a fraction of its relatively small population of scattered interneurons is a good example of an intrinsic modulatory system. These tonically active neurons appear to be critical elements in the striatal circuitry controlling motor planning, movement, and associative learning (Graybiel et al., 1994; Bennett and Wilson, 1999). Theoretically, intrinsic neuromodulation produces local changes in neuronal computation within a circuit. In view of the largely asynaptic nature of the neostriatal ACh system, its influence is certainly exerted beyond point‐to‐point synaptic connections, and presumably reaches a variety of more‐or‐less distant cellular targets within the neostriatal circuitry. As an intrinsic component, the level of activity of this ACh system would be directly dependent on that of the overall circuitry, the latter being itself modulated by extrinsic inputs (e.g., DA and 5‐HT). It is tempting to speculate that the activity of such a circuitry is reflected by fluctuations in the ambient level of its intrinsic neuromodulators (see Descarries et al., 1997). On the other hand, the ambient levels of extrinsic neuromodulators should more closely depend on the activity of these systems themselves. Much as the progressive unraveling of the numerous modulatory systems, several fairly recent data and concepts pertaining to chemical neurotransmission per se emphasize the previously unsuspected multiplicity of transmission modes in the CNS. The coexistence of transmitter substances within the same neurons (Ho¨kfelt et al., 1980; Merighi, 2002) is now recognized as a common if not universal trait in the CNS, much as the release of transmitter from dendrites, and presumably from cell bodies (Cheramy et al., 1981; He´ry et al., 1982). Spillover beyond the synaptic clefts has been demonstrated for most of the highly if not exclusively synaptic, amino acid transmitters (Isaacson et al., 1993; Kullmann, 2000). There is also ample evidence that multiple metabotropic and/or ionotropic receptors exist for every neurotransmitter/modulator, and receptors for a wide variety of transmitters, and even different receptor subtypes for the same transmitter, are expressed by a given neuron. Moreover, whenever visualized by electron microscopic immunocytochemistry, most neuronal receptors, whether somatodendritic and/or axonal, appear to be located on extra‐ as well as intrasynaptic portions of the plasma membrane (e.g., Riad et al., 2000). Lastly, a growing number of signaling cascades, accounting for short‐, medium‐, and long‐term effects, have been identified for most

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

1

neurotransmitter/modulators. All these mechanisms and properties are obviously needed to support the coordinated activity of a complex nervous system, carrying out multiple functions simultaneously and capable of processing highly diversified information and to react, adapt, and learn from it.

References Ade`r JP, Room P, Postema F, Korf J. 1980. Bilaterally diverging axon collaterals and contralateral projections from rat locus coeruleus neurons, demonstrated by fluorescent retrograde double labeling and norepinephrine metabolism. J Neural Transm 49: 207-218. Albanese A, Minciacchi D. 1983. Organization of the ascending projection from the ventral tegmental area. A multiple fluorescent retrograde tracer study in the rat. J Comp Neurol 216: 406-420. Alvarez FJ, Dewey DE, McMillin P, Fyffe REW. 1999. Distribution of cholinergic contacts on Renshaw cells in the rat spinal cord: A light microscopic study J Physiol 515: 787-797. Ande´n NE, Fuxe K, Hamberger B, Ho¨kfelt T. 1966. A quantitative study of the nigro‐striatal dopamine neuron system in the rat. Acta Physiol Scand 67: 306-312. Antonopoulos J, Dinopoulos A, Dori I, Parnavelas JG. 1997. Distribution and synaptology of dopaminergic fibers in the mature and developing lateral septum of the rat. Brain Res Dev Brain Res 102: 135-141. Antonopoulos J, Dori I, Dinopoulos A, Chiotelli M, Parnavelas JG. 2002. Postnatal development of the dopaminergic system of the striatum in the rat. Neuroscience 110: 245-256. Antonopoulos J, Latsari M, Dori I, Chiotelli M, Parnavelas JG, et al. 2004. Noradrenergic innervation of the developing and mature septal area of the rat. J Comp Neurol 476: 80-90. Aoki C, Venkatesan C, Go CG, Forman R, Kurose H. 1998. Cellular and subcellular sites for noradrenergic action in the monkey dorsolateral prefrontal cortex as revealed by the immunocytochemical localization of noradrenergic receptors and axons. Cereb Cortex 8: 269-277. Armstrong DM, Saper CB, Levey AI, Wainer BH, Terry RD. 1983. Distribution of cholinergic neurons in rat brain: Demonstrated by the immunocytochemical localization of choline acetyltransferase. J Comp Neurol 216: 53-68. Audet MA, Descarries L, Doucet G. 1989. Quantified regional and laminar distribution of the serotonin innervation in the anterior half of adult rat cerebral cortex. J Chem Neuroanat 2: 29-44. Audet MA, Doucet G, Oleskevich S, Descarries L. 1988. Quantified regional and laminar distribution of the noradrenaline innervation in the anterior half of the adult rat cerebral cortex. J Comp Neurol 274: 307-318.

Aznavour N, Mechawar N, Descarries L. 2002. Comparative analysis of the cholinergic innervation in the dorsal hippocampus of adult mouse and rat. A quantitative immunocytochemical study. Hippocampus 12: 406-417. Aznavour N, Mechawar N, Watkins KC, Descarries L. 2003. Fine structural features of the acetylcholine innervation in the developing neostriatum of rat. J Comp Neurol 460: 280-291. Aznavour N, Watkins KC, Descarries L. 2005. Postnatal development of the cholinergic innervation in the dorsal hippocampus of rat: A quantitative light and electron microscopic immunocytochemical description. J Comp Neurol 486: 61-75. Barber RP, Phelps PE, Houser CR, Crawford GD, Salvaterra PM, et al. 1984. The morphology and distribution of neurons containing choline acetyltransferase in the adult rat spinal cord: An immunocytochemical study. J Comp Neurol 229: 329-346. Beaudet A, Descarries L. 1978. The monoamine innervation of rat cerebral cortex: Synaptic and nonsynaptic relationships. Neuroscience 3: 851-860. Bennett BD, Wilson CJ. 1999. Spontaneous activity of neostriatal cholinergic interneurons in vitro. J Neurosci 19: 5586-5596. Bjo¨rklund A, Lindvall O. 1984. Dopamine‐containing systems in the CNS. Handbook of Chemical Neuroanatomy, Vol. 2. Classical Transmitters in the CNS. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 55-122. Bjo¨rklund A, Skagerberg G. 1979. Evidence for a major spinal cord projection from the diencephalic A11 dopamine cell group in the rat using transmitter‐specific fluorescent retrograde tracing. Brain Res 177: 170-175. Bowker RM, Westlund KN, Sullivan MC, Coulter JD. 1982. Organization of descending serotonergic projections to the spinal cord. Prog Brain Res 57: 239-298. Cadete‐Leite A, Pereira PA, Madeira MD, Paula‐Barbosa MM. 2003. Nerve growth factor prevents cell death and induces hypertrophy of basal forebrain cholinergic neurons in rats withdrawn from prolonged ethanol intake. Neuroscience 119: 1055-1069. Che´dotal A, Umbriaco D, Descarries L, Hartman BK, Hamel E. 1994. Light and electron microscopic immunocytochemical analysis of the neurovascular relationships of choline acetyltransferase and vasoactive intestinal polypeptide nerve terminals in the rat cerebral cortex. J Comp Neurol 343: 57-71.

15

16

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

Cheney DL, Le Fevre HF, Racagni G. 1975. Choline acetyltransferase activity and mass fragmentographic measurement of acetylcholine in specific nuclei and tracts of rat brain. Neuropharmacology 14: 801-809. Cheramy A, Leviel V, Glowinski J. 1981. Dendritic release of dopamine in the substantia nigra. Nature 289: 537-542. Cohen Z, Ehret M, Maıˆtre M, Hamel E. 1995. Ultrastructural analysis of tryptophane hydroxylase immunoreactive nerve terminals in the rat cerebral cortex and hippocampus: Their associations with local blood vessels. Neuroscience 66: 555-569. Cohen Z, Molinatti G, Hamel E. 1997. Astroglial and vascular interactions of noradrenaline terminals in the rat cerebral cortex. J Cereb Blood Flow Metab 17: 894-904. Contant C, Umbriaco D, Garcia S, Watkins KC, Descarries LD. 1996. Ultrastructural characterization of the acetylcholine innervation in adult rat neostriatum. Neuroscience 71: 937-947. Coyle JT, Yamamura HI. 1976. Neurochemical aspects of the ontogenesis of cholinergic neurons in the rat brain. Brain Res 118: 429-440. Cozzari C, Howard J, Hartman B. 1990. Analysis of epitopes on choline acetyltransferase (ChAT) using monoclonal antibodies (Mabs). Soc Neurosci Abstr 16: 200. Dahlstro¨m A, Fuxe K. 1964. Evidence for the existence of monoamine‐containing neurons in the central nervous system I. Demonstration of monoamines in the cell bodies of brain stem neurons. Acta Physiol Scand 62 (Suppl. 232): 1-55. DeFelipe J, Jones EG. 1988. A light and electron microscopic study of serotonin‐immunoreactive fibers and terminals in the monkey sensory‐motor cortex. Exp Brain Res 71: 171-182. DeFelipe J, Hendry SHC, Hashikawa T, Jones EG. 1991. Synaptic relationships of serotonin‐immunoreactive terminal baskets of GABA neurons in the cat auditory cortex. Cereb Cortex 1: 117-133. Descarries L, Beaudet A. 1983. The use of radioautography for investigating transmitter‐specific neurons. Handbook of Chemical Neuroanatomy, Vol. 1, Methods in Chemical Neuroanatomy. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 286-364. Descarries L, Saucier G. 1972. Disappearance of the locus coeruleus in the rat after intraventricular 6‐hydroxydopamine. Brain Res 37: 310-316. Descarries L, Beaudet A, Watkins KC. 1975. Serotonin nerve terminals in adult rat neocortex. Brain Res 100: 563-588. Descarries L, Doucet G, Lemay B, Se´gue´la P, Watkins KC. 1988. Structural basis of cortical monoamine function. Neurotransmitters and Cortical Function. Avoli M, Reader TA, Dykes RW, Gloor P, editors. New York: Plenum Publishing, pp. 321-332.

Descarries L, Gisiger V, Steriade M. 1997. Diffuse transmission by acetylcholine in the CNS. Prog Neurobiol 53: 603-625. Descarries L, Lemay B, Doucet G, Berger B. 1987. Regional and laminar density of the dopamine innervation in adult rat cerebral cortex. Neuroscience 21: 807-824. Descarries L, Mechawar N, Aznavour N. 2002. Structural determinants of the roles of acetylcholine in cerebral cortex. Progress in Brain Research, Vol. 145, Acetylcholine in the Cerebral Cortex. Descarries L, Krnjevic K, Steriade M, editors. Amsterdam: Elsevier; pp. 45-58. Descarries L, Watkins KC, Garcia S, Beaudet A. 1982. The serotonin neurons in nucleus raphe dorsalis of adult rat. A light and electron microscope radioautographic study. J Comp Neurol 207: 239-254. Descarries L, Watkins KC, Garcia S, Bosler O, Doucet G. 1996. Dual character, asynaptic and synaptic, of the dopamine innervation in adult rat neostriatum: A quantitative autoradiographic and immunocytochemical analysis. J Comp Neurol 375: 167-186. Descarries L, Watkins KC, Lapierre Y. 1977. Noradrenergic axon terminals in the cerebral cortex of rat. III. Topometric ultrastructural analysis. Brain Res 133: 197-222. Dinopoulos A, Dori I. 1995. The development of the serotonergic fiber network of the lateral ventricles of the rat brain: A light and electron microscopic immunocytochemical analysis. Exp Neurol 133: 73-84. Dinopoulos A, Dori I, Parnavelas JG. 1993. Serotonergic innervation of the mature and developing lateral septum of the rat: A light and electron microscopic immunocytochemical analysis. Neuroscience 55: 209-222. Dinopoulos A, Dori I, Parnavelas JG. 1995. Serotonergic innervation of the lateral geniculate nucleus of the rat during postnatal development: A light and electron microscopic immunocytochemical analysis. J Comp Neurol 363: 532-544. Dinopoulos A, Dori I, Parnavelas JG. 1997. The serotonin innervation of the basal forebrain shows a transient phase during development. Brain Res Dev Brain Res 99: 38-52. Dori I, Dinopoulos A, Blue ME, Parnavelas JG. 1996. Regional differences in the ontogeny of the serotonergic projection to the cerebral cortex. Exp Neurol 138: 1-14. Dori IE, Dinopoulos A, Parnavelas JG. 1998. The development of the synaptic organization of the serotonergic system differs in brain areas with different functions. Exp Neurol 154: 113-125. Doucet G, Descarries L, Garcia S. 1986. Quantification of the dopamine innervation in adult rat neostriatum. Neuroscience 19: 427-445. Eckenstein F, Baughman RW. 1984. Two types of cholinergic innervation in cortex, one co‐localized with vasoactive intestinal polypeptide. Nature 309: 153-155.

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS Eckenstein F, Thoenen H. 1983. Cholinergic neurons in the rat cerebral cortex demonstrated by immunohistochemical localization of choline acetyltransferase. Neurosci Lett 36: 211-215. Erickson SL, Sesack SR, Lewis DA. 2000. The dopamine innervation of monkey entorhinal cortex: Postsynaptic targets of tyrosine hydroxylase terminals. Synapse 36: 47-56. Fallon JH, Loughlin SE. 1982. Monoamine innervation of the forebrain: Collateralization. Brain Res Bull 9: 295-307. Fournier GN, Semba K, Rasmusson DD. 2004. Modality‐and region‐specific acetylcholine release in the rat neocortex. Neuroscience 126: 257-262. Fritschy JM, Grzanna R. 1990. Demonstration of two separate descending noradrenergic pathways to the rat spinal cord: Evidence for an intragriseal trajectory of locus coeruleus axons in the superficial layers of the dorsal horn. J Comp Neurol 291: 553-582. Gauthier J, Parent M, Le´vesque M, Parent A. 1999. The axonal arborization of single nigrostriatal neurons in rats. Brain Res 834: 228-232. Graybiel AM, Aosaki T, Flaherty AW, Kimura M. 1994. The basal ganglia and adaptive motor control. Science 265: 1826-1831. Gritti I, Mainville L, Jones BE. 1993. Codistribution of GABA‐ with acetylcholine‐synthesizing neurons in the basal forebrain of the rat. J Comp Neurol 329: 438-457. Guyenet PG, Crane JK. 1981. Non‐dopaminergic nigro‐striatal pathway. Brain Res 213: 291-305. Hedreen JC, Chalmers JP. 1972. Neuronal degeneration in rat brain induced by 6‐hydroxydopamine: A histological and biochemical study. Brain Res 47: 1-36. He´ry F, Faudon M, Ternaux JP. 1982. In vivo release of serotonin in two raphe nuclei (raphe dorsalis and magnus) of the cat. Brain Res Bull 8: 123-129. Ho¨kfelt T, Fuxe K, Goldtein M, Johansson O. 1974. Evidence for the existence of adrenaline neurons in the rat brain. Brain Res 66: 235-251. Ho¨kfelt T, Johansson O, Goldstein M. 1984a. Central catecholamine neurons as revealed by immunohistochemistry with special reference to adrenaline neurons. Handbook of Chemical Neuroanatomy, Vol. 2, Classical Transmitters in the CNS. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 157-276. Ho¨kfelt T, Ma˚rtensson, Bjo¨rklund A, Kleinau S, Goldstein M. 1984b. Distributional maps of tyrosine‐hydroxylase‐immunoreactive neurons in the rat brain. Handbook of Chemical Neuroanatomy, Vol. 2, Classical Transmitters in the CNS. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 277-379. Ho¨kfelt T, Lundberg JM, Schultzberg M, Johansson O, Ljungdahl A, et al. 1980. Coexistence of peptides and

1

putative transmitters in neurons. Adv Biochem Psychopharmacol 22: 1-23. Ho¨kfelt T, Phillipson O, Goldtein M. 1979. Evidence for a dopaminergic pathway in the rat descending from the A11 cell group to the spinal cord. Acta Physiol Scand 107: 393-395. Hoover DB, Muth EA, Jacobowitz DM. 1978. A mapping of the distribution of acetylcholine, choline acetyltransferase and acetylcholinesterase in discrete areas of rat brain. Brain Res 153: 295-306. Huang A, Noga BR, Carr PA, Fedirchuk B, Jordan LM. 2000. Spinal cholinergic neurons activated during locomotion: Localization and electrophysiological characterization. J Neurophysiol 83: 3537-3547. Inagaki N, Yamatodani A, o‐Yamamoto M, Tohyama M, Watanabe T, et al. 1988. Organization of histaminergic fibers in the rat brain. J Comp Neurol 273: 283-300. Isaacson JS, Solis JM, Nicoll RA. 1993. Local and diffuse synaptic actions of GABA in the hippocampus. Neuron 10: 165-175. Johnston MV, McKinney M, Coyle JT. 1981. Neocortical cholinergic innervation: A description of extrinsic and intrinsic components in the rat. Exp Brain Res 43: 159-172. Kalsbeek A, Voorn P, Buijs RM, Pool CW, Uylings HB. 1988. Development of the dopaminergic innervation in the prefrontal cortex of the rat. J Comp Neurol 269: 58-72. Katz PS, Frost WN. 1996. Intrinsic neuromodulation: Altering neuronal circuits from within. Trends Neurosci 19: 54-61. Kullmann DM. 2000. Spillover and synaptic cross talk mediated by glutamate and GABA in the mammalian brain. Prog Brain Res 125: 339-351. Latsari M, Antonopoulos J, Dori I, Chiotelli M, Dinopoulos A. 2004. Postnatal development of the noradrenergic system in the dorsal lateral geniculate nucleus of the rat. Brain Res Dev Brain Res 149: 79-83. Latsari M, Dori I, Antonopoulos J, Chiotelli M, Dinopoulos A. 2002. Noradrenergic innervation of the developing and mature visual and motor cortex of the rat brain: A light and electron microscopic immunocytochemical analysis. J Comp Neurol 445: 145-158. Lebrand C, Cases O, Adelbrecht C, Doye A, Alvarez C, et al. 1996. Transient uptake and storage of serotonin in developing thalamic neurons. Neuron 17: 823-835. Levey AI, Wainer BH, Rye DB, Mufson EJ, Mesulam MM. 1984. Choline acetyltransferase‐immunoreactive neurons intrinsic to rodent cortex and distinction from acetylcholinesterase neurons. Neuroscience 13: 341-353. Levitt P, Rakic P, Goldman‐Rakic P. 1984. Comparative assessment of monoamine afferents in mammalian cerebral cortex. Monoamine Innervation of Cerebral Cortex. Descarries L, Reader TA, Jasper HH, editors. New York: Alan R. Liss; pp. 41-59.

17

18

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

Lindvall O, Bjo¨rklund A. 1974a. The glyoxylic acid fluorescence histochemical method: A detailed account of the methodology for the visualization of central catecholamine neurons. Histochemistry 39: 97-127. Lindvall O, Bjo¨rklund A. 1974b. The organization of the ascending catecholamine neuron systems in the rat brain as revealed by the glyoxylic acid fluorescence method. Acta Physiol Scand (Suppl. 412): 1‐48. Lindvall O, Bjo¨rklund A. 1978. Organization of catecholamine neurons in rat central nervous system. Handbook of Psychopharmacology, Vol. 9. Iversen L, Iversen S, Snyder SN, editors. New York: Plenum Press; pp. 139-231. Loughlin SE, Fallon JH. 1984. Substantia nigra and ventral tegmental area projections to cortex: Topography and collateralization. Neuroscience 11: 425-435. Loughlin SE, Foote SL, Fallon SH. 1982. Locus coeruleus projections to cortex: Topography, morphology and collateralization. Brain Res Bull 9: 287-294. Lysakowski A, Wainer BH, Bruce G, Hersh LB. 1989. An atlas of the regional and laminar distribution of choline acetyltransferase immunoreactivity in rat cerebral cortex. Neuroscience 28: 291-336. Markham JA, Vaughn JE. 1990. Ultrastructural analysis of choline acetyltransferase‐immunoreactive sympathetic preganglionic neurons and their dendritic bundles in rat thoracic spinal cord. Synapse 5: 299-312. Marlier L, Sandillon F, Poulat P, Rajaofetra, Geffard M, Privat A. 1991. Serotonergic innervation of the dorsal horn of rat spinal cord: Light and electron microscopic immunocytochemical study. J Neurocytol 20: 310-322. Mechawar N, Descarries L. 2001. The cholinergic innervation develops early and rapidly in the rat cerebral cortex: A quantitative immunocytochemical study. Neuroscience 108: 555-567. Mechawar N, Cozzari C, Descarries L. 2000. Cholinergic innervation of adult rat cerebral cortex: A quantitative immunocytochemical description. J Comp Neurol 428: 305-318. Mechawar N, Watkins KC, Descarries L. 2002. Ultrastructural features of the acetylcholine innervation in the developing parietal cortex of rat. Neuroscience 111: 83-94. Merighi A. 2002. Costorage and coexistence of neuropeptides in the mammalian CNS. Prog Neurobiol 66: 161-190. Mesulam MM. 1988. Central cholinergic pathways: Neuroanatomy and some behavioral implications. Neurotransmitters and Cortical Function: From Molecules to Mind. Avoli M, Reader TA, Gloor D, Dykes RW, editors. New York: Plenum Press; pp. 237-260. Mesulam MM, Mufson EJ, Wainer BH, Levey AI. 1983. Central cholinergic pathways in the rat: An overview based on an alternative nomenclature (Ch1–Ch6). Neuroscience 10: 1185-1201.

Milner TA, Morrison SF, Abate C, Reis DJ. 1988. Phenylethanolamine N‐methyltransferase‐containing terminals synapse directly on sympathetic preganglionic neurons in the rat. Brain Res 17: 205-222. Moore RY, Card JP. 1984. Noradrenaline‐containing neuron systems. Handbook of Chemical Neuroanatomy, Vol. 2, Classical Transmitters in the CNS. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 123-156. Morrison JH, Foote SL, Bloom FE. 1984. Regional, laminar, developmental, functional characteristics of noradrenaline and serotonin innervation patterns in monkey cortex. Monoamine Innervation of Cerebral Cortex. Descarries L, Reader TA, Jasper HH, editors. New York: Alan R. Liss; pp. 61-75. Morrison JH, Molliver ME, Grzanna R, Coyle JT. 1981. The intracortical trajectory of the coeruleo‐cortical projection in the rat: A tangentially organized cortical afferent. Neuroscience 6: 139-18. Mrini A, Soucy JP, Lafaille F, Lemoine P, Descarries L. 1995. Quantification of the serotonin hyperinnervation in adult rat neostriatum after neonatal 6‐hydroxydopamine lesion of nigral dopamine neurons. Brain Res 669: 303-308. Mrzljak L, Pappy M, Leranth C, Goldman-Rakic PS. 1995. Cholinergic synaptic circuitry in the macaque prefrontal cortex. J Comp Neurol 357: 603-617. Nagai T, Satoh K, Imamoto K, Maeda T. 1981. Divergent projections of catecholamine neurons of the locus coeruleus as revealed by fluorescent retrograde double labeling techniques. Neurosci Lett 23: 117-123. Nagy JI, Yamamoto T, Jordan LM. 1993. Evidence for the cholinergic nature of c‐terminals associated with subsurface cisterns in a‐motoneurons of rat. Synapse 15: 17-32. Oleskevich S, Descarries L. 1990. Quantified distribution of the serotonin innervation in adult rat hippocampus. Neuroscience 34: 19-33. Oleskevich S, Descarries L, Lacaille JC. 1989. Quantified distribution of the noradrenaline innervation in the hippocampus of adult rat. J Neurosci 9: 3803-3815. Oleskevich S, Descarries L, Watkins KC, Se´gue´la P, Daszuta A. 1991. Ultrastructural features of the serotonin innervation in adult rat hippocampus. An immunocytochemical description in single and serial thin sections. Neuroscience 42: 777-791. Oorschot DE. 1996. Total number of neurons in the neostriatal, pallidal, subthalamic, substantia nigral nuclei of the rat basal ganglia: A stereological study using the Cavalieri and optical dissector methods. J Comp Neurol 366: 580-599. Panula P, Pirvola U, Auvinen S, Airaksinen MS. 1989. Histamine immunoreactive nerve fibers in the rat brain. Neuroscience 28: 585-610. Papadopoulos GC, Parnavelas JG, Buijs RM. 1989. Light and electron microscopic immunocytochemical analysis of the

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS dopamine innervation of the rat visual cortex. J Neurocytol 18: 303-310. Phelps PE, Barber RP, Houser CR, Crawford GD, Salvaterra PM, et al. 1984. Postnatal development of neurons containing choline acetyltransferase in rat spinal cord: An immunocytochemical study. J Comp Neurol 229: 347-361. Phelps PE, Houser CR, Vaughn JE. 1985. Immunocytochemical localization of choline acetyltransferase within the rat neostriatum: A correlated light and electron microscopic study of cholinergic neurons and synapses. J Comp Neurol 238: 286-307. Poulat P, Marlier L, Rajaofetra N, Privat A. 1992. 5‐Hydroxytryptamine, substance P and thyrotropin‐releasing hormone synapses in the intermediolateral cell column of the rat thoracic spinal cord. Neurosci Lett 136: 19-22. Prensa L, Parent A. 2001. The nigrostriatal pathway in the rat: A single‐axon study of the relationship between dorsal and ventral tier nigral neurons and the striosome/matrix striatal compartments. J Neurosci 21: 7247-7260. Rajaofetra N, Ridet JL, Poulat P, Marlier L, Sandillon F, et al. 1992. Immunocytochemical mapping of noradrenergic projections to the rat spinal cord with an antiserum against noradrenaline. J Neurocytol 21: 481-494. Rea MA, Simon JR. 1981. Regional distribution of cholinergic parameters within the rat striatum. Brain Res 219: 317-326. Riad M, Garcia S, Watkins KC, Jodoin N, Doucet E´, et al. 2000. Somatodendritic localization of 5‐HT1A and preterminal axonal localization of 5‐HT1B serotonin receptors in adult rat brain. J Comp Neurol 417: 181-194. Ridet JL, Rajaofetra N, Teilhac JR, Geffard M, Privat A. 1993. Evidence for non‐synaptic serotonergic and noradrenergic innervation of the rat dorsal horn and possible involvement of neuron‐glia interactions. Neuroscience 52: 143-157. Ridet JL, Sandillon F, Rajaofetra N, Geffard M, Privat A. 1992. Spinal dopaminergic system of the rat: Light and electron microscopic study using an antiserum against dopamine, with particular emphasis on synaptic incidence. Brain Res 598: 233-241. Room P, Postema F, Korf J. 1981. Divergent axon collaterals of rat locus coeruleus. Brain Res 221: 219-230. Rye DB, Wainer BH, Mesulam MM, Mufson EJ, Saper CB. 1984. Cortical projections arising from the basal forebrain: A study of cholinergic and noncholinergic components employing combined retrograde tracing and immunohistochemical localization of choline acetyltransferase. Neuroscience 13: 627-643. Saper CB. 1984. Organization of cerebral cortical afferent systems in the rat. II. Magnocellular basal nucleus. J Comp Neurol 222: 313-342. Scha¨fer MK, Eiden LE, Weihe E. 1998. Cholinergic neurons and terminal fields revealed by immunohistochemistry for

1

the vesicular acetylcholine transporter. I. Central nervous system. Neuroscience 84: 331-359. Se´gue´la P, Watkins KC, Descarries, L. 1988. Ultrastructural features of dopamine axon terminals in the anteromedial and suprarhinal cortex of adult rat. Brain Res 442: 11-22. Se´gue´la P, Watkins KC, Descarries L. 1989. Ultrastructural relationships of serotonin axon terminals in the cerebral cortex of the adult rat. J Comp Neurol 289: 129-142. Se´gue´la P, Watkins KC, Geffard M, Descarries L. 1990. Noradrenaline axon terminals in adult rat neocortex: An immunocytochemical analysis in serial thin sections. Neuroscience 35: 249-264. Seiger A, Olson L. 1973. Late prenatal ontogeny of central monoamine neurons in the rat: Fluorescence histochemical observations. Z Anat Entwicklungsgesch 140: 281-318. Semba K. 1992. Development of central cholinergic neurons. Handbook of Chemical Neuroanatomy, Vol. 10, Ontogeny of Transmitters and Peptides in the CNS. Bjo¨rklund A, Ho¨kfelt T, Tohyama M, editors. Amsterdam: Elsevier; pp. 33-62. Semba K. 2004. Phylogenetic and ontogenetic aspects of the basal forebrain cholinergic neurons and their innervation of the cerebral cortex. Descarries L, Krnjevic K, Steriade M, editors. Progress in Brain Research, Vol. 145, Acetylcholine in the Cerebral Cortex. Amsterdam: Elsevier; pp. 3-43. Sherriff FE, Henderson Z. 1994. A cholinergic propriospinal innervation of the rat spinal cord. Brain Res 634: 150-154. Sherriff FE, Henderson ZH, Morrison JFB. 1991. Further evidence for the absence of a descending cholinergic projection from the brainstem to the spinal cord in the rat. Neurosci Lett 128: 52-56. Shirouzu M, Anraku T, Iwashi TA, Yoshida M. 1990. A new dopaminergic terminal plexus in the ventral horn of the spinal cord. Immunohistochemical studies at the light and electron microscopic levels. Experientia 46: 201-204. Skagerberg G, Bjo¨rklund A. 1985. Topographic principles in the spinal projections of serotonergic and non‐ serotonergic brainstem neurons in the rat. Neuroscience 15: 445-480. Skagerberg G, Bjo¨rklund A, Lindvall O, Schmidt RH. 1982. Origin and termination of the diencephalo‐spinal dopamine system in the rat. Brain Res Bull 9: 237-244. Smiley JF, Goldman‐Rakic PS. 1993. Heterogeneous targets of dopamine synapses in monkey prefrontal cortex demonstrated by serial section electron microscopy: A laminar analysis using the silver‐enhanced diaminobenzidine sulfide (SEDS) immunolabeling technique. Cereb Cortex 3: 223-238. Smiley JF, Goldman‐Rakic PS. 1996. Serotonergic axons in monkey prefrontal cerebral cortex synapse predominantly on interneurons as demonstrated by serial section electron microscopy. J Comp Neurol 367: 431-443.

19

20

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

Smiley JF, Morrell F, Mesulam MM. 1997. Cholinergic synapses in human cerebral cortex: An ultrastructural study in serial sections. Exp Neurol 144: 361-368. Smiley JF, Williams SM, Szigeti K, Goldman‐Rakic PS. 1992. Light and electron microscopic characterization of dopamine‐immunoreactive axons in human cerebral cortex. J Comp Neurol 32: 325-335. Steinbusch HWM. 1981. Distribution of serotonin in the central nervous system of the rat—cell bodies and terminals. Neuroscience 6: 557-618. Steindler DA. 1981. Locus coeruleus neurons have axon collaterals that branch to the forebrain and cerebellum. Brain Res 223: 367-373. Swanson LW. 1976. The locus coeruleus: A cytoarchitectonic Golgi and immunohistochemical study in the albino rat. Brain Res 110: 39-56. Swanson LW. 1982. The projections of the ventral tegmental area and adjacent regions: A combined fluorescent retrograde tracer and immunofluorescence study in the rat. Brain Res Bull 9: 321-353. Swanson LW, Hartman BK. 1975. The central adrenergic system: An immunofluorescence study of the location of cell bodies and their efferent connections in the rat utilizing dopamine‐b‐hydroxylase as a marker. J Comp Neurol 163: 467-506. Swanson LW, Sawchenko PE, Be´rod A, Hartman BK, Helle KB, et al. 1981. An immunohistochemical study of the organization of catecholaminergic cells and terminal fields in the paraventricular and supraoptic nuclei of the hypothalamus. J Comp Neurol 196: 271-285. Takagi H, Morishima Y, Matsuyama T, Hayashi H, Watanabe T, et al. 1986. Histaminergic axons in the neostriatum and cerebral cortex of the rat: A correlated light and electron microscopic immunocytochemical study using histidine decarboxylase as a marker. Brain Res 364: 155-168. Turrini P, Casu MA, Wong TP, De Koninck Y, Ribeiro‐Da‐ Silva A, et al. 2001. Cholinergic nerve terminals establish classical synapses in the rat cerebral cortex: Synaptic pattern and age‐related atrophy. Neuroscience 105: 277-285. Umbriaco D, Garcia S, Beaulieu C, Descarries L. 1995. Relational features of acetylcholine, noradrenaline, serotonin and GABA axon terminals in the stratum radiatum of adult rat hippocampus (CA1). Hippocampus 5: 605-620. Umbriaco D, Watkins KC, Descarries L, Cozzari C, Hartman BK. 1994. Ultrastructural and morphometric features of the acetylcholine innervation in adult rat parietal cortex: An electron microscopic study in serial sections. J Comp Neurol 348: 351-373.

Ungerstedt U. 1971. Stereotaxic mapping of the monoamine pathways in the rat brain. Acta Physiol Scand 367: (Suppl) 1-48. Vaucher E, Hamel E. 1995. Cholinergic basal forebrain neurons project to cortical microvessels in the rat: Electron microscopic study with anterogradely transported Phaseolus vulgaris leucoagglutinin and choline acetyltransferase immunocytochemistry. J Neurosci 15: 7427-7441. Verney C, Berger B, Adrien J, Vigny A, Gay M. 1982. Development of the dopaminergic innervation of the rat cerebral cortex. A light microscopic immunocytochemical study using anti-tyrosine hydroxylase antibodies. Brain Res 281: 41-52. Volpicelli LA, Levey AI. 2004. Muscarinic acetylcholine receptor subtypes in cerebral cortex and hippocampus. Prog Brain Res 145: 59-66. Watanabe T, Taguchi Y, Shiosaka S, Tanaka J, Kubota H, et al. 1984. Distribution of histaminergic neuron system in the central nervous system of rats; a fluorescent immunocytochemical analysis with histidine decarboxylase as a marker. Brain Res 295: 13-25. Westlund KN, Bowker RM, Ziegler MG, Coulter JD. 1981. Origins of spinal noradrenergic pathways demonstrated by retrograde transport of antibody to dopamine‐b‐hydroxylase. Neurosci Lett 25: 243-249. Westlund KN, Bowker RM, Ziegler MG, Coulter JD. 1982. Descending noradrenergic projections and their spinal terminations. Prog Brain Res 57: 219-238. Westlund KN, Bowker RM, Ziegler MG, Coulter JD. 1983. Noradrenergic projections to the spinal cord of the rat. Brain Res 263: 15-31. Woolf NJ. 1991. Cholinergic systems in mammalian brain and spinal cord. Prog Neurobiol 37: 475-524. Woolf NJ, Butcher LL. 1981. Cholinergic neurons in the caudate‐putamen complex proper are intrinsically organized: A combined Evans blue and acetylcholinesterase analysis. Brain Res Bull 7: 487-507. Woolf NJ, Butcher LL. 1986. Cholinergic systems in the rat brain. III. Projections from the pontomesencephalic tegmentum to the thalamus, tectum, basal ganglia, basal forebrain. Brain Res Bull 16: 603-637. Zaborszky L, Duque A. 2000. Local synaptic connections of basal forebrain neurons. Behav Brain Res 115: 143-158. Zoli M, Jansson A, Sykova´ E, Agnati LF, Fuxe K. 1999. Volume transmission in the CNS and its relevance for neuropsychopharmacology. Trends Pharmacol Sci 20: 142-150.

2

Brain Neurons Partly Expressing Monoaminergic Phenotype: Distribution, Development, and Functional Significance in Norm and Pathology

M. V. Ugrumov

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23

2 2.1 2.1.1 2.1.2 2.1.3 2.2 2.3

Brain Neurons Partly Expressing the MA‐ergic Phenotype in Adult Mammals . . . . . . . . . . . . . . . Neurons Expressing Individual Enzymes of MA Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hypothalamus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Striatum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Other Brain Regions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bienzymatic TH- and AADC-Expressing Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Non-MA-ergic Neurons Expressing the MA Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

26 26 26 29 31 32 33

3 3.1 3.1.1 3.1.2 3.2

Brain Neurons Partly Expressing MA‐ergic Phenotype in Mammals in Ontogenesis . . . . . . . . . Neurons Expressing Individual Enzymes of MA Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hypothalamus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Extrahypothalamic Regions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Non-MA-ergic Neurons Expressing the MA Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

35 35 35 39 42

4

Functional Properties and Functional Significance of the Neurons Partly Expressing the MA‐ergic Phenotype . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monoenzymatic Neurons Expressing TH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monoenzymatic Neurons Expressing AADC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ensembles of Monoenzymatic Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Non‐MA‐ergic Neurons Expressing the MA Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

44 44 47 48 51

4.1 4.2 4.3 4.4 5

Tuberoinfundibular Neurons Partly Expressing DA‐ergic Phenotype in Hyperprolactinemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52

6 6.1 6.2 6.3 6.4

Striatal Neurons Partly Expressing DA‐ergic Phenotype in Parkinson’s Disease . . . . . . . . . . . . . . Monoenzymatic TH‐Expressing Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monoenzymatic AADC‐Expressing Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bienzymatic TH‐ and AADC‐Expressing Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Origin, Functional Properties, and Functional Significance of Striatal Neurons Partly or Completely Expressing the DA‐ergic Phenotype . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

#

2008 Springer ScienceþBusiness Media, LLC.

53 54 57 59 59

22

2 7 7.1 7.2 7.3

Brain neurons partly expressing monoaminergic phenotype

Regulation of the Partial Expression of MA‐ergic Phenotype by the Brain Neurons in Norm and Pathology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of the Partial Expression of MA‐ergic Phenotype by Neural Afferents . . . . . . . . . . . . . . Paracrine Regulation of the Partial Expression of MA‐ergic Phenotype by Diffusive Factors . . Hormonal Regulation of the Partial Expression of MA‐ergic Phenotype . . . . . . . . . . . . . . . . . . . . . . . .

62 62 64 65

Brain neurons partly expressing monoaminergic phenotype

2

Abstract: In addition to the monoaminergic (MA‐ergic) neurons possessing the whole set of enzymes of monoamine (MA) synthesis from the precursor amino acid and the MA membrane transporter, the neurons partly expressing the MA‐ergic phenotype have been first discovered almost twenty years ago. Most of the neurons expressing individual enzymes of MA synthesis lack the MA transporter. These so‐ called monoenzymatic neurons are widely distributed throughout the brain in adult mammals being even more numerous than MA‐ergic neurons. Individual enzymes of MA synthesis are expressed continuously or transiently over certain periods of ontogenesis and in adulthood under functional insufficiency of the MA‐ergic neurons, e.g., under their chronic stimulation or in certain neurodegenerative diseases. The earlier data suggest an important functional role of monoenzymatic neurons. Most monoenzymatic neurons possess enzymes of dopamine (DA) synthesis, tyrosine hydroxylase (TH), or aromatic L‐amino acid decarboxylase (AADC). TH and AADC are enzymatically active in a substantial number of monoenzymatic neurons being capable to convert L‐tyrosine to L‐3,4‐dihydroxyphenylalanine (L‐DOPA) and L‐DOPA to DA or serotonin, respectively. L‐DOPA produced in monoenzymatic TH‐neurons is supposed to play a role of a neurotransmitter or a neuromodulator providing its action on the target neurons via catecholamine receptors. Moreover, L‐DOPA released from the monoenzymatic TH‐neurons is captured by monoenzymatic AADC‐neurons or dopaminergic (DA‐ergic) and serotoninergic neurons for DA synthesis (Kannari et al., 2006). Such cooperative synthesis of MAs is considered as a compensatory reaction under the failure of MA‐ergic neurons, e.g., in neurodegenerative diseases like hyperprolactinemia and Parkinson’s disease which are developed primarily because of the degeneration of DA‐ergic neurons of the tuberoinfundibular system and the nigrostriatal system, respectively. Noteworthy, the neurotoxin‐induced increased level of prolactin returns with time to the normal level due to stimulation of DA synthesis by the neurons of the tuberoinfundibular system, most probably because of the turning on cooperative synthesis of DA by monoenzymatic neurons. The same compensatory mechanism is supposed to be used under the failure of the nigrostriatal DA‐ergic system that is manifested by the increased number of monoenzymatic neurons in the striatum of animals with neurotoxin‐induced parkinsonism and in humans with Parkinson’s disease. Expression of the enzymes of MA synthesis in non‐MA‐ergic neurons is controlled by intercellular signals such as classical neurotransmitters (catecholamines), neurotrophic factors (brain‐derived neurotrophic factor, glia‐derived neurotrophic factor), and perhaps hormones (prolactin, estrogens, progesterone). Thus, a substantial number of the brain neurons express partly the MA‐ergic phenotype, mostly individual complementary enzymes of MA synthesis, serving to produce MAs in cooperation that is considered as a compensatory reaction under the failure of MA‐ergic neurons. List of Abbreviations: AADC, aromatic L‐amino acid decarboxylase; AN, arcuate nucleus; DA, dopamine; DOPA, 3,4‐dihydroxyphenylalanine; GABA, g‐aminobutyric acid; GTP, guanosine triphosphate; MA(s), monoamine(s); MPTP, 1‐methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine; 6‐OHDA, 6‐hydroxydopamine; SCN, suprachiasmatic nucleus; TH, tyrosine hydroxylase; VMAT2, vesicular monoamine transporter 2

1

Introduction

Mechanisms of the regulation of the brain development, plasticity, and integration via chemical neurotransmission are the crucial issue of Neuroscience. The neural and neuroendocrine regulations of most important functions are under the control of dozens or even hundreds of neurotransmitters, neuromodulators, and neurohormones. Monoamines (MAs), catecholamines (dopamine (DA), noradrenaline, and adrenaline), and serotonin are among the most important classical neurotransmitters which are widely distributed all over the brain in ontogenesis and adulthood (Squire et al., 2003). The most frequent MAs, DA and serotonin, are produced enzymatically from L‐tyrosine and L‐tryptophan (the precursor amino acids) in the cytosol of both cell bodies and processes (> Figures 2‐1 and > 2‐2). L‐tyrosine is converted first to L‐3,4‐dihydroxyphenylalanine (L‐DOPA) by tyrosine hydroxylase (TH), the rate‐limiting enzyme of catecholamine synthesis, and then to DA by aromatic L‐amino acid decarboxylase (AADC). In turn, L‐tryptophan is transformed first to 5‐hydroxytryptophan with tryptophan hydroxylase, the rate limiting enzyme of serotonin synthesis, and then to serotonin by AADC

23

24

2

Brain neurons partly expressing monoaminergic phenotype

. Figure 2‐1 Schematic representation of the functioning of dopaminergic and serotoninergic neurons. A, precursor amino acid, L‐tyrosine or L‐tryptophan; D, aromatic L‐amino acid decarboxylase; M, monoamine, dopamine or serotonin; T, tyrosine hydroxylase or tryptophan hydroxylase; TC, target cell; X, intermediate synthetic product, L‐3, 4‐dihydroxyphenylalanine or 5‐hydroxytryptophan, , dopamine or serotonin transporter; , receptor to dopamine or serotonin; , secretory granule at high magnification; , vesicular monoamine transporter 2

. Figure 2‐2 Synthetic pathways of serotonin (a) and dopamine (b)

Brain neurons partly expressing monoaminergic phenotype

2

(> Figure 2‐2). Noteworthy, the enzymatic activity of AADC greatly exceeds that of TH and tryptophan hydroxylase (Moore et al., 1985). DA and serotonin, the final synthetic products in DA‐ergic and serotoninergic neurons, are captured from cytosol to the secretory granules (¼dense core vesicles) by the vesicular membrane transporter 2 (VMAT2) (> Figure 2-1) (Hoffman et al., 1998; Weihe and Eiden, 2000). In noradrenergic and adrenergic neurons, DA is subsequently converted into noradrenaline and adrenaline by intragranular enzymes, DA b‐hydroxylase and phenyletanolamine N‐methyltransferase, respectively (Squire et al., 2003). MAs stored in secretory granules are discharged via exocytosis. After the action on the targets and partial enzymatic degradation, MAs are captured by the MA‐specific membrane transporter into the monoaminergic (MA‐ergic) neurons for subsequent reutilization (> Figure 2-1) (Hoffman et al., 1998). Although MAs act on the neurons in ontogenesis and adulthood via the same specific receptors their final effects are quite different. They provide a long‐lasting irreversible (¼morphogenetic, imprinting) action on the differentiating target neurons regulating the expression of their specific phenotype in ontogenesis and a short‐term reversible action on the differentiated target neurons controlling their functional activity in adulthood (Ugrumov, 1997). In the early sixties, the MA‐ergic neurons containing catecholamines and serotonin have been first detected on sections with the histofluorescent technique (Dahlstro¨m and Fuxe, 1964) that gave an opportunity to map the catecholaminergic and serotoninergic neurons in the brain. Later, this mapping has been confirmed and slightly modified by using additional morphological approaches; autoradiography of the neurons following the administration of the radiolabeled MAs (Beaudet and Descarries, 1979), and immunostaining of the enzymes of MA synthesis (Ho¨kfelt et al., 1984) or MAs (Steinbusch, 1984). The pioneer morphological studies of the Swedish and Dutch groups were followed by an accumulation of physiological evidence that the clusters of MA‐ergic neurons or the so‐called MA‐ergic centers located in a number of brain regions are involved in neural and neuroendocrine regulations of central and visceral functions, e.g., general metabolism, water–mineral metabolism, memory, different types of behavior, circadian rhythms, etc. (Montange and Calas, 1988; Weiner et al., 1988). Although the MA‐ergic neurons have been already studied for about 40 years, there is a discrepancy in their definition. Indeed, until presently, the MA‐ergic neurons are identified on sections by detecting individual specific markers or an incomplete set of specific markers: (1) MAs (mono‐immunolabeling) (Bjo¨rklund and Lindvall, 1984; Steinbusch, 1984); (2) individual, most often first rate‐limiting enzymes of MA synthesis (mono‐immunolabeling) (Ho¨kfelt et al., 1984); (3) the whole set of the enzymes of MA synthesis (double‐ and triple‐labeling) (Ikemoto et al., 1999); (4) MA membrane transporters (mono‐ and double labeling) (Lorang et al., 1994; Hoffman et al., 1998); (5) colocalization of individual enzymes of MA synthesis and MAs (double‐immunolabeling) (Ershov et al., 2002a, b); (6) colocalization of individual enzymes of MA synthesis and the MA transporter (double‐immunolabeling and/or a combination of immunocytochemistry with in situ hybridization) (Lorang et al., 1994; Betarbet et al., 1997; Cossette et al., 2005a); (7) VMAT2 and its colocalization with the earlier markers (Hoffman et al., 1998; Weihe et al., 2006). Based on the data accumulated over the last 40 years, one may define the MA‐ergic neurons as those having specific machinery providing the MA turnover, i.e., the neurons expressing the MA transporter, the whole set of the enzymes of MA synthesis from the precursor amino acid, and VMAT2. However, VMAT2 is a semi‐specific marker being common for all MA‐ergic neurons, catecholaminergic, and serotoninergic neurons (Hoffman et al., 1998; Weihe and Eiden, 2000). The neurons partly expressing MA‐ergic phenotype have been first discovered in the eighties when the brain regions containing the TH‐immunoreactive (IR) neurons but lacking AADC‐immunoreactive neurons, and vice versa were detected (Jaeger et al., 1983, 1984; Okamura et al., 1988b). The existence of the so‐called monoenzymatic neurons was definitely proved by using double‐immunolabeling of the enzymes of DA and serotonin synthesis (Meister et al., 1988; Ikemoto et al., 1998a, b; Ershov et al., 2002a, b). Another type of neurons partly expressing MA‐ergic phenotype possess the MA transporter but lack either the whole set of the enzymes of MA synthesis or only the first rate‐limiting enzyme (De Vitry et al., 1986; Ugrumov et al., 1989a; Hoffman et al., 1998). The number of neurons partly expressing the MA‐ergic phenotype in ontogenesis and in adulthood of mammals with neurodegenerative diseases (hyperprolactinemia and Parkinson’s disease) appears to exceed those in normal adult mammals (Meredith et al., 1999; Porritt et al., 2000; Ershov et al., 2002a).

25

26

2

Brain neurons partly expressing monoaminergic phenotype

The discovery of neurons partly expressing the MA‐ergic phenotype has raised a number of crucial questions: (1) what is their functional significance; (2) whether they are numerous and widely distributed over the brain; (3) if the conventional classification of MA‐ergic neurons based on histofluorescence of MAs (Dahlstro¨m and Fuxe, 1964; Bjo¨rklund and Lindvall, 1984) or immunostaining of the first rate‐limiting enzymes of MA synthesis (Ho¨kfelt et al., 1984) or MAs (Steinbusch, 1984) is still valid. The answer to the last question should a priori be negative. In fact, the neurons possessing only one enzyme fail to synthesize MAs from the precursor amino acid. The existence of the MA transporter also cannot be considered as a specific marker of MA‐ergic neurons. In fact, some nonserotoninergic neurons possessing the serotonin transporter cannot synthesize serotonin from L‐tryptophan (De Vitry et al., 1986). The first attempt to improve the Swedish and Dutch classification of MA‐ergic systems has been made in eighties by mapping of the neurons expressing only AADC. The clusters of these neurons were called ‘‘D’’ groups (Jaeger et al., 1984). Although extensive data have been already accumulated in literature on the neurons partly expressing the MA‐ergic phenotype, their functional role, and the regulation remain far from complete understanding. According to the most promising hypothesis, the neurons partly expressing MA‐ergic phenotype serve to compensate a functional deficiency of MA‐ergic neurons (Ugrumov et al., 2002, 2004; Ershov et al., 2005) arising as a consequence of: (1) a transient decrease of the neuron functional activity; (2) a irreversible decrease of the neuron functional activity, e.g., under neurodegeneration; (3) an increased need of the brain in MAs that cannot be satisfied even by highly stimulated MA‐ergic neurons. The goal of this review is to evaluate the data concerning the location in the brain, development, regulation, and functional significance of the neurons partly expressing MA‐ergic phenotype in norm and pathology.

2

Brain Neurons Partly Expressing the MA‐ergic Phenotype in Adult Mammals

2.1 Neurons Expressing Individual Enzymes of MA Synthesis Particular attention in literature and in this review is focused on the so‐called monoenzymatic neurons expressing individual enzymes of DA synthesis, TH or AADC that is explained by their particularly high frequency and wide distribution in the brain.

2.1.1 Hypothalamus The first attempt to estimate the frequency of monoenzymatic neurons has already shown that at least in the hypothalamus, one of the largest DA‐ergic center in the brain (A12, A13, A14), a number of TH‐ immunoreactive neurons (Van den Pol et al., 1984) greatly exceeded that of DA‐containing (fluorescent) neurons (Bjo¨rklund and Lindvall, 1984). It meant that numerous TH‐containing neurons lack AADC being incapable of DA synthesis. Later, the most interesting information about the monoenzymatic neurons was obtained when studying the hypothalamic arcuate nucleus (AN), suprachiasmatic nucleus (SCN), and the magnocellular supraoptic, paraventricular, and accessory nuclei. Arcuate nucleus. The initial immunocytochemical studies with mono‐immunostaining of the enzymes of DA synthesis have already shown incomplete overlapping in the distribution of TH‐immunoreactive neurons and AADC‐immunoreactive neurons in the AN in rats. The former were distributed all through the AN whereas the latter were mainly located dorsomedially (Okamura et al., 1988b). Similar distribution of TH‐immunoreactive neurons and AADC‐immunoreactive neurons is also a characteristic of the AN in other mammals including primates (Komori et al., 1991; Kitahama et al., 1998). It has been suggested that at least the ventrolateral portion of the AN contains monoenzymatic TH‐expressing neurons (Okamura et al., 1988b, c; Komori et al., 1991; Zoli et al., 1993) that was further proven by using double‐immunolabeling of the enzymes of DA synthesis (> Figures 2-3 and > 2-4) (Ershov et al., 2002a, b). Besides the ventrolateral region of the AN, the monoenzymatic TH‐neurons are located in

Brain neurons partly expressing monoaminergic phenotype

2

. Figure 2‐3 Schematic drawing of the distribution of monoenzymatic TH‐neurons (filled circle), monoenzymatic AADC‐ neurons (opened circle), and bienzymatic (TH and AADC) neurons (asterisk) in the arcuate nucleus of adult rats. Arcuate nucleus is shown unilaterally. DM, dorsomedial region of the arcuate nucleus; VL, ventrolateral region of the arcuate nucleus

the rest of the nucleus. In contrast to monoenzymatic TH‐neurons, most monoenzymatic AADC‐neurons and bienzymatic neurons are located in the dorsomedial region of the AN, and only occasional neurons of either type are scattered in the ventrolateral region (> Figures 2‐3 and > 2‐4) (Ershov et al., 2002a). A population of monoenzymatic TH‐neurons is twice as small as the population of monoenzymatic AADC‐ neurons and the population of bienzymatic neurons in the AN of adult rats (Ershov et al., 2002a). From these quantitative data, it indirectly follows that relatively large populations of monoenzymatic neurons may be of functional importance. Suprachiasmatic nucleus. Besides AN, the SCN, a pacemaker of the circadian rhythmic activity (Klein et al., 1991) is a promising model for the study of monoenzymatic neurons. It has been demonstrated with double‐immunostaining in all studied mammals (rats, hamsters, sheep, shrews, and humans) that the SCN contains numerous monoenzymatic AADC‐neurons (D13) (Jaeger et al., 1983; Battaglia et al., 1995; Kitahama et al., 1998; Ishida et al., 2002) and occasional monoenzymatic TH‐neurons (Battaglia et al., 1995). The former population is located in the ventral and ventrolateral regions of the nucleus whereas the latter population is distributed at the periphery of the nucleus (> Figure 2-5) (Novak and Nunez, 1998). Hypothalamic ‘‘magnocellular’’ nuclei. Since the mid‐eighties, it has been repeatedly demonstrated that vasopressinergic neurons of the hypothalamic ‘‘magnocellular’’ nuclei, the supraoptic nucleus, the dorsolateral paraventricular nucleus, and ‘‘accessory’’ nuclei, are capable to coexpress TH in all studied mammals, rodents, and primates. In contrast to rodents, in humans TH is expressed not only in magnocellular vasopressinergic neurons but also in magnocellular oxytocinergic neurons (Panayotacopoulou et al., 1994). Rare magnocellular neurons coexpress TH in norm whereas their number multiplies under functional stimulation (> Figure 2-6) (Kiss and Mezey, 1986; Abramova et al., 2002). TH synthesized in neuronal cell

27

28

2

Brain neurons partly expressing monoaminergic phenotype

. Figure 2‐4 The number of monoenzymatic TH‐expressing neurons (TH), monoenzymatic aromatic L‐amino acid decarboxylase (AADC)‐expressing neurons, and bienzymatic (TH and AADC) neurons in the ventrolateral (dotted line) and dorsomedial (solid line) regions of the arcuate nucleus (AN) of male rats at the 21st embryonic day (E21), the ninth postnatal day (P9) and in adulthood (Ershov et al., 2002a). Mean SEM. *P < 0.05

bodies is transported via axons and accumulated in axonal terminals in the pituitary posterior lobe (Abramova et al., 2000). Up to now, all the attempts to detect AADC in vasopressinergic neurons were unsuccessful (Kitahama et al., 1998). Other hypothalamic regions. In rodents, monoenzymatic TH‐neurons were detected in the periventricular nucleus, near the SCN (Battaglia et al., 1995), in the lateral preoptic area (Vincent and Hope, 1990), and the zona incerta (Skagerberg et al., 1988) whereas the monoenzymatic AADC‐neurons were found in the dorsomedial nucleus (D12) (Jaeger et al., 1984), premamillary nucleus (D8) (Jaeger et al., 1984; Karasawa et al., 1994; Kitahama et al., 1998), zona incerta (D10) (Skagerberg et al., 1988), lateral hypothalamus (D11), and in the medial preoptic area (Vincent and Hope, 1990; Kitahama et al., 1998).

Brain neurons partly expressing monoaminergic phenotype

2

. Figure 2‐5 Schematic representation of the distribution of tyrosine hydroxylase‐immunopositive neurons (a) and aromatic L‐amino acid decarboxylase‐immunopositive neurons (b) in the hamster suprachiasmatic nuclei (SCN) (Novak and Nunez, 1998). The neurons are represented as dots. OC, optic chiasma; III, third ventricle

2.1.2 Striatum Rodents. Although dozens or even hundreds of immunocytochemical studies have been devoted to the nigrostriatal DA‐ergic system in intact mammals, only few of them described the neurons (cell bodies) containing enzymes of DA synthesis in the striatum and in the near limbic system, mostly nucleus accumbens in normal rodents and primates (Tashiro et al., 1989a, b; Betarbet et al., 1997; Mura et al., 2000; Lopez‐Real et al., 2003). In rats, the number of AADC‐immunoreactive neurons did not exceed 20 per striatum (Tashiro et al., 1989a) whereas TH‐immunoreactive neurons were either as numerous as AADC‐ neurons in number or even fewer (Tashiro et al., 1989b; Lopez‐Real et al., 2003). Besides the striatum, the TH‐immunoreactive neurons (1–3 per 40‐mm thick section) were found with mono‐immunolabeling in the nucleus accumbens of rodents. In most studies, no zonality in the distribution of the striatal monoenzymatic neurons has been recognized, though according to some authors the AADC‐immunoreactive neurons were mainly located in the subcallosal area of the dorsomedial, dorsal, and the dorsolateral striatum (Lopez‐ Real et al., 2003). Most TH‐immunoreactive neurons and AADC‐immunoreactive neurons had a round or oval cell body, which ranged from 10 to 20 mm in diameter, and extended 1–2 spiny processes, whereas rare neurons possessed several spiny dendrites (Tashiro et al., 1989a, b, 1990). Monkeys. In the primates (monkeys and humans), the number of the striatal neurons expressing enzymes of DA synthesis greatly exceeded that in the rodents (Betarbet et al., 1997; Cossette et al., 2005a). In normal

29

30

2

Brain neurons partly expressing monoaminergic phenotype

. Figure 2‐6 Vasopressinergic neurons containing tyrosine hydroxylase, peptide (a, b) and mRNA (c, d), in supraoptic nucleus of intact (a, c) and salt‐loaded (b, d) adult rats (modified from Abramova et al., 2002). OT, optic tract; SON, supraoptic nucleus

monkeys (rhesus, macaques), TH‐immunoreactive neurons detected with mono‐immunolabeling were found in the caudate nucleus and putamen, mostly at their periphery (Dubach et al., 1987; Betarbet et al., 1997; Tande´ et al., 2006). Dorsally, the neurons were concentrated toward the dorsal border of the striatum near the corpus callosum. Ventrally, the striatal distribution of TH‐immunoreactive neurons appeared to be continuous with DA‐ergic neuron populations in the ventrobasal regions of the forebrain. Along the lateral edge of the putamen, the neurons were present in the neuropil and in the adjacent white matter. A few neurons were also located within the anterior limb of the internal capsule where it separated the caudate from the putamen. The AADC‐immunoreactive neurons (mono‐labeling) were distributed sparsely in the caudate nucleus, putamen, and the nucleus accumbens in the monkeys (Ikemoto et al., 1998a). Most of the TH‐immunoreactive neurons were round or oval in shape and small in size (10–18 mm) having from two to five primary aspiny processes. A few ( Figure 2-7) (Cossette et al., 2005a; Huot et al., 2007). Moreover, a small number of TH‐neurons were found in ventral margin of the nucleus accumbens (Ikemoto et al., 1998a). The most frequent TH‐immunoreactive neurons (58%) have a medium‐sized (diameter 20–24 mm) or small‐sized (10–15 mm) cell bodies with 3–4 varicose aspiny dendrites (Ikemoto et al., 1998a; Cossette et al., 2005a; Huot et al., 2007). Although aspiny TH‐neurons are distributed throughout the rostrocaudal extent of the caudate nucleus and the putamen, they predominate in the ventral striatum (> Figure 2-7). The

Brain neurons partly expressing monoaminergic phenotype

2

. Figure 2‐7 Schematic representation of the distribution of tyrosine hydroxylase‐immunoreactive aspiny (circle) and spiny (star) neurons on frontal sections through the human basal ganglia in rostrocaudal (a, b) extension (Cossette et al., 2005a). AC, anterior commissure; CD, caudate nucleus; CL, claustrum; GPe, external segment of the globus pallidus; GPi, internal segment of the globus pallidus; IC, internal capsule; NA, nucleus accumbens; PUT, putamen; LV, lateral ventricle

minor population of TH‐IR neurons is represented by large neurons (6.5%) (25–35 mm) with spiny dendrites and few TH‐immunoreactive cells displaying mixed neuron–glial morphology. The latter have round or ovoid cell body (10–12 mm) with up to 10 aspiny processes. In contrast to aspiny TH‐IR neurons, the spiny TH neurons were detected in the caudate nucleus but not in the putamen (> Figure 2-7). The TH neurons with neuron–glial morphology are confined to the striatal bridges that link the caudate nucleus and the putamen rostrally. A single human striatum contains in average 331 TH‐immunoreactive neurons. However this number could have been increased by about 10 times if the prolific zone, containing about 3000 cells, observed in some brains had been taken into account (Cossette et al., 2005a). According to the mono‐immunolabeling study, rather numerous AADC‐immunoreactive neurons were distributed in the entire rostrocaudal extent of the striatum and the nucleus accumbens (> Figure 2-8) though their number varies significantly from human to human. The most numerous AADC‐neurons were located in the putamen (
120 neurons per 50 mm thick section), less numerous ones were situated in the nucleus accumbens (
90 neurons per 50 mm thick section), and the least number of AADC‐ immunoreactive neurons was observed in the caudate nucleus (
30 neurons per 50 mm thick section) (Ikemoto et al., 2003). Most AADC‐neurons are fusiform, bipolar, or multipolar in shape, medium to large in size (15–30 mm) having thick dendritic arbors (Ikemoto et al., 1997).

2.1.3 Other Brain Regions Cortex. Rather numerous monoenzymatic TH‐neurons and AADC‐neurons were detected with the double‐ immunolabeling of TH and AADC in the cortex, e.g., in the human anterior cingulate cortex (Ikemoto et al., 1999). These neurons lack VMAT2 (Weihe et al., 2006). The certain zonality in the distribution of TH‐neurons within the cortex has been recognized: the highest concentration is a characteristic of multi‐ associative cortex, a moderate concentration is typical for the unimodal associative cortex, and a minor concentration is an attribute of the visual, sensory, and motor primary areas (Gaspar et al., 1987). Substantia nigra and the ventral tegmental area. In addition to numerous DA‐ergic neurons innervating the striatum, the compact zone of the substantia nigra contains rare monoenzymatic TH‐neurons and AADC‐neurons that was shown initially in humans (Ikemoto et al., 1998b). In contrast to the substantia

31

32

2

Brain neurons partly expressing monoaminergic phenotype

. Figure 2‐8 Schematic representation of the distribution of aromatic L‐amino acid decarboxylase‐immunoreactive neurons (dots) in the human striatum (Ikemoto et al., 1997). Acc, nucleus accumbens; Cn, caudate nucleus; Pu, putamen

nigra, rather numerous monoenzymatic TH‐neurons and AADC‐neurons were found in the ventral tegmental area (Ikemoto et al., 1998b). Other brain regions. Besides the hypothalamus, cortex, and the striatum, monoenzymatic TH‐neurons were found in the basal ganglia and in the globus pallidus in particular that was initially shown in the humans (Komori et al., 1991). Moreover, the clusters of monoenzymatic TH‐neurons were detected in hamsters in the nucleus of the horizontal limb of the diagonal band, pericentral nucleus of the inferior colliculus, lateral parabrachial nucleus, the dorsal motor nucleus of the vagus, and the substantia innominata, beneath the rostral globus pallidus (Vincent and Hope, 1990). Rare TH‐neurons are located in the anterior olfactory nucleus in mice (Nagatsu et al., 1990) and rats (Meredith et al., 1999). The monoenzymatic AADC‐neurons were observed in rodents in the anterior olfactory nucleus, pretectal nucleus (D5), the nucleus of the solitary tract (D2) (Karasawa et al., 1994), and in the internal division of the lateral parabrachial nucleus (Vincent and Hope, 1990). Moreover, a small number of TH‐ immunoreactive and AADC‐immunoreactive Purkinje cells were detected with mono‐immunostaining in the cerebellum (Sakai et al., 1995). Taking into account that the number of TH‐immunoreactive neurons is twice as large as AADC‐immunoreactive neurons, at least half of TH‐immunoreactive neurons should be monoenzymatic in nature (Sakai et al., 1995).

2.2 Bienzymatic TH- and AADC-Expressing Neurons According to the double‐immunolabeling studies, the striatum of rodents, monkeys, and humans contains bienzymatic (TH and AADC) neurons in addition to the monoenzymatic ones (Ikemoto et al., 1998a; Weihe et al., 2006). However, in contrast to true DA‐ergic neurons, they lack VMAT2 (Weihe et al., 2006). The bienzymatic neurons are distributed along the ventral margin of the rostral nucleus accumbens and the caudate nucleus. They were small‐ to medium‐sized (10–15 mm), elongated in shape, having one or two processes (Ikemoto et al., 1998a).

Brain neurons partly expressing monoaminergic phenotype

2

2.3 Non-MA-ergic Neurons Expressing the MA Transporters Up to the present, no special studies of the expression of the MA transporters in non‐MA‐ergic neurons have been made by using multi‐labeling of at least two first enzymes of MA (DA or serotonin) synthesis and the MA transporter. Nevertheless, according to indirect evidence, at least two types of non‐MA‐ergic neurons expressing the MA transporter may be recognized: the monoenzymatic neurons and the neurons lacking enzymes of MA synthesis. Some neurons express the serotonin transporter while others – the DA transporter (> Figures 2-9 and > 2-10) (Lorang et al., 1994; Hoffman et al., 1998).

. Figure 2‐9 Neurons containing either tyrosine hydroxylase or dopamine transporter mRNA on successive coronal levels of the rat hypothalamus in rostrocaudal extension (a–d) (Lorang et al., 1994). Open squares, TH‐immunoreactive neurons; open circles, DAT mRNA‐containing and TH‐immunonegative neurons; solid circles, double‐labeled (TH‐immunoreactive, DAT mRNA‐containing) neurons. AH, anterior hypothalamic nucleus; AN, arcuate nucleus; DM, dorsomedial nucleus; LPA, lateral preoptic area; ME, median eminence; MPA, medial preoptic area; OC, optic chiasm; PV, paraventricular nucleus; VM, ventromedial nucleus; III, third ventricle; ZI, zona incerta

33

34

2

Brain neurons partly expressing monoaminergic phenotype

. Figure 2‐10 Schematic representation of the distribution of neurons containing vesicular monoamine transporter 2 mRNA (left hemisphere of each level) or serotonin (open square) and dopamine (filled triangles) transporter mRNAs (right hemisphere of each level) at coronal successive levels in rostrocaudal extension of the adult rat hypothalamus (Hoffman et al., 1998). Open circles, overlap of vesicular monoamine transporter 2 mRNA with either serotonin transporter or dopamine transporter; filled circles, neurons expressing vesicular monoamine transporter 2 mRNA which do not correspond to neurons expressing either serotonin transporter or dopamine transporter mRNAs. AC, anterior commissure; AH, anterior hypothalamic nucleus; AN, arcuate nucleus; DM, dorsomedial nucleus; DP, dorsal premamillary nucleus; LH, lateral hypothalamus; PH, posterior hypothalamic nucleus; MP, medial preoptic area; MV, mamillary nucleus, ventral part; OC, optic chiasm; PV, paraventricular nucleus; VM, ventromedial nucleus; VP, ventral premamillary nucleus; SC, suprachiasmatic nucleus; ZI, zona incerta; III, third ventricle

Neurons expressing the serotonin transporter. Up to the present, the only one population of non‐MA‐ ergic neurons expressing a serotonin transporter has been recognized with certainty. These small oval mono‐ and bipolar neurons with short unbranched processes were detected in the hypothalamic dorsomedial nucleus in rats. They contain serotonin transporter mRNA (> Figure 2-10) (Hoffman et al., 1998) and the functionally active protein that is manifested by: (1) the radiolabeling of the neurons following 3 H‐serotonin injections to the cerebral ventricles (Beaudet and Descarries, 1979; Ugrumov et al., 1986);

Brain neurons partly expressing monoaminergic phenotype

2

(2) the appearance of the serotonin fluorescent neurons after the intraventricular injections of serotonin (Fuxe and Ungerstedt, 1968); and (3) the neuron degeneration provoked by 5,7‐dihydroxytryptamine, the neurotoxin of serotoninergic neurons (Frankfurt and Azmitia, 1983). These neurons are not serotoninergic in nature since they lack tryptophan hydroxylase, the first rate‐limiting enzyme of serotonin synthesis, and the presence of AADC is questionable (Frankfurt et al., 1981; Ugrumov et al., 1989a). Similar neurons were found in the substantia nigra, the ventral tegmental area, the nucleus sagulum, the Barrington nucleus, and the parvicellular reticular nucleus (Hoffman et al., 1998). Neurons expressing the DA transporter. Although there is no convincing evidence of the existence of non‐ MA‐ergic neurons expressing the DA transporter, some data may be interpreted in favor of this idea. One can state that at least a part of the striatal monoenzymatic TH‐neurons coexpress the DA transporter since it is colocalized in the whole population of striatal TH‐immunoreactive neurons (Porritt et al., 2000; Cossette et al., 2005b; Tande´ et al., 2006) composed of monoenzymatic TH‐neurons and bienzymatic (TH and AADC) neurons (Lopez‐Real et al., 2003). In this context, the author’s statement that all the TH‐immunoreactive neurons coexpressing the DA transporter are obligatory DA‐ergic in nature (Porritt et al., 2000; Cossette et al., 2005a, b; Tande´ et al., 2006) appears to be doubtful. In addition to monoenzymatic TH‐neurons, the DA transporter is coexpressed in the neurons lacking TH. Since the authors did not use a triple‐labeling technique, the DA transporter coexpression in monoenzymatic AADC‐neurons cannot be excluded. The neurons possessing the DA transporter but lacking TH have been detected in the zona incerta (> Figure 2-9), ventral premamillary nucleus (Lorang et al., 1994), Barrington nucleus, and in the pontine tegmentum (Hoffman et al., 1998).

3

Brain Neurons Partly Expressing MA‐ergic Phenotype in Mammals in Ontogenesis

Major data on the monoenzymatic neurons in the brain in ontogenesis are rather fragmentary with an exception of the developing hypothalamus and the AN in particular.

3.1 Neurons Expressing Individual Enzymes of MA Synthesis 3.1.1 Hypothalamus Arcuate nucleus. According to the mono‐immunolabeling data, TH‐immunoreactive neurons first appear in the developing AN in rats on the 17–18th day of prenatal life (Daikoku et al., 1986; Ugrumov et al., 1989b). In the initial studies, these neurons were mistakenly considered as DA‐ergic neurons based on the conventional Swedish classification (Daikoku et al., 1986; Ugrumov et al., 1989b). Later, it has been shown by using mono‐immunostaining of TH and AADC on adjacent mirror sections that only TH is expressed in the neurons of the AN on the 18th fetal day (Balan et al., 2000). At that time and later, on the 20–21st embryonic day, most monoenzymatic TH‐neurons were located in the ventrolateral region of the nucleus (> Figure 2-11) (Balan et al., 2000). In contrast to TH‐immunoreactive neurons, the AADC‐immunoreactive neurons first appear in the rat AN on the 20th fetal day. Since this time onward, monoenzymatic AADC‐immunoreactive neurons are located mainly in the dorsomedial region of the nucleus (> Figure 2-11) being in perinatal rats as numerous as monoenzymatic TH‐immunoreactive neurons (> Figure 2-4). Subsequent application of the double‐ immunolabeling of TH and AADC has shown that the AN of rat fetuses contains more than 99% monoenzymatic TH‐neurons and AADC‐neurons (1:1), and less than 1% bienzymatic neurons at the end of prenatal life (> Figure 2-12) (Balan et al., 2000; Ershov et al., 2002a). The total number of monoenzymatic TH‐neurons, monoenzymatic AADC‐neurons, and bienzymatic DA‐ergic neurons in the AN was about 850, 850, and 17, respectively (> Figure 2-4). From the ninth postnatal day to adulthood, the number of monoenzymatic TH‐neurons decreases significantly (
430 neurons per nucleus), the number of monoenzymatic AADC‐neurons remains at the

35

36

2

Brain neurons partly expressing monoaminergic phenotype

. Figure 2‐11 Tyrosine hydroxylase‐immunoreactive neurons (a) and aromatic L‐amino acid decarboxylase‐immunoreactive neurons (b) (Balan et al., 2000); schematic representation of their distribution in the arcuate nucleus of rat fetuses on the 21st day of intrauterine development. AN, arcuate nucleus; DM, dorsomedial region of the AN; ME, median eminence; VL, ventrolateral region of the AN; III, third ventricle

same level as in fetuses (
850 neurons per nucleus), and the number of bienzymatic neurons greatly increases (
1200 neurons per nucleus). However in adulthood, the monoenzymatic neurons are still as numerous as the bienzymatic ones (> Figure 2-4) (Ershov et al., 2002a). Although the monoenzymatic and bienzymatic neurons are distributed all through the AN in postnatal rats, most monoenzymatic TH‐neurons are located in the ventrolateral region whereas the monoenzymatic AADC‐neurons and bienzymatic neurons are concentrated in the dorsomedial region (> Figures 2-4 and > 2-12) (Ershov et al., 2002a). From the end of prenatal life to adulthood, the monoenzymatic TH‐neurons, AADC‐neurons and bienzymatic neurons were often seen in close topographic relations (Ershov et al., 2002a). According to confocal microscopy, the neurons containing enzymes of DA synthesis are in appositions both at the level of the distal axons in the median eminence and at the level of cell bodies in the AN (> Figures 2-13 and > 2-14) (Ershov et al., 2002b). The frequency of these contacts increases gradually with age, mainly due to the overgrowth and ramification of the neuron processes. In the previous ontogenetic studies, the fibers mono‐immunostained for TH or possessing histofluorescent DA detected in the developing median eminence have a priori been considered as bienzymatic DA‐ergic in nature (Ibata et al., 1982; Ugrumov et al., 1989c). A subsequent application of the double‐ immunostaining of the enzymes of DA synthesis first made it possible to change this consideration by evaluating the ingrowth of monoenzymatic and bienzymatic fibers to the median eminence in ontogenesis, separately. The density of monoenzymatic TH‐axons, i.e., the number of axons per unit of the volume of the

Brain neurons partly expressing monoaminergic phenotype

2

. Figure 2‐12 The number of monoenzymatic tyrosine hydroxylase‐immunoreactive neurons (TH), monoenzymatic aromatic L‐amino acid hydroxylase‐immunoreactive neurons (AADC), and bienzymatic tyrosine hydroxylase‐ immunoreactive and aromatic L‐amino acid hydroxylase‐immunoreactive neurons (TH and AADC) in the arcuate nucleus of rat fetuses on the 21st day of intrauterine development (Ershov et al., 2002a). *P < 0.05

median eminence is maximal at the end of prenatal life whereas it decreases gradually during postnatal period (> Figure 2-15) (Ershov et al., 2002b). The age variation in the density of monoenzymatic TH‐axons might be a consequence of: (1) coexpression of AADC in the neurons initially synthesizing only TH; (2) establishment of either neural or hormonal inhibitory controls of TH synthesis (see > Section 8); (3) apoptosis of some monoenzymatic TH‐neurons. However, it should be emphasized that the decrease of the density of monoenzymatic TH‐axons in the median eminence with age does not obligatory mean a decrease of their total number as the volume of the median eminence enlarges significantly in rats during postnatal period (Ugrumov et al., 1985a). In contrast to monoenzymatic TH‐axons, the density of monoenzymatic AADC‐axons increases progressively from the end of fetal life to adulthood (> Figure 2-15) (Ershov et al., 2002b) though the number of monoenzymatic AADC‐neurons does not change over the same period (> Figure 2-4) (Ershov et al., 2002a, b). This seeming controversy might be explained by: (1) an intense ramification of the axonal projections of monoenzymatic AADC‐neurons of the AN; (2) the ingrowth of monoenzymatic AADC‐ axons to the median eminence from the outside of the AN (Jaeger et al., 1984). The density of bienzymatic axons increases gradually during pre‐ and postnatal period that correlates well with the increasing number of bienzymatic neurons in the AN (> Figures 2-4 and > 2-15). From this correlation it indirectly follows that a great portion of bienzymatic axons in the median eminence belongs to the neurons of the AN. Suprachiasmatic nucleus. Although the information about the monoenzymatic neurons in the developing SCN is quite limited, it is known that it contains monoenzymatic AADC‐neurons in young rats (P10), the only studied period of ontogenesis, as in adulthood (Battaglia et al., 1995). Moreover, rare TH‐immunoreactive most probably monoenzymatic neurons were first detected with the electron microscopic immunocytochemistry in the rat SCN even earlier, at the end of prenatal life. From this time onward, they are located in the periphery of the nucleus (Ugrumov et al., 1994).

37

38

2

Brain neurons partly expressing monoaminergic phenotype

. Figure 2‐13 Schematic representation of topographic relations between monoenzymatic tyrosine hydroxylase (TH)‐neurons and monoenzymatic aromatic L‐amino acid decarboxylase (AADC)‐neurons in the arcuate nucleus and median eminence as well as of hypothetical L‐DOPA transfer from the former to the latter (Ugrumov et al., 2002). BV, blood vessels of the hypophysial portal circulation; DA, dopamine

The data on the transient innervation of the SCN with the monoenzymatic TH‐fibers in ontogenesis appears to be of particular interest (Ugrumov et al., 1989c; Beltramo et al., 1994). Indeed, it has been demonstrated that the ventral and ventromedial portions of the SCN are intensely innervated by TH‐immunoreactive fibers in young rodents (> Figure 2-16) (Ugrumov et al., 1989c) that is in contrast to adults. Irrespectively of the sex, TH‐immunoreactive fibers first appear in the rat SCN on the second postnatal day. Thereafter, the number of fibers increases rapidly until reaching maximum at the 10th postnatal day. Then, the fiber density decreases rapidly (> Figure 2-16) up to puberty (Beltramo et al., 1994). TH‐immunoreactive fibers occurred to be non‐MA‐ergic in nature as they lack AADC (Battaglia et al., 1995) and transporters of catecholamines and serotonin. Indeed, 6‐hydroxydopamine (6‐OHDA) and 5,7‐ dihydroxytryptamine, neurotoxins of DA‐ergic and serotoninergic neurons, which are captured by the MA transporters, did not provoke the degeneration of TH‐immunoreactive fibers (> Figure 2-16) (Beltramo et al., 1994). Most TH‐immunoreactive fibers innervating the SCN belong to a loose accumulation of small monoenzymatic TH‐neurons scattered in the vicinity of the SCN, mainly in the ventral portion of the anterior hypothalamic nucleus (Mirochnik et al., 2002). These TH‐immunoreactive neurons first appear in rats just after birth, followed by their continuous increase in number for subsequent two weeks. Then, the population of TH‐immunoreactive neurons diminishes rapidly and disappears by puberty, synchronously with the disappearance of the monoenzymatic TH‐fibers in the SCN. The axons of TH‐immunoreactive neurons of

Brain neurons partly expressing monoaminergic phenotype

2

. Figure 2‐14 Axo‐somatic specialized‐like contact (arrowhead) between monoenzymatic tyrosine hydroxylase (TH)‐immunoreactive axon (light grey) and monoenzymatic aromatic L‐amino acid decarboxylase (AADC)‐immunoreactive cell body in the arcuate nucleus of the prepubertal rat (Ershov et al., 2002b). Confocal microscopy: double‐labeling of TH and AADC (a) and mono‐labeling of AADC (b) and TH (c). Dotted line, outlined AADC‐immunoreactive neuron

this location are often oriented toward the SCN, and one may follow some of them on thick sections up to the SCN. In addition to monoenzymatic TH‐neurons in the anterior hypothalamic nucleus, rare TH‐ neurons located in the periphery of the SCN or in the next periventricular nucleus (Battaglia et al., 1995) appear to contribute to its transient innervation. Hypothalamic magnocellular nuclei. In ontogenesis, TH is initially expressed in the magnocellular vasopressinergic neurons of rats at the end of the second postnatal week. This is manifested by the appearance of rare neurons possessing TH‐immunoreactive material and TH mRNA in the supraoptic nucleus (Ugrumov, 2002). In contrast to rodents, in humans, TH is expressed not only in differentiating vasopressinergic neurons but also in oxytocinergic neurons (Panayotacopoulou et al., 1994).

3.1.2 Extrahypothalamic Regions The neurons which are TH‐immunoreactive transiently over certain periods of ontogenesis were recognized in many areas of the developing brain. First TH‐immunoreactive neurons increase in number reaching the peak, and then they decrease gradually in number and finally disappear. The transient neuron populations of this kind were found: (1) in the raphe nucleus of the laboratory shrew over the first two weeks of postnatal life (Karasawa et al., 1997); (2) in the anterior olfactory nucleus of mice from the 16th fetal day to puberty with the plateau during the second postnatal week (Nagatsu et al., 1990); (3) in the olfactory bulbs in fetuses of rats and humans (Verney et al., 1996; Izvolskaia et al., 2006); (4) in the limbic system, the bed

39

40

2

Brain neurons partly expressing monoaminergic phenotype

. Figure 2‐15 Density of monoenzymatic TH‐immunoreactive fibers (TH), monoenzymatic aromatic L‐amino acid decarboxylase‐ immunoreactive fibers (AADC), and bienzymatic (TH&AADC) fibers in the median eminence of rats at the 21st embryonic day (E21), the ninth postnatal day (P9) and in adulthood (Ershov et al., 2002b). Mean SEM. *P < 0.05

nucleus of the stria terminalis, and the central nucleus of amygdala in rats from the 17th prenatal day to puberty with the maximal concentration in the early postnatal period (Verney et al., 1988); (5) in the medial geniculate nucleus in mice from the first to the fourth postnatal week with the maximal number during the third week of life (Nagatsu et al., 1996); (6) in the neocortex in rats for the first three weeks of postnatal life (Berger et al., 1985), in the striatum in rat fetuses (> Figure 2-17) (Sorokin et al., unpublished). Some differentiating TH‐neurons synthesize classical neurotransmitters, serotonin (Karasawa et al., 1997), or neuropeptides: gonadotropin‐releasing hormone, somatostatin, and substance P (Verney et al., 1988, 1996; Izvolskaia et al., 2006). According to the double‐immunolabeling studies, differentiating TH neurons do not coexpress AADC (Berger et al., 1985; Verney et al., 1988; Nagatsu et al., 1990, 1996), apparently except the serotoninergic neurons of the raphe nucleus (Karasawa et al., 1997). According to the author’s opinion, the disappearance of TH‐immunoreactive neurons in ontogenesis is explained by: (1) the degeneration of the neurons coexpressing TH; (2) the turning off TH‐expression with age; and (3) the decrease of TH synthesis and hence of the intracellular content of the TH‐immunoreactive material under the level undetectable with immunocytochemistry (Karasawa et al., 1997). The studies of the monoenzymatic AADC‐neurons in ontogenesis are less numerous than those of monoenzymatic TH‐neurons. Nevertheless, it has been shown that between fourteen D groups recognized in adults, nine groups (D1, D4–D7, D10–D12, and D14) first appear in rats during prenatal period, whereas the remaining five groups become detectable with immunocytochemistry during the first postnatal week (Jaeger and Teitelman, 1992). There is a ventrodorsal gradient in the appearance of AADC‐immunoreactive neurons in the developing brain: the neurons of the ventral groups attain AADC‐immunoreactivity prior to the neurons of the dorsal groups.

Brain neurons partly expressing monoaminergic phenotype

2

. Figure 2‐16 Strong innervation of the suprachiasmatic nucleus (SCN) in rats at the tenth postnatal day by tyrosine hydroxylase(TH)‐immunoreactive fibers (a, b) (Ugrumov et al., 1989c) which are resistant to 6‐hydroxydopamine (6‐OHDA) (C, left side) that is in contrast to tyrosine hydroxylase‐immunoreactive fibers of the paraventricular nucleus (PVN) (C, right side) in the same animals (c) (Beltramo et al., 1994). *P < 0.05

. Figure 2‐17 Tyrosine hydroxylase-immunoreactive neurons (arrows) in the striatum and neural epithelium around the lateral ventricles of rat fetuses on the 21st day of intrauterine development (Sorokin et al., unpublished)

Monoenzymatic AADC‐immunoreactive neurons first appear in the rat brain at the 15th embryonic day. At that time, all the neurons are gathered together representing a large cluster located in the intermediate region of the lateral diencephalic vesicle. Later, this accumulation gives rise first to ventral groups located in the lateral hypothalamic area (D11) and in the anterior hypothalamus, medial to the medial forebrain bundle (D14), and then to dorsal groups distributed in the dorsal mesencephalon (D5) and in the mid‐hypothalamus in its rostrocaudal extent. Rostrally, this group extends to the level of the bed nucleus segregating into the most anterior cluster adjacent to the lateral preoptic area (D11) and the accumulation located just beyond the anterior commissure (D14). From the 16th to the 19th prenatal day, a number of additional clusters of monoenzymatic AADC‐immunoreactive neurons become visible. They are

41

42

2

Brain neurons partly expressing monoaminergic phenotype

located in the dorsal mesencephalon (D4) and the dorsal diencephalon (D6, D7, and D10), as well as in the dorsomedial nucleus (D12), thalamus (D7), and the lateral habenular nucleus (D6). From the earlier data it follows that most accumulations of AADC‐neurons appear in rats by the end of intrauterine development, except groups D2, D3, D8, D9, and D13 which appear during the early postnatal period (Jaeger and Teitelman, 1992).

3.2 Non-MA-ergic Neurons Expressing the MA Transporters Neurons partly expressing the serotoninergic phenotype, the high‐affinity serotonin uptake, but lacking either both enzymes of serotonin synthesis or tryptophan hydroxylase have been discovered in the rat diencephalon in ontogenesis (Ugrumov et al., 1986, 1989a) as in adulthood (see, > Section 3.3). In contrast to adult rats, in fetal and young rats these neurons are more widely distributed in the forebrain, and transiently express the serotonin transporter (Ugrumov et al., 1986, 1989a; Gaspar et al., 2003). The neurons of this kind were found in the hypothalamus: the dorsomedial nucleus, the SCN, and the preoptic area (> Figure 2-18), as well as in the thalamus, limbic cortex, retina, and superior olivary nucleus of rodents from the 15th embryonic day to the tenth postnatal day (Ugrumov et al., 1986, 1989a; Lebrand et al., 1998; Gaspar et al., 2003). The neurons are quite different in morphology even within the same brain area. Indeed, the neurons of the hypothalamic dorsomedial nucleus differed considerably in their appearance from those in the anterior

. Figure 2‐18 Schematic representation of the distribution of nonmonoaminergic neurons expressing serotonin transporter in the diencephalon of perinatal rats (Ugrumov et al., 1986, 1989c). AC, anterior commissure; DM, dorsomedial nucleus; LV, lateral ventricle; MFB, medial forebrain bundle; OC, optic chiasm; ON, optic nerve; PA, preoptic area; SC, suprachiasmatic nucleus; III, third ventricle

Brain neurons partly expressing monoaminergic phenotype

2

hypothalamus. The former population was represented by uni‐ or bipolar neurons small in size (5–8 mm) and round in shape with slightly ramified processes, whereas the latter population composed of bipolar neurons larger in size (6  8  15 25 mm) and oval in shape with long widely ramified processes (> Figure 2-19) (Ugrumov et al., 1986, 1989a). . Figure 2‐19 Neurons expressing serotonin transporter (arrows) in the preoptic area of rat fetuses (18th fetal day) (a, b) and in dorsomedial nucleus in young rats (tenth postnatal day) (c, d) at low (a, c) and high (b, d, e) magnifications. Neurons become serotonin‐immunoreactive after systemic treatment with serotonin precursors (a–d) and capture specifically intraventricularly injected tritiated serotonin (E). Both types of labeling were prevented by preliminary injection of inhibitor of serotonin uptake (Ugrumov et al., 1986, 1989a). DM, dorsomedial nucleus; PA, preoptic area; III, third ventricle

The neurons of the SCN and the dorsomedial nucleus possessing the serotonin transporter have been detected in fetal and neonatal rats with autoradiography following the 3H‐serotonin intraventricular injection (Ugrumov et al., 1986), whereas the existence of non‐MA‐ergic neurons expressing the serotonin transporter in the lateral preoptic area was proven by using a combination of immunocytochemistry for serotonin and a pharmacological approach. The latter became detectable with immunocytochemistry for serotonin but only after the animal treatment with L‐tryptophan or 5‐hydroxytryptophan, serotonin precursors, and pargyline, the inhibitor of MA oxidase. However, this effect has been abolished by the preliminary treatment with fluoxetine, an inhibitor of the serotonin uptake (Ugrumov et al., 1989a). According to the author’s interpretation, the pretreatment with the serotonin precursor stimulated serotonin synthesis and release in serotoninergic neurons that was followed by the serotonin uptake from the extracellular space to nonserotoninergic neurons expressing the serotonin transporter (Ugrumov et al., 1989a). In addition to the in vivo studies, the hypothalamic neurons possessing the serotonin transporter have been detected in the primary tissue culture of the embryonic hypothalamus (De Vitry et al., 1986). Although a number of studies have provided indirect evidence of the existence of the non‐MA‐ergic neurons expressing the DA transporter in the brain of adult mammals, the information about the neurons of this kind in ontogenesis is not still available.

43

44

2 4

Brain neurons partly expressing monoaminergic phenotype

Functional Properties and Functional Significance of the Neurons Partly Expressing the MA‐ergic Phenotype

The discovery of non‐MA‐ergic neurons expressing individual enzymes of MA synthesis has raised a question about their functional properties and functional significance.

4.1 Monoenzymatic Neurons Expressing TH TH is represented in monoenzymatic neurons by the same isoforms (Marsais et al., 2002) as in DA‐ergic neurons of the substantia nigra (Nagatsu, 1995). Furthermore, in most monoenzymatic neurons, TH is enzymatically active providing L‐DOPA synthesis from L‐tyrosine as a final synthetic product. This was proven by a number of observations made in the brain areas possessing mostly monoenzymatic TH‐ neurons: (1) the neurons being immunoreactive for L‐DOPA but not for DA (Meister et al., 1988; Okamura et al., 1988a, b, c); (2) the neurons lacking DA histofluorescence even after systemic administration of exogenous L‐DOPA (Zoli et al., 1993); and (3) the neurons showing a high level of L‐DOPA synthesis (> Figure 2-20) (Melnikova et al., 1999; Ugrumov et al., 2002). The neurons with L‐DOPA synthesis as a final synthetic product were detected with certainty in the AN, substantia nigra, ventral tegmental region, and the raphe nucleus (Mons et al., 1989). . Figure 2‐20 þ L‐DOPA concentration and K ‐stimulated release in the mediobasal hypothalamus (arcuate nucleus and median eminence) of rat fetuses at the 21st day of intrauterine development (Melnikova et al., 1999). Mean SEM. *P < 0.05

Nevertheless in some monoenzymatic neurons, TH fails to convert L‐tyrosine to L‐DOPA most probably because of the absence of tetrahydrobiopterin, the specific cofactor, synthesized with guanosine triphosphate (GTP) cyclohydroxylase I (Nagatsu et al., 1997). Indeed, the neurons of this kind: hypothalamic magnocellular vasopressinergic neurons (Marsais et al., 2002), cerebellar Purkinje cells (Sakai et al., 1995),

Brain neurons partly expressing monoaminergic phenotype

2

the neurons of the developing anterior olfactory nucleus (Nagatsu et al., 1990), and the neurons of the geniculate nucleus (Nagatsu et al., 1996) are immunoreactive for TH, but immunonegative for L‐DOPA and GTP cyclohydroxylase I. Therefore, the Weihe et al. (2006) suggestion to call all the monoenzymatic TH‐neurons ‘‘dopaergic’’ neurons is incorrect. At least a part of monoenzymatic TH‐neurons does not coexpress the DA transporter. For instance, the monoenzymatic TH‐neurons in the ventrolateral portion of the AN lack the DA transporter mRNA and protein (> Figure 2-21) (Hoffman et al., 1998) that is in line with the biochemical observation of the low‐ affinity DA uptake in the whole AN (Moore et al., 1985; Melnikova et al., 1999). Similar conclusion has been

. Figure 2‐21 Tyrosine hydroxylase (TH), dopamine transporter (DAT) and vesicular monoamine transporter 2 (VMAT2) expression in the arcuate nucleus: TH expression in both dorsomedial (arrow) and ventrolateral (arrowhead) regions of the arcuate nucleus (a); DAT expression in the dorsomedial but not in the ventrolateral region (b); VMAT2 expression in the dorsomedial region (c) (Hoffman et al., 1998). ME, median eminence; III, third ventricle

45

46

2

Brain neurons partly expressing monoaminergic phenotype

made when studying the SCN of young rats (Beltramo et al., 1994). It has been shown that the monoenzymatic TH‐fibers innervated the SCN during the early postnatal period (Ugrumov et al., 1989c; Beltramo et al., 1994) lack the DA and serotonin transporters. This was proven by the failure of 6‐OHDA and 5,7‐dihydroxytryptamine, neurotoxins of DA‐ergic, and serotoninergic neurons, to provoke the degeneration of TH‐immunoreactive fibers (> Figure 2-16) (Beltramo et al., 1994). The TH‐immunoreactive neurons lacking the DA transporter are widely distributed in the brain, particularly in the diencephalon. They were found in the periventricular nucleus (A14), preoptic area (A15), supraoptic nucleus, and the posterior hypothalamus (A11), as well as outside the diencephalon in the bed nucleus of the stria terminalis (A15), rostral liner nucleus, rostral periaqueductal gray (A11), over the interpeduncular nucleus (> Figure 2-9) (Lorang et al., 1994). The authors probably made a mistake considering these TH‐immunoreactive neurons as DA‐ergic neurons (Lorang et al., 1994). That is why they found it difficult to interpret their functional significance being obliged to declare that DA produced by these ‘‘DA‐ergic’’ neurons provide a paracrine action on the target brain cells or an endocrine action on the pituitary lactrotrophes. This is probably not the true as the pituitaries of mice lacking the DA transporter fail to develop lactrotrophes (Bosse et al., 1997). The difficulty in interpretation of the functioning of diencephalic TH‐immunoreactive neurons lacking the DA transporter may be overcome if they are considered as monoenzymatic neurons producing L‐DOPA as a final synthetic product. Although L‐DOPA synthesis in most monoenzymatic TH‐neurons has been proven, the mechanism of its storage and release remains uncertain. The fact that all the monoenzymatic TH‐neurons irrespective of their location in the brain of the studied animals (rodents and monkeys) lack VMAT2 (> Figure 2-21) (Hoffman et al., 1998; Weihe et al., 2006), makes it questionable the intragranular accumulation of L‐DOPA. Nevertheless, it has been demonstrated ex vivo and in the primary cell culture of the AN of rat fetuses containing numerous monoenzymatic TH‐neurons but almost lacking bienzymatic DA‐ergic neurons (Ershov et al., 2002a) that a large amount of L‐DOPA is discharged under membrane depolarization (> Figure 2-20) (Melnikova et al., 1999). These data have definitely shown that the monoenzymatic TH‐neurons synthesize and store L‐DOPA which releases in response to the adequate physiological stimuli. L‐DOPA synthesis in the monoenzymatic TH‐neurons raised a question about its functional significance. Three possibilities are discussed in literature. According to the first suggestion, L‐DOPA plays a role of the intracellular signal controlling neuronal metabolism, including synthesis and release of non‐MA neurotransmitters and neuromodulators, mainly of neuropeptides. In fact, TH is coexpressed in peptidergic neurons: (1) permanently in adulthood, e.g., in the neurons of the AN producing GABA (g‐aminobutyric acid), neurotensin, somatostatin, growth hormone‐releasing hormone, dinorphin, and galanin (Everitt et al., 1986; Chaillou et al., 1998); (2) in adulthood transiently under certain physiological conditions, e.g., in magnocellular vasopressinergic neurons under functional stimulation (Yagita et al., 1994); (3) transiently during certain periods of ontogenesis that was shown for the differentiating neurons producing somatostatin and substance P (Verney et al., 1988). It should be mentioned that some authors have considered a priori peptidergic neurons coexpressing TH as catecholaminergic in nature though they did not attempt to detect AADC (Everitt et al., 1986). In addition to vasopressinergic neurons, TH but not AADC is probably coexpressed in a wide range of peptidergic neurons. This suggestion is strongly supported by the overlapping in the distribution of neurotensin‐, galanin‐, growth hormone‐releasing hormone‐, and corticotrophin‐producing neurons coexpressing TH with the monoenzymatic TH‐neurons in the ventrolateral part of the AN (Okamura et al., 1985; Everitt et al., 1986), which almost lacks monoenzymatic AADC‐neurons and bienzymatic DA‐ergic neurons (Ershov et al., 2002a). TH is colocalized in differentiating neurons not only with neuropeptides but also with classical neurotransmitters, e.g., serotonin, or the key enzymes of their synthesis, e.g., choline acetyltransferase (Tinner et al., 1989; Karasawa et al., 1997). The authors believe that these neurons produce L‐DOPA which contributes to the regulation of the neuron differentiation (Verney et al., 1996; Izvolskaia et al., 2006). However, all the attempts to detect L‐DOPA in differentiating noncatecholaminergic TH‐expressing neurons were unsuccessful (Karasawa et al., 1997). According to the second suggestion, L‐DOPA synthesized in monoenzymatic TH‐neurons plays a role of an intercellular signal like classical neurotransmitters or neuromodulators (Misu et al., 2003). The authors have raised a number of arguments in favor of this hypothesis: (1) Caþ2‐dependent mechanism of L‐DOPA

Brain neurons partly expressing monoaminergic phenotype

2

release from the striatum that has been shown in vivo (Nakamura et al., 1992) and in vitro (Goshima et al., 1988); (2) stimulating effect of L‐DOPA via most probably presynaptic b‐adrenoreceptors on the noradrenaline release from the hypothalamic slices that has been shown under the pharmacological inhibition of AADC (Goshima et al., 1991); (3) dose‐dependent stimulating or inhibitory effects of L‐DOPA on the DA release from the DA‐ergic nigrostriatal system mediated most probably via presynaptic D2 receptors (Fisher et al., 2000); and (4) dose‐dependent inhibitory action of L‐DOPA on AADC activity and DA release in the DA‐ergic nigrostriatal system (Fisher et al., 2000). The observation of close topographic relations of monoenzymatic TH‐neurons with the presumptive targets for L‐DOPA, e.g., with the vasoactive intestinal polypeptide‐producing neurons in the SCN in young rats (Battaglia et al., 1995) may also be interpreted in favor of the earlier hypothesis. Although the hypothesis about the L‐DOPA functioning as a neurotransmitter is attractive, some arguments should be considered with caution. Indeed, some principal arguments supporting this concept have derived from the experiments with the AADC inhibition and the presumptive shutdown of catecholamine synthesis (Misu et al., 2003). In practice, it is quiet difficult to inhibit completely the enzymes of synthesis of classical neurotransmitters when using a pharmacological approach. The observation of monoenzymatic TH‐immunoreactive fibers (Ershov et al., 2002b) or the L‐DOPA‐ immunoreactive fibers abutting on the primary capillary plexus of the hypophysial portal circulation in the median eminence (Misu et al., 2003) is considered a morphological evidence of the L‐DOPA delivery from the monoenzymatic TH‐axons to the hypophysial portal circulation. In this particular case, L‐DOPA may play a role of a neurohormone reaching the pituitary via bloodstream and acting on the glandular cells as a neurohormone.

4.2 Monoenzymatic Neurons Expressing AADC In most studied monoenzymatic AADC‐neurons, AADC was shown to be capable of converting L‐DOPA to DA and 5‐hydroxytryptophan to serotonin (Karasawa et al., 1994; Ishida et al., 2002). Either precursor is known to be captured to monoenzymatic AADC‐neurons by the membrane transporter of large neutral amino acids (Sugaya et al., 2001; Ferna´ndez et al., 2005). DA synthesis in monoenzymatic AADC‐neurons has been proven by the appearance of DA‐histofluorescence or DA‐immunoreactive materials after the systemic administration of exogenous L‐DOPA. This was shown for monoenzymatic AADC‐neurons of the SCN (D13) (Ishida et al., 2002), premamillary nucleus (D18), pretectal nucleus (D15), and the nucleus of the solitary tract (D2) (Karasawa et al., 1994). Furthermore, the monoenzymatic AADC‐neurons, e.g., the neurons of the dorsomedial nucleus and some hypothalamic neurons in culture become serotonin‐ immunoreactive after the systemic administration of 5‐hydroxytryptophan, but not L‐tryprophan. The preliminary pharmacological inhibition of AADC has abolished this effect thereby confirming the AADC activity in the monoenzymatic neurons (De Vitry et al., 1986). Interestingly, at least a part of monoenzymatic AADC‐neurons lacks the MA transporters. This has been specifically confirmed by the inability of monoenzymatic AADC‐neurons of the SCN for uptake of the radiolabeled serotonin and DA in young and adult rats (Beaudet and Descarries, 1979; Bosler and Calas, 1982; Ugrumov et al., 1986). Although DA synthesis in monoenzymatic AADC‐neurons has been proven, the mechanism of DA storage and release remains uncertain, particularly because of the absence of VMAT2 (Weihe et al., 2006). Nevertheless, DA is probably synthesized in cytosol of monoenzymatic AADC‐neurons as in catecholaminergic neurons. Regarding the DA release, it has been demonstrated ex vivo and in the primary cell culture that under membrane depolarization a large amount of DA is discharged from the AN of rat fetuses (> Figure 2-22) (Melnikova et al., 1998, 1999) containing a large number of monoenzymatic AADC‐ neurons but almost lacking bienzymatic (TH and AADC) neurons (> Figure 2-12) (Ershov et al., 2002a). On one hand these data strongly suggest that the monoenzymatic AADC‐neurons synthesize and store DA, and on the other hand they are capable of releasing DA in response to the adequate physiological stimuli. Still, the source of L‐DOPA for DA synthesis in monoenzymatic AADC‐neurons, mechanism of the DA store and release should be clarified. Although a wide distribution of monoenzymatic AADC‐neurons all over the brain has been repeatedly reported (Jaeger et al., 1984), little is known about their functional significance. In this context, an

47

48

2

Brain neurons partly expressing monoaminergic phenotype

. Figure 2‐22 Dopamine (DA) concentration and Kþ‐stimulated release in: (a) dissected mediobasal hypothalamus (arcuate nucleus & median eminence) of rat fetuses on the 21st day of intrauterine development; (b) primary cell culture of mediobasal hypothalamus of rat fetuses taken on the 17th day of intrauterine development and maintained for 7 days culture (Melnikova et al., 1999). Mean SEM. *P < 0.05

evaluation of intraneuronal colocalization of AADC with neurotransmitters or neuromodulators is of particular importance. Still, only the AADC colocalization with vasopressin in the SCN neurons has been definitely proven by using double‐immunolabeling technique (Jaeger et al., 1983). Nevertheless, a number of other monoenzymatic AADC‐neurons appear to coexpress neuropeptides also. This suggestion is supported by the overlapping in the distribution of monoenzymatic AADC‐neurons and the enkephalin‐producing neurons in the lateral parabrachial nucleus (D3), as well as the substance P‐ and neurotensin‐producing neurons of the nucleus of the solitary tract (D2) (Jaeger et al., 1984). Taking into account that the monoenzymatic AADC‐neurons were often seen in close topographic relations with cerebral blood vessels (Ugrumov et al., 1989a; Karasawa et al., 1994), one may suggest that these neurons synthesize DA and serotonin from the nearest precursors circulating in blood (Melnikova et al., 2006).

4.3 Ensembles of Monoenzymatic Neurons All the previous attempts to evaluate a functional significance of monoenzymatic neurons were restricted to TH‐neurons and AADC‐neurons, separately. Our idea to consider the neurons expressing individual complementary enzymes of DA synthesis as a functional unit occurred to be more productive for understanding of their physiological role. According to our hypothesis, L‐DOPA synthesized in monoenzymatic TH‐neurons is discharged to the intercellular space and thereafter captured by monoenzymatic AADC‐neurons for further conversion to DA (> Figure 2-23) (Ugrumov et al., 2002). Bearing in mind that neurotransmitters and apparently L‐DOPA diffuse for a long distance along the intercellular clefts (Schneider et al., 1994), the cooperative synthesis of DA may be realized even if monoenzymatic TH‐neurons and AADC‐neurons are located far from each other. The AN of fetal rats containing more than 99% monoenzymatic TH‐neurons and AADC‐neurons, in proportion 1:1 and less than 1% bienzymatic neurons has been used as a model to test the hypothesis about the cooperative synthesis of DA by non‐dopaminergic (non‐DA‐ergic) neurons. It has been demonstrated in the ex vivo and in vitro (primary tissue culture) study that despite a minor number of bienzymatic neurons in the AN of fetal rats, the DA concentration in this local region is higher (> Figure 2-22) (Melnikova et al.,

Brain neurons partly expressing monoaminergic phenotype

2

. Figure 2‐23 Schematic representation of the hypothesis on: (a) cooperative synthesis of dopamine (DA) by monoenzymatic neurons expressing complementary enzymes of DA synthetic pathway (left side); (b) strengthening of DA synthesis in DA‐ergic neurons due to L‐DOPA produced by monoenzymatic tyrosine hydroxylase‐expressing neurons (centre); (c) turning on DA synthesis in serotoninergic neurons due to L‐DOPA produced by monoenzymatic tyrosine hydroxylase‐expressing neurons (right side). AADC, aromatic L‐amino acid decarboxylase; TH, tyrosine hydroxylase; TryH, tryptophan hydroxylase

1999) than in the whole diencephalon or in the whole brain which contains a lot of bienzymatic DA‐ergic neurons (Coyle and Henry, 1973). Furthermore, the L‐DOPA concentration in the AN of fetal rats exceeds three times that of DA (> Figures 2‐20 and > 2‐22) (Melnikova et al., 1999), whereas in the brain regions containing the accumulations of true DA‐ergic neurons, only trace amounts of L‐DOPA are detectable as the enzymatic activity of AADC greatly exceeds that of TH (Moore et al., 1985). Taken together these data have been considered as the indirect indication of DA cooperative synthesis by monoenzymatic neurons. The hypothesis about cooperative synthesis of DA was additionally supported by the observation of close topographic relations between monoenzymatic TH‐neurons and AADC‐neurons in the rat AN that was shown with double‐immunofluorecent labeling of the enzymes in confocal microscopy (Ershov et al., 2002b). Besides simple appositions, sort of specialized‐like junctions were observed. These axo‐somatic contacts were formed by ramified monoenzymatic TH‐axons spreading along a cell body of monoenzymatic AADC‐neurons (> Figure 2-14) (Ershov et al., 2002b). Close topographic relations between monoenzymatic neurons are supposed to serve for increasing an efficacy of the L‐DOPA transfer from the monoenzymatic TH‐neurons to the monoenzymatic AADC‐neurons (> Figure 2-13). The contacts between monoenzymatic TH‐neurons and AADC‐neurons have been observed not only at the level of cell bodies in the AN but also at the level of distal axons including axonal terminals in the external zone of the median eminence (> Figure 2-13). Apart from axo‐axonal contacts, numerous monoenzymatic TH‐axons and AADC‐axons abut on the primary capillary plexus of the hypophysial portal circulation, giving rise to axo‐vascular contacts. The particularly high density of the axo‐vascular contacts of this kind was observed in the lateral region of the median eminence. Despite the great value of double‐immunoflurescent observations in confocal microscopy, the resolution of this technique is not

49

50

2

Brain neurons partly expressing monoaminergic phenotype

sufficient for recognition of true appositions between monoenzymatic neurons. Nevertheless, the very recent electron microscopic study with the double‐immunolabeling of the enzymes of DA synthesis confirmed definitely the existence of direct contacts between monoenzymatic TH‐neurons and AADC‐ neurons (Sorokin et al., unpublished). Despite convincingness of the earlier ex vivo and in vitro data, they may be considered only as indirect evidence of the DA cooperative synthesis in the AN since the input of the minor population of bienzymatic DA‐ergic neurons to DA synthesis cannot be ignored. All the attempts to eliminate completely bienzymatic neurons by using 6‐OHDA were unsuccessful because of the low affinity uptake of DA and hence neurotoxin in this particular brain region (Ershov et al., 2005). The only way to definitely prove the cooperative synthesis of DA by monoenzymatic neurons was to inhibit the L‐DOPA transfer from the monoenzymatic TH‐neurons to monoenzymatic AADC‐neurons. If the hypothesis is valid, this action should provoke a decrease of DA synthesis by monoenzymatic neurons but not by DA‐ergic neurons. In a whole, this should result in a drop of the content and hence synthesis of DA in the AN. Three types of the experimental model could theoretically be used to achieve this result: (1) an inhibition of L‐DOPA release from monoenzymatic TH‐neurons; (2) an anchoring of L‐DOPA in the extracellular space; and (3) an inhibition of the L‐DOPA uptake by the monoenzymatic AADC‐neurons. Finally, the third model has been chosen (> Figure 2-24) (Ugrumov et al., 2004). The presumptive L‐DOPA . Figure 2‐24 Schematic representation (a) and results (b, c) of the experiment showing that the competitive inhibition of neutral amino acid and L‐DOPA transporter with L‐tyrosine (a) resulted in the decrease of dopamine (DA) synthesis in cell suspension of the arcuate nucleus of rat fetuses on the 21st day of prenatal life containing mostly monoenzymatic TH‐neurons and AADC‐neurons (b) and in the increase of DA synthesis in cell suspension of the substantia nigra of the same fetuses containing mostly DA‐ergic neurons (c) (Ugrumov et al., 2004) Tyr (), incubation of cell suspension in the absence of L‐tyrosine; Tyr (þ), incubation of cell suspension in the presence of L‐tyrosine. AADC, aromatic L‐amino acid decarboxylase; TH, tyrosine hydroxylase. Filled oval, neutral amino acid and L‐DOPA transporter

uptake by monoenzymatic AADC‐neurons was inhibited under static or perifusion incubation of the cell suspension of the AN of fetal rats (21st embryonic day) in the presence of a relatively high amount of L‐tyrosine which competes with L‐DOPA for the membrane transporter (> Figure 2-24). The cell suspension of the ventral mesencephalon (substantia nigra) of the same fetuses containing a lot of DA‐ergic

Brain neurons partly expressing monoaminergic phenotype

2

neurons has been used as a control. The total amount of L‐DOPA or DA in the incubation medium and cell extracts after the incubation has been considered as an index of their synthesis rate. The L‐tyrosine administration resulted in almost 50% increase of the total amount of L‐DOPA in the cell suspensions of the AN and the substantia nigra (Ugrumov et al., 2004). This result was quite predictable as L‐tyrosine stimulates L‐DOPA synthesis in any cell with enzymatically active TH because of the semi‐ saturation of TH with L‐tyrosine under physiological concentration (Dairman, 1972). The influence of L‐tyrosine on DA synthesis in the substantia nigra was opposite in sign to that in the AN. It stimulated significantly DA synthesis in DA‐ergic neurons of the substantia nigra as a substrate of synthesis, and inhibited DA synthesis in the AN as a competitive inhibitor of the L‐DOPA transporter (> Figure 2-24). This result was considered as convincing evidence of cooperative synthesis of DA by monoenzymatic TH‐neurons and AADC‐neurons (Ugrumov et al., 2004). Despite detection of a relatively high level of DA synthesis in the AN of rat fetuses ex vivo and in the cell culture (Melnikova et al., 1999), it remained uncertain whether the DA amount is sufficient to provide the inhibitory control of the adenohypophysial prolactin secretion in fetuses as in adulthood (McCann et al., 1984). The pharmacological model of the inhibition of D2 receptors on the adenohypophysial lactotropes by the systemic administration of haloperidol was used to solve this issue (Melnikova et al., 1998). It has been shown that the inhibitory control of the adenohypophysial prolactin secretion is established in rats during last 2 days of the intrauterine development, almost in the absence of bienzymatic DA‐ergic neurons in the AN. This result is considered as an additional argument in favor of cooperative synthesis of DA by monoenzymatic neurons in the AN (Melnikova et al., 1998). L‐DOPA synthesized in monoenzymatic TH neurons may be involved in cooperative synthesis of DA when captured not only to monoenzymatic AADC‐neurons but also to DA‐ergic and serotoninergic neurons (> Figure 2-23) (Arai et al., 1995; Karasawa et al., 1995; Ugrumov et al., 2002; Kannari et al., 2006). Bearing in mind that the enzymatic activity of AADC greatly exceeds that of TH and tryptophan hydroxylase, the admission of extracellular L‐DOPA to catecholaminergic neurons and serotoninergic neurons promotes catecholamine synthesis or triggers DA synthesis, respectively (> Figure 2-23). Cooperative synthesis of DA by monoenzymatic TH‐neurons and AADC‐neurons as well as by monoenzymatic TH‐neurons and either DA‐ergic neurons or serotoninergic neurons is considered as a compensatory reaction under the failure of DA‐ergic neurons (Ugrumov et al., 2002, 2004). The former takes place in the brain regions containing monoenzymatic TH‐neurons and AADC‐neurons, (e.g., in the AN), whereas the latter might be a characteristic of the brain regions possessing monoenzymatic TH‐neurons and either DA‐ergic neurons or serotoninergic neurons (e.g., the AN and the striatum). In addition to monoenzymatic AADC‐neurons, the glial cells and endothelial cells of blood vessels possess enzymatically active AADC (Hardebo et al., 1980; Juorio et al., 1993). These cells are probably capable of synthesizing DA from L‐DOPA derived either from the monoenzymatic TH‐neurons or from general circulation (Melnikova et al., 2006). It should be emphasized that the penetration of DA synthesized in endothelial cells to the brain is prohibited by the blood–brain barrier.

4.4 Non‐MA‐ergic Neurons Expressing the MA Transporters The neurons partly expressing the serotoninergic phenotype have been first recognized in the eighties in rodents, both in adulthood (Frankfurt and Azmitia, 1983; De Vitry et al., 1986) and in ontogenesis (Ugrumov et al., 1986, 1989a), by using autoradiography and a combination of pharmacological approach and immunocytochemistry for serotonin. Later, this discovery has been confirmed by using the direct technical approaches, double‐labeling of serotonin and the serotonin transporter, and sophisticated animal genetic models (Gaspar et al., 2003). It has been definitely shown that these neurons possess the serotonin transporter but lack tryptophan hydroxylase. However, the expression of AADC remains to be under question (De Vitry et al., 1986). It should be stressed that the neurons partly expressing the serotoninergic phenotype is an attribute of a wide range species, from lobsters to humans (Verney et al., 2002; Richards et al., 2003). The non‐MA‐ergic neurons possessing the mechanism of the serotonin uptake most probably serve to capture and store serotonin released from the next serotoninergic axons arising from the raphe nucleus.

51

52

2

Brain neurons partly expressing monoaminergic phenotype

This conclusion is derived from the observation of the accumulation of serotonin‐immunoreactive material in these neurons following the pharmacological stimulation of serotonin synthesis (L‐tryptophan, 5‐hydroxytryptophan, and pargyline) that may be abolished by the preliminary pharmacological inhibition of the serotonin uptake (Ugrumov et al., 1989a). The earlier suggestion is strongly supported by the fact that the non‐MA‐ergic neurons expressing the serotonin transporter are often located around cerebral ventricles (the dorsomedial nucleus, SCN), close to the supraependymal and subependymal plexes of serotoninergic fibers (Ugrumov et al., 1985b), or in the vicinity to the medial forebrain bundle (lateral preoptic area) composed of serotoninergic and other fibers (> Figure 2-18) (Ugrumov et al., 1989a). There is a certain similarity in functioning of the neurons and platelets which do not synthesize serotonin but capture it from plasma with the serotonin transporter (Maurer‐Spurej, 2005). Non‐MA‐ergic neurons serving to store serotonin most probably compensate the local serotonin deficiency that might be of particular importance in ontogenesis during so‐called critical period of serotonin action on the target‐neurons as a morphogenetic factor (Lauder, 1993; Ugrumov, 1997). If the earlier neurons possess AADC in addition to the serotonin transporter, they may synthesize serotonin from extracellular 5‐hydroxytryptophan including that circulating in blood. Indeed, the neurons abutting on the blood vessels become serotonin‐immunoreactive only after systemic pretreatment of the animals with either pargiline, the MA oxidase inhibitor, or 5‐hydroxytryptophan (Ugrumov et al., 1989a). In this case, serotonin may contribute to the regulation of the vascular tone as a vasoconstrictor (Ugrumov et al., 1989a). Although a number of data in adult mammals may be interpreted in favor of the existence of the non‐ MA‐ergic neurons expressing the DA transporter but lacking either TH or AADC, no direct evidence has been yet obtained (see > Section 3.2). Moreover, no data are available about these neurons in the brain in ontogenesis. Nevertheless, by analogy with the non‐MA‐ergic neurons expressing the serotonin transporter, one can suggest that the non‐MA‐ergic neurons expressing the DA transporter serve for capturing and storage of extracellular DA, as well as for DA synthesis from the extracellular L‐DOPA if expressing AADC additionally. It is important to ascertain in the future study whether the DA accumulated in the non‐DA‐ergic neurons can be further discharged. Indeed, Hoffman et al. (1998), who failed to detect VMAT2 mRNA in neurons expressing DA transporter but lacking TH, have stated that these neurons are capable of removing extracellular DA but may not release it.

5

Tuberoinfundibular Neurons Partly Expressing DA‐ergic Phenotype in Hyperprolactinemia

Tuberoinfundibular system consists of DA‐ergic neurons located in the AN and projecting their axons to the median eminence toward the primary vascular plexus of the hypophysial portal circulation (> Figure 2-25). DA delivered from DA‐ergic axons to the hypophysial portal circulation arrives with the bloodstream to the anterior lobe providing an inhibitory control of prolactin secretion via D2 receptors on lactotropes (McCann et al., 1984). Degeneration of DA‐ergic neurons of the tuberoinfundibular nucleus (¼AN in rodents) in humans leads to the development of the syndrome of hyperprolactinemia and finally to the disturbance of reproduction (Serri et al., 2003). Hyperprolactinemia is reproduced in rats by the injection of 6‐OHDA (neurotoxin) (> Figure 2-26) to the cerebral ventricles (Ershov et al., 2005). One to two weeks after the 6‐OHDA administration, 50–60% DA‐ergic neurons of the AN are degenerated (Ershov et al., 2005) that results in 50% decrease of DA synthesis and in doubling of prolactin concentration in plasma (> Figure 2-26) (Ziyazetdinova et al., unpublished). A relatively low level of the neurotoxin‐induced degeneration of DA‐ergic neurons in the AN compared to DA‐ergic neurons of other location in the brain, e.g., in the substantia nigra (Smith and Helme, 1974; Jonsson, 1983) is explained by the low‐affinity uptake of DA in the AN (Demarest and Moore, 1980; Annunziato et al., 1980;Moore et al., 1985) and by the relatively small number of the neurons expressing the DA transporter (Bosler and Calas, 1982; Hoffman et al., 1998). The prolactin concentration in plasma returns to the normal level in rats 1–1.5 months after the treatment with neurotoxin that is a manifestation of the compensation of the functional insufficiency of

Brain neurons partly expressing monoaminergic phenotype

2

. Figure 2‐25 Schematic representation of dopaminergic tuberoinfundibular and nigrostriatal systems and their characteristics in norm and pathology

the tuberoinfundibular DA‐ergic system (> Figure 2-26) (Ziyazetdinova et al., unpublished). In theory, the compensatory mechanisms may be represented by: (1) intensified DA synthesis in survived DA‐ergic neurons due to the increased synthesis and/or activity of TH; (2) stimulated cooperative DA synthesis by monoenzymatic neurons due to the increased number of monoenzymatic neurons or the increased activity of the enzymes in preexisting monoenzymatic neurons; (3) enhanced sensibility of lactotropes to DA as a result of the elevated expression of D2 receptors; and (4) attenuated sensibility of lactotropes to stimulating effects of prolactin‐releasing neurohormones like thyrotropin‐releasing hormone and vasoactive intestinal polypeptide. In reality, normalization of the prolactin secretion in the neurotoxin‐treated rats is accompanied with the increased number of the monoenzymatic TH‐neurons and AADC‐neurons (Ershov et al., 2005) and the augmentation of DA synthesis to normal level (Ziyazetdinova et al., unpublished) that was shown both in vivo and in vitro (perifusion of slices of the AN). Taken together these data strongly suggest that the functional insufficiency of the tuberoinfundibular DA‐ergic system is compensated at least in part by the intensified cooperative synthesis of DA by monoenzymatic neurons. Thus, the functional insufficiency of the tuberoinfundibular DA‐ergic system, e.g., in hyperprolactinemia, is compensated due to the stimulation of DA synthesis most probably by the monoenzymatic neurons.

6

Striatal Neurons Partly Expressing DA‐ergic Phenotype in Parkinson’s Disease

Nigrostriatal DA‐ergic system consists of DA‐ergic neurons located in the compact zone of the substantia nigra which project their axons to the striatum (> Figure 2-25). DA released from the distal axons in the striatum plays a key role in the regulation of a motor behavior. That is why the degeneration of nigrostriatal DA‐ergic neurons in humans leads to the disturbance of the motility which is the crucial mechanism of the Parkinson’s disease pathogenesis (Agid, 1991). Noteworthy, clinical symptoms first appear 25–30 years after the onset of the Parkinson’s disease under degeneration of about 70–80% DA‐ergic neurons in the

53

54

2

Brain neurons partly expressing monoaminergic phenotype

. Figure 2‐26 Schematic representation of the experiment (a) and results (b–d) showing the 6‐hydroxydopamine‐induced degeneration of 50% bienzymatic (TH and AADC) neurons in the arcuate nucleus of adult rats is followed first by decreased DA synthesis and increased prolactin secretion (14th day) and then by the normalization of dopamine synthesis and prolactin secretion. Moreover, the number of monoenzymatic neurons increased significantly (Ershov et al., 2005; unpublished data). AN, arcuate nucleus; AADC, aromatic L‐amino acid decarboxylase; TH, tyrosine hydroxylase, 6‐OHDA, 6‐hydroxydopamine

substantia nigra (> Figure 2-27) and the loss of about 70% DA in the striatum (Agid, 1991). This raised a question what mechanisms of the brain plasticity serve to compensate the failure of the nigral DA‐ergic neurons for a long time and what may be the role of the striatal neurons partly expressing the DA‐ergic phenotype in this phenomenon.

6.1 Monoenzymatic TH‐Expressing Neurons The degeneration of the DA‐ergic neurons in the substantia nigra is accompanied by the increased number of the striatal TH‐immunoreactive neurons in all mammals, rodents (Lopez‐Real et al., 2003), monkeys, and humans studied so far (Betarbet et al., 1997; Porritt et al., 2000; Palfi et al., 2002). As observed from the double‐labeling studies, most of them are monoenzymatic (> Figure 2-28) (Lopez‐Real et al., 2003; Sorokin et al., unpublished). In animals, the maximal – 3.5–7‐fold – increase of TH‐immunoreactive neurons after the degeneration of nigral DA‐ergic neurons has been observed in monkeys (> Figure 2-29) (Betarbet et al., 1997; Palfi et al., 2002). In fact, the total number of the TH‐immunoreactive neurons in the denervated

Brain neurons partly expressing monoaminergic phenotype

2

. Figure 2‐27 Schematic representation of timing characteristics of degeneration of nigral dopaminergic neurons with age in norm and in Parkinson’s disease and its relevance to presymptomatic and symptomatic phases of the disease (adapted from Agid, 1991). Initial number of nigral dopaminergic neurons is considered as 100%

striatum in monkeys greatly exceeded that in rodents (140 neurons per 50 mm‐thick section in monkeys vs. occasional neurons per 40 mm‐thick section in rats). Apparently, this cannot be explained by the species‐ specific difference in the total number of all the striatal neurons only (Betarbet et al., 1997). The decreased number (by sixfold) of the striatal TH‐immunoreactive neurons after degeneration of the nigral DA‐ergic neuron has been described so far in only one paper devoted to the striatum of patients with Parkinson’s disease (Huot et al., 2007). According to the authors’ opinion, the long‐term treatment of these patients with L‐DOPA resulted in a partial compensation of the striatal DA deficiency thereby exerting a hypothetical negative feedback on the TH expression in the striatal neurons. In animals with degenerated nigral DA‐ergic neurons as in intact or control animals, most TH‐immunoreactive neurons were located in the dorsal striatum in the subcallosal area and in the ventral striatum around the anterior commissure (> Figure 2-30) (Meredith et al., 1999; Palfi et al., 2002; Lopez‐ Real et al., 2003). Less numerous TH‐immunoreactive neurons were found scattered in other striatal regions (> Figure 2-30) (Lopez‐Real et al., 2003). Two morphological types of TH‐immunoreactive neurons have been distinguished (Lopez‐Real et al., 2003). Most neurons (99%) of the first type were unipolar, oval in shape and small in size (6–12 mm) having one or two aspiny processes. These were most probably interneurons (Betarbet et al., 1997; Lopez‐Real et al., 2003). Rare neurons ( Figure 2-31) (Betarbet et al., 1997; Lopez‐Real et al., 2003). These neurons were similar in appearance to projection neurons (Lopez‐Real et al., 2003). Some authors emphasized that the TH‐immunoreactive neurons in the denervated striatum were more heavily stained than those in the intact striatum (Betarbet et al., 1997). The 6‐OHDA injection to the substantia nigra in rats resulted in the appearance of new striatal TH‐immunoreactive neurons on the next day and in the progressive increase of their number for two to three subsequent weeks. Then, the TH‐immunoreactive neurons gradually decreased in number for 2–3 months (> Figure 2-32) (Meredith et al., 1999; Lopez‐Real et al., 2003). Similar observations were made in mice after their combined systemic treatment with 1‐methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine (MPTP) and 3‐nitropropionic acid. The latter is a semi‐specific neurotoxin providing a general damaging action on the striatal tissue. After the treatment with 6‐OHDA, first TH‐immunoreactive neurons appear in the striatum on the next day after administration of toxins (5.3 neurons per striatum). However, later the number of these neurons first increased rapidly for 3 days reaching maximum on the fourth day (135 neurons per striatum) and then decreased gradually and disappeared by the 80th day (Nakahara et al., 2001).

55

2

. Figure 2‐28 Mono‐ (A, B, E, F) and double‐immunolabeled (C, D) neurons for either tyrosine hydroxylase (A, B, C, E, F) or aromatic L‐amino acid decarboxylase (D) in the rat striatum after 6‐hydroxydopamine‐induced degeneration of nigral dopaminergic neurons (Sorokin et al., unpublished). A‐D, double‐immunofluorescent for tyrosine hydroxylase and aromatic L‐amino acid decarboxylase in confocal microscopy; E, F, mono‐immunolabeling with peroxidase for tyrosine hydroxylase in conventional microscopy. Arrow, immunoreactive neurons. Bar scale: A, B – 10 mm; C, D – 20 mm; E, D – 40 mm

56 Brain neurons partly expressing monoaminergic phenotype

Brain neurons partly expressing monoaminergic phenotype

2

. Figure 2‐29 Tyrosine hydroxylase‐immunoreactive cell counts in the striatum of control and 1‐methyl‐4‐phenyl‐1,2,3,6‐ tetrahydropyridine(MPTP)‐treated monkeys (Betarbet et al., 1997). *P < 0.05

. Figure 2‐30 Schematic representation of the distribution of tyrosine hydroxylase‐immunoreactive neurons (a) and of aromatic L‐amino acid decarboxylase‐immunoreactive neurons (b) in the rat striatum after 6‐hydroxydopamine‐induced degeneration of nigral dopaminergic neurons (Lopez‐Real et al., 2003). ac, anterior commissure; cc, corpus callosum; v, lateral ventricle

6.2 Monoenzymatic AADC‐Expressing Neurons According to Tashiro et al. (1989b), the number of the striatal AADC‐immunoreactive neurons in rats increases four times under degeneration of the nigral DA‐ergic neurons (12–60 neurons per striatum vs. 3–15 neurons per striatum in the control). Noteworthy, the frequency of the striatal AADC‐immunoreactive neurons appears to be proportional to the damaged area of the compact zone of the substantia nigra. As in intact and control animals, in the neurotoxin‐treated animals most AADC‐immunoreactive neurons are concentrated in the striatum in the subcallosal region, along the lateral ventricle and scattered in other striatal regions (> Figure 2-30) showing a partial overlapping with the location of less numerous

57

58

2

Brain neurons partly expressing monoaminergic phenotype

. Figure 2‐31 Aspiny (a) and spiny (b) tyrosine‐hydroxylase‐immunoreactive neurons in the striatum of 1‐methyl‐4‐phenyl‐ 1,2,3,6‐tetrahydropyridine(MPTP)‐treated monkeys (Betarbet et al., 1997). The insets in a and b show a magnified image of a portion of the dendrite denoted by the arrows

. Figure 2‐32 The number of TH‐immunoreactive neurons at six rostrocaudal levels of the rat striatum (S1–S6) 1, 2, 3, and more than 4 weeks after 6‐hydroxydopamine injection to the raphe nucleus (Meredith et al., 1999)

TH‐immunoreactive neurons (Lopez‐Real et al., 2003). The frequency of the AADC neurons decreased in the rostrocaudal extension of the striatum (Mura et al., 1995, 2000). Striatal AADC‐immunoreactive neurons are small in size (6–10 mm) and oval or spindle‐like in form possessing aspiny processes. They look like interneurons (Mura et al., 1995; Meredith et al., 1999; Lopez‐ Real et al., 2003). According to Meredith et al. (1999), the AADC‐immunoreactive neurons almost disappear with time after the neurotoxin injection, in rats after three weeks.

Brain neurons partly expressing monoaminergic phenotype

2

6.3 Bienzymatic TH‐ and AADC‐Expressing Neurons It has been confirmed by using the double‐immunolabeling of the enzymes of DA synthesis that the striatum of 6‐OHDA‐treated animals contains bienzymatic neurons (> Figure 2-28) as that in intact (control) animals (Lopez‐Real et al., 2003; Sorokin et al., unpublished). However, the proportion of the striatal monoenzymatic and bienzymatic neurons varied when injecting the neurotoxin to different compartments of the nigrostriatal system. Among the neurons expressing enzymes of DA synthesis, most neurons were bienzymatic after the neurotoxin injection to the striatum whereas their portion did not exceed 30% when the neurotoxin was injected to the medial forebrain bundle (Lopez‐Real et al., 2003). It should be emphasized that in bienzymatic neurons the concentration of the TH‐immunoreactive material was always higher than that of AADC‐immunoreactive material. Therefore, the authors supposed that even a part of the TH‐immunoreactive but AADC‐immunonegative neurons may in fact coexpress both enzymes but AADC is not visible because of its low concentration (Lopez‐Real et al., 2003). The observation of the striatal bienzymatic neurons raised a question whether these neurons are DA‐ergic in nature coexpressing the DA transporter. Although nobody has yet attempted to use triple‐ labeling for solving this issue, it was shown with the double‐immunolabeling technique that in the normal and denervated striatum of the monkeys and humans almost all TH‐immunoreactive neurons coexpress the DA transporter and VMAT2. The number of a such type of the neurons multiplies under degeneration of nigral DA‐ergic neurons (Betarbet et al., 1997; Porritt et al., 2000; Cossette et al., 2005; Tande´ et al., 2006). According to Porritt et al. (2000), the TH‐immunoreactive neurons coexpressing the DA transporter are distributed in the striatum of patients with Parkinson’s disease as follows: 39.9% neurons are located in the putamen, 11.6% in the caudate nucleus, 16.3% in the globus pallidus externa, 6.3% in the globus pallidus interna, and 25.9% in the internal capsule and ansa lenticularis. In addition to the DA transporter, the TH‐immunoreactive neurons coexpress the nuclear orphan receptor Nurr1 (Cossette et al., 2004, Hout and Parent, 2007), a transcription factor essential for the expression of the DA‐ergic phenotype by midbrain neurons (Saucedo‐Cardenas et al., 1998). In addition to bienzymatic neurons expressing the DA transporter and VMAT2, the bienzymatic neurons lacking VMAT2 have been recently found thereby raising the question about DA storage and release in these neurons (Weihe et al., 2006). The same discussion may be addressed to this issue as in the case of monoenzymatic AADC‐neurons (see, > Section 5.2).

6.4 Origin, Functional Properties, and Functional Significance of Striatal Neurons Partly or Completely Expressing the DA‐ergic Phenotype If such a large number of striatal neurons expressing TH and AADC somehow contribute to DA synthesis, this compensatory mechanism may be of a substantial functional importance. Therefore, a number of studies have addressed this issue by reproducing parkinsonism in animals, mostly in rodents and monkeys. The authors attempted to determine the origin, functional significance, and regulation of the neurons expressing enzymes of DA synthesis. The degeneration of DA‐ergic neurons in the substantia nigra in animals was provoked by electrochemical lesion or more often by the administration of 6‐OHDA and MPTP. The latter is administered systemically and transformed in the brain to 1‐methyl‐4‐pyridinium, a specific neurotoxin of DA‐ergic neurons, by MA oxidase B (Nakahara et al., 2001). All the earlier manipulations resulted in the appearance of the striatal neurons possessing enzymes of DA synthesis (Betarbet et al., 1997; Meredith et al., 1999; Palfi et al., 2002; Lopez‐Real et al., 2003). This was shown by using immunocytochemistry, in situ hybridization, the reverse transcriptase reaction followed by polymerase chain reaction (Nakahara et al., 2001). Although the striatal neurons expressing enzymes of DA synthesis were detected in parkinsonian animals most often with mono‐immunolabeling, the major TH‐immunoreactive neurons and AADC‐immunoreactive neurons were supposed to be monoenzymatic as they differ to a certain extent in location, size, and morphology (Meredith et al., 1999). Origin of the striatal neurons partly or completely expressing the DA‐ergic phenotype. Two sources of the striatal neurons partly or completely expressing the DA‐ergic phenotype are considered in literature: the

59

60

2

Brain neurons partly expressing monoaminergic phenotype

stem cells of the lateral ventricles and the preexisting neurons, ‘‘silent’’ in norm and expressing enzymes of DA synthesis and the DA transporter under the local DA depletion. The first hypothesis appears to be nonvaluable as the neurons expressing enzymes of DA synthesis and the DA transporter first appear in the striatum on the next day after the neurotoxin administration, i.e., during the too short period for the origin of neuron from the stem cells, its migration to the striatum, and its differentiation. Furthermore, the cells derived from stem cells and migrated to the striatum do not express specific neuronal markers, NeuN, TH, AADC, or GABA‐decarboxylase even three weeks after their appearance (Nakahara et al., 2001). Therefore, the second hypothesis appears to be more convincing. In fact, the enzymes of DA synthesis are most probably coexpressed in preexisting GABA‐ergic neurons under the local DA deficiency that was shown in the in vivo and in vitro studies by using the double‐immuno‐labeling technique (Max et al., 1996; Betarbet et al., 1997; Mura et al., 2000). Functional properties and functional significance of the striatal neurons partly or completely expressing the DA‐ergic phenotype. Despite the absence of direct evidence of the enzymatic activity of TH in the striatal neurons, most authors consider that TH is capable to convert L‐tyrosine to L‐DOPA. The identification of the principal neurotransmitter or neuromodulator in the striatal neurons coexpressing TH may be useful also for understanding their functional significance. It has been demonstrated both in vivo (Betarbet et al., 1997) and in cell culture (Max et al., 1996) that TH is coexpressed in the GABA‐ergic neurons. In the denervated striatum of monkeys, 99% TH‐immunoreactive neurons are GABA‐ergic and only 1% synthesize calbindin, a marker of striatal projection neurons, or parvalbumin, a marker of a distinct set of striatal interneurons (Betarbet et al., 1997). A substantial number of the TH‐immunoreactive neurons expresses the subunit NR1 of the N‐methyl‐D‐aspartate glutamate receptors (26% TH‐neurons) or the subunit GluR1 of the a‐amino‐3‐hydroxy‐5‐methyl‐isoxazole‐4‐propionic acid glutamate receptors (75% TH‐neurons) though no TH‐immunoreactive neurons express subunits GluR2/3 of the a‐amino‐3‐hydroxy‐5‐methyl‐ isoxazole‐4‐propionic acid type of glutamate receptors or subunits mGluR1/5 of the metabotropic glutamate receptors (Betarbet and Greenamyre, 1999). AADC in monoenzymatic striatal neurons is capable of converting L‐DOPA to DA that was proven by the concomitant appearance of the AADC‐immunoreactive neurons and the DA‐immunoreactive neurons identical by morphology and location in the striatum of the rats following the 6‐OHDA‐induced degeneration of the nigral DA‐ergic neurons (Meredith et al., 1999). Similar DA‐immunoreactive neurons were observed in the striatum following the systemic administration of exogenous L‐DOPA (> Figure 2-33)

. Figure 2‐33 Schematic representation of the distribution of aromatic L‐amino acid decarboxylase‐immunoreactive neurons (a) and dopamine‐immunoreactive neurons (b) in the rat striatum after L‐DOPA administration (Mura et al., 2000). AC, anterior commissure; CC, corpus callosum; LV, lateral ventricle

Brain neurons partly expressing monoaminergic phenotype

2

(Mura et al., 1995, 2000). Apparently, DA may be synthesized in monoenzymatic AADC‐neurons not only from exogenous L‐DOPA but also from endogenous L‐DOPA circulating with blood (Melnikova et al., 2006) or synthesizing in the striatal monoenzymatic TH‐neurons. As monoenzymatic TH‐neurons, most monoenzymatic AADC‐neurons were GABA‐ergic, and only rare neurons (4.6%) were peptidergic synthesizing calretenin. All the attempts to detect AADC in peptidergic neurons producing somatostatin and parvalbumin were unsuccessful (Cossette et al., 2005b). One cannot exclude that DA produced in monoenzymatic AADC‐neurons interacts functionally with colocalized GABA or calretinin. Apart from monoenzymatic AADC‐neurons and DA‐ergic fibers survived after the neurotoxin administration, endogenous L‐DOPA can be converted to DA in the striatal serotoninergic fibers (Arai et al., 1996; Lopez et al., 2001; Lopez‐Real et al., 2003; Kannari et al., 2006) that is considered as a compensatory mechanism under the local DA deficiency. This suggestion is supported by a number of observations in the denervated striatum: (1) the colocalization of DA in the serotoninergic fibers after systemic administration of L‐DOPA (Maeda et al., 2005); (2) hyperinnervation of the striatum with serotoninergic fibers; and (3) fourfold decrease of the DA extracellular concentration after the additional degeneration of serotoninergic fibers (Tanaka et al., 1999). In addition to serotoninergic fibers, surviving DA‐ergic fibers and monoenzymatic AADC‐neurons, DA is believed to be synthesized in the striatum in the glial cells and endothelial cells containing AADC (Hardebo et al., 1980). However, this suggestion should be taken with caution as: (1) Nakamura et al. (2000) failed to observe the DA‐immunoreactive cells in the primary striatal culture following the L‐DOPA administration; (2) DA synthesized in the endothelial cells probably cannot penetrate to the brain tissue because of the blood–brain barrier. Degeneration of the nigral DA‐ergic neurons is accompanied by the appearance of the neurons expressing enzymes of DA synthesis not only in the striatum but also in adjacent brain regions. Monoenzymatic TH‐immunoreactive neurons appear in the nucleus accumbens and the olfactory bulbs whereas AADC‐neurons become visible in the cortex and in the bed nucleus of the stria terminalis. If these neurons contribute to DA synthesis, it may further diffuse to the striatum providing a volume transmission effect on the target neurons. This is in agreement with the Schneider et al. (1994) data showing that extracellular DA diffuses in the denervated striatum for a particularly long distance (5–7 mm) that is possible due to the loss of DA‐ergic fibers and hence the decrease of the DA uptake (Bergstrom and Garris, 2003; Bezard et al., 2003). The expression of the enzymes of DA synthesis in non‐DA‐ergic neurons outside the striatum probably represents one of the mechanisms of the so‐called ‘‘passive stabilization’’ serving to maintain a normal level of extracellular DA in the denervated striatum without compensatory change of the DA uptake and release (Bergstrom and Garris, 2003). By analogy with hyperprolactinemia following degeneration of the tuberoinfundibular DA‐ergic neurons, one may expect that the DA cooperative synthesis by striatal monoenzymatic neurons is turned on as a compensatory reaction under degeneration of the nigral DA‐ergic neurons in Parkinson’s disease. In fact, the number of the striatal neurons expressing enzymes of DA synthesis in patients with Parkinson’s disease exceeded that of normal humans (Porritt et al., 2000). About 66, 000 neurons expressing either TH or DA transporter have been detected postmortem in the striatum and in the close basal ganglia like globus pallidus and internal capsule in humans with Parkinson’s disease (Porritt et al., 2000). It should be emphasized that this population is only twice as little compared to the whole population of DA‐ergic neurons in the substantia nigra of the same patients and is as large as the population of DA‐ergic neurons of the substantia nigra innervating the putamen in normal humans. Noteworthy, the implantation of only a slightly higher number of embryonic DA‐ergic neurons (80, 000) to patients with Parkinson’s disease is sufficient to induce the substantial but temporal improvement of their status (Ugrumov, 2001). Compensatory mechanisms in Parkinson’s disease. A delay for 25–30 years in the appearance of the initial symptoms of Parkinson’s disease after the onset of the nigral DA‐ergic neuron degeneration is apparently explained by turning on the compensatory processes which are mostly realized in the striatum (Bezard et al., 2003). In addition to the expression of the enzymes of DA synthesis in non‐DA‐ergic neurons, they include: (1) an increase of TH activity and DA synthesis in the surviving nigral DA‐ergic neurons (Agid et al., 1973; Mogi et al., 1988) despite of downregulation of the TH synthesis (Sherman and Moody, 1995); (2) the increased release of DA from the rest of axon terminals (Bernheimer et al., 1973; Zhang et al., 1988); (3) a decrease of the expression of the DA transporter and the DA uptake (Uhl et al.,

61

62

2

Brain neurons partly expressing monoaminergic phenotype

1994; Laihinen et al., 1995; Garris et al., 1997); (4) the enhanced sensibility of the target‐neurons according to upregulation of DA receptors (Bezard et al., 2003); (5) the DA diffusion from intact to denervated regions along the intercellular clefts that is a milestone of the hypothesis on ‘‘the passive stabilization’’ (Bergstrom and Garris, 2003), on one hand, and fits well in the hypothesis on ‘‘the volume or extrasynaptic transmission’’ (Zoli et al., 1998; Vizi, 2000), on the other. The earlier compensatory mechanisms result in maintaining of the normal concentration of DA in denervated striatum under the continuous degeneration of the nigral DA‐ergic neurons (Robinson and Whishaw, 1988; Garris et al., 1997; Bezard et al., 2003). Obviously the list of the compensatory mechanisms is not limited by those described earlier that makes it necessary to extend these studies. After the initial appearance of symptoms, the Parkinson’s disease is developed rapidly as a result of the failure of DA‐ergic system on one hand, and on the other hand by the exhaustion of compensatory resources. Thus, the functional failure of the nigrostriatal DA‐ergic system is temporarily compensated by turning on the mechanisms of the brain plasticity, including the expression of enzymes of DA synthesis in the striatal non‐DA‐ergic neurons which probably contribute to DA synthesis.

7

Regulation of the Partial Expression of MA‐ergic Phenotype by the Brain Neurons in Norm and Pathology

From the earlier data it follows that the frequency of monoenzymatic neurons varies in certain brain regions in ontogenesis and in adulthood under different functional conditions or in pathology. For instance, the number of monoenzymatic neurons decreases in the AN in ontogenesis and increases in adulthood under degeneration of DA‐ergic neurons (Ershov et al., 2002a, 2005). The latter phenomenon is also true for striatal monoenzymatic neurons in adulthood after degeneration of the nigral DA‐ergic neurons (Lopez‐ Real et al., 2003). Another example is represented by magnocellular vasopressinergic neurons of the supraoptic, paraventricular, and accessory nuclei which coexpress TH in norm under chronic functional stimulation (> Figure 2-6) (Abramova et al., 2002), chronic deficiency of vasopressin (Kiss and Mezey, 1986), as well as in pathology under certain metabolic disorders (Fetisov et al., 1997), and perinatal hypoxia (Panayotacopoulou et al., 1994).

7.1 Regulation of the Partial Expression of MA‐ergic Phenotype by Neural Afferents Although the regulation of the expression of the enzymes of DA synthesis in non‐MA‐ergic neurons remains uncertain, some data suggest that neural afferents or more precisely their neurotransmitters contribute to this control. In fact, the number of monoenzymatic neurons increases in certain brain regions, the supraoptic nucleus, the AN and the striatum, after their surgical deafferentation (Kiss and Mezey, 1986; Daikoku et al., 1986; Betarbet et al., 1997; Lopez‐Real et al., 2003) or, on the contrary, decreased in number along with their innervation in ontogenesis (Sorokin et al., unpublished). The earlier hypothesis has been first tested by using vasopressinergic neurons, one of the most promising cell models to solve this problem. Indeed, it has been shown till now that for these cells they may transiently coexpress TH during the osmotic stimulation. This suggests an existence of extracellular signals which can turn on the TH expression after the onset of osmotic stimulation and turn it off after normalization of the water‐mineral metabolism (Yagita et al., 1994). Indeed, it has been recently demonstrated that the TH expression in vasopressinergic neurons is inhibited by noradrenergic afferents and hence noradrenaline via adrenoreceptors (> Figure 2-34). This conclusion derived from the fact that the administration in vivo of the a1‐adrenoreceptor antagonist increased whereas the administration of the a1‐adrenoreceptor agonist decreased the concentrations of TH mRNA and protein in osmotically stimulated vasopressinergic neurons in young rats (Ugrumov, 2002; Abramova et al., unpublished). The catecholaminergic inhibitory control of the expression of the enzymes of MA synthesis in non‐MA‐ ergic neurons may be an attribute not only of vasopressinergic neurons but also of the neurons located

Brain neurons partly expressing monoaminergic phenotype

2

. Figure 2‐34 Schematic drawing of the inhibitory influence of noradrenergic afferents and noradrenaline on tyrosine hydroxylase expression in vasopressinergic neurons of supraoptic nucleus. dots, noradrenaline; asterisk, tyrosine hydroxylase; filled circles, vasopressin secretory granules; filled triangle, adrenoreceptor

outside the magnocellular nuclei. In this context, the non‐MA‐ergic neurons of the AN, striatum, and the olfactory bulbs appear to be under the DA‐ergic afferent inhibitory control. This suggestion is supported by the observations in rats of: (1) the increased number of monoenzymatic TH‐neurons in the AN (Ershov et al., 2005), striatum (Tashiro et al., 1989a, b), and the olfactory bulbs (Tashiro et al., 1990) under degeneration of their afferent DA‐ergic neurons located in the AN, substantia nigra, and ventral tegmental area, respectively; (2) the coexpression of TH in GABA‐ergic neurons after degeneration of their DA‐ergic afferents (Meredith et al., 1999); (3) the existence of numerous TH‐immunoreactive neurons in the striatum and the limbic system during prenatal and early postnatal periods in rats, before the establishment of their DA‐ergic synaptic innervation, and (4) the disappearance of TH‐immunoreactive neurons by puberty (Sorokin et al., unpublished), after completion of synaptogenesis. The inhibitory action on the TH expression if exists, is provided by DA rather than by L‐DOPA as L‐DOPA does not influence the expression of at least AADC in the striatal neurons (Lopez‐Real et al., 2003). The hypothesis about the catecholaminergic inhibitory control of the enzymes of MA synthesis in non‐ MA‐ergic neurons should be carefully tested in the future because of ambiguity of the interpretations of

63

64

2

Brain neurons partly expressing monoaminergic phenotype

some supporting arguments. In fact, the synchronization of the disappearance of the neurons expressing enzymes of DA synthesis and the establishment of the afferent catecholaminergic synaptic innervation in ontogenesis may be a result not only of the catecholaminergic inhibitory control of the enzymes expression in this local brain area but also of the neuron migration to other brain regions or their apoptosis. Furthermore, the following data also appear to contradict this idea: (1) the monoenzymatic neurons expressing TH are usually located in the denervated striatum in close vicinity of surviving DA‐ergic fibers (Porritt et al., 2000; Lopez‐Real et al., 2003); (2) DA promotes the stimulating influence of some neurotropic factors, e.g., ‘‘brain‐derived neurotrophic factor’’, on TH expression in the primary cell culture of the striatum (Du and Iacovitti, 1995).

7.2 Paracrine Regulation of the Partial Expression of MA‐ergic Phenotype by Diffusive Factors Neurotrophic or growth factors which are produced to a great extent by glial cells appear to be among the most efficient chemical signals stimulating the expression of the enzymes of DA synthesis in non‐MA‐ergic neurons. This suggestion has derived from the observation of the synchronization of the increased secretion of neurotrophic factors (Nakajima et al., 2001) with the expression of the enzymes of DA synthesis in the striatal non‐DA‐ergic neurons under degeneration of nigral DA‐ergic neurons. The intensity of both processes increased continuously until reaching maximum 2–3 weeks following the neurotoxin injection (Meredith et al., 1999; Nakajima et al., 2001). The neurotrophic factors were shown to stimulate the surviving of DA‐ergic neurons and the local reparative processes in the denervated striatum of adult animals, as well as in the striatum of ageing animals (Du and Iacovitti, 1995; Tomac et al., 1995). They occur to be of particular efficiency in the brain of primates. This is manifested by the multiple increase of the number of the neurons expressing TH and the DA transporter in the intact striatum of old monkeys or in the DA‐ergic denervated striatum in young monkeys after the intracerebral injections of the vector of the glia‐derived neurotrophic factor to both groups of the animals (Palfi et al., 2002). A number of data obtained in the in vivo and in vitro studies have definitely proven that the neurotrophic factors promoted the expression of the enzymes of DA synthesis in the striatal neurons. The fibroblast growth factor and the brain‐derived neurotrophic factor occurred to be particularly efficient in stimulation of the TH expression in non‐DA‐ergic neurons in the denervated striatum, and their action is mediated by DA. Although the glial‐derived neurotrophic factor and the ciliary neurotrophic factor provide similar action on the striatal neurons, they are less efficient (Du and Iacovitti, 1995). The administration of neurotrophic factors to the striatal tissue culture induced the appearance of the initial TH‐immunoreactive cells in 12 hours. Then, TH‐immunoreactive neurons increased rapidly in number reaching maximum 18 hours after the treatment that was followed by the gradual decrease in the neuron number for subsequent 4 days (Du and Iacovitti, 1995). The action of neurotrophic factors has been shown to be mediated via specific receptors. The glia‐derived neurotrophic factor provides its action through a multireceptor complex composed of a novel glycosylphosphatidylinositol‐anchored glia‐derived neurotrophic factor receptor‐a and the receptor tyrosine kinase product of the c‐ret proto‐oncogene (Durbec et al., 1996; Trupp et al., 1997). The expression of this receptor in the developing brain in rats is maximal in the early postnatal period that coincides with a high level of the TH expression (Sorokin et al., unpublished) and minimal in adulthood (Trupp et al., 1997). In addition to catecholamines (see earlier), vasopressin is also considered as an extracellular signal providing an inhibitory control of the TH expression in vasopressinergic neurons. This was indirectly confirmed by the decreased level of TH in the neurons of the magnocellular nuclei in the vasopressin deficient Brattleboro rats treated with exogenous vasopressin (Kiss and Mezey, 1986). A list of chemical signals controlling the expression of the enzymes of DA synthesis in non‐DA‐ergic neurons apparently is not limited to catecholamines, DA, vasopressin, and neurotrophic factors. By contrast with these extracellular signals, some signals may provide the opposite effects when acting on the neurons in different brain regions or in the same region but over different periods of ontogenesis. For instance, the serotonin deficiency provoked by p‐chlorophenylalanine, an inhibitor of serotonin synthesis, resulted in

Brain neurons partly expressing monoaminergic phenotype

2

the increased number of TH‐immunoreactive neurons in the dorsal motor vagal nucleus (Kitahama et al., 1987) and in the decrease of the TH content in the differentiating neurons of the AN that was shown both in vivo and in the primary cell culture (Melnikova et al., 2001). Noteworthy, serotonin as a morphogenetic diffusive factor provides a long‐lasting effect on differentiating TH‐neurons in the AN that is manifested by the maintaining of a relatively low level of TH in adult offspring treated with p‐chlorophenylalanine during a certain period of the intrauterine development (Melnikova et al., 2001). The only weak point of the earlier studies was the use of the TH mono‐immunolabeling as it is impossible to precise whether the monoenzymatic TH‐neurons or bienzymatic DA‐ergic neurons are the targets for serotonin. Only occasional studies have been somehow related to the regulation of the expression of AADC in monoenzymatic neurons. It has been demonstrated in rats that the AADC gene expression and AADC activity in monoenzymatic neurons of the SCN strongly depend on the circadian rhythms being maximal during daylight hours and minimal over nocturnal period. Although the regulation of AADC expression in these neurons by extracellular signals is quite probable, their nature remains uncertain (Ishida et al., 2002).

7.3 Hormonal Regulation of the Partial Expression of MA‐ergic Phenotype The regulation of the expression of the enzymes of DA synthesis in non‐MA‐ergic neurons is provided probably not only by the brain‐derived factors but also by hormones. However, no direct evidence of this suggestion is so far available. The most promising model to solve this issue in the future appears to be the AN containing a great portion of monoenzymatic TH‐neurons and AADC‐neurons (Ershov et al., 2002a, b), which synthesize DA in cooperation (Ugrumov et al., 2004), in addition to DA‐ergic bienzymatic neurons . It has been repeatedly demonstrated that the DA‐producing neurons of the AN contribute to the regulation of reproduction providing the inhibitory control of the pituitary prolactin secretion (McCann et al., 1984; Moore et al., 1985). Moreover, DA produced in the AN contributes to the regulation of the gonadotropin‐releasing hormone secretion by the neurons of the anterior forebrain. In turn, DA synthesis in the AN is under the feedback control of the pituitary (prolactin) and gonadal (estrogens, progesterone) reproductive hormones (Arbogast and Voogt, 1991). The hormonal regulation of the DA‐producing neurons in the AN is supported by the observation of receptors for prolactin (Arbogast and Voogt, 1997; Lerant and Freeman, 1998), estrogens (Jones and Naftolin, 1990; Hou et al., 2003;Mitchell et al., 2003), and progesterone (Warembourg et al., 1993; Dufourny et al., 2005) on TH‐expressing neurons, as well as by detection of the progesterone receptors on AADC‐containing neurons (Warembourg et al., 1993). The earlier data were considered as the proof of the receptor expressions in DA‐ergic neurons (Jones and Naftolin, 1990; Arbogast and Voogt, 1991, 1997; Lerant and Freeman, 1998; Hou et al., 2003) though the authors did not attempt to detect both enzymes of DA synthesis in these neurons by using double‐labeling technique. Therefore, it remains uncertain whether the hormone receptors are expressed in DA‐ergic and/or in monoenzymatic neurons. Up to the present, only occasional studies have provided indirect evidence of the expression of receptors for hormones of reproduction both in the DA‐ergic and monoenzymatic neurons. In fact, the number of monoenzymatic TH‐neurons and AADC‐neurons increased significantly under hyperprolactinemia provoked by 6‐OHDA‐induced degeneration of DA‐ergic neurons (Ershov et al., 2005). Apparently, this compensatory mechanism is responsible for normalization with time of the prolactin secretion (Fenske and Wuttke, 1976). The decrease of the density of monoenzymatic TH‐fibers in the median eminence synchronous to a change of the concentrations of prolactin and progesterone in plasma in the rats during postnatal period (Arbogast and Voogt, 1991; Ershov et al., 2002b) may be considered as another argument in favor of the expression of the receptors for reproduction hormones in monoenzymatic TH‐neurons. Furthermore, the data showing that the TH‐expressing neurons located in the dorsomedial area of the AN, mostly bienzymatic, and those located in the ventrolateral area of the AN, mostly monoenzymatic, are differently regulated by reproduction hormones (Lerant and Freeman, 1998) appear to be particularly promising for further strengthening of our knowledge about the hormonal regulation of the

65

66

2

Brain neurons partly expressing monoaminergic phenotype

partial expression of the MA‐ergic phenotype. Noteworthy, the sensitivity to reproduction hormones, e.g., prolactin is an attribute not only of DA‐producing neurons of the tuberoinfundibular system but also of those in other brain regions including the nigrostriatal system (Chen and Ramirez, 1989). Thus, the expression of the enzymes of DA synthesis in monoenzymatic non‐DA‐ergic neurons appears to be under the control of classical neurotransmitters, paracrine diffusive factors, and hormones.

Acknowledgments This study was supported by the grants of: the program of the Presidium of the Russian Academy of Sciences ‘‘Basic sciences for medicine’’, the program of the Department of Biological Sciences ‘‘Physiological mechanisms of the regulation of homeostasis in the systemic control of the animal behavior’’, RGNF 06‐06‐ 000‐10A, Scientific Schools‐6352.2006.4., RFBR 05-04-48829, RFBR-OFI 07-04-12211, PICS 07-04-92173.

References Abramova M, Calas A, Thibault J, Ugrumov M. 2000. Tyrosine hydroxylase in vasopressinergic axons of the pituitary posterior lobe of rats under salt‐loading as a manifestation of neurochemical plasticity. Neural Plast 7: 179-191. Abramova M, Marsais F, Calas A, Thibault J, Ugrumov M. 2002. Dynamical study of tyrosine hydroxylase expression and its correlation with vasopressin turnover in the magnocellular neurons of the supraoptico‐posthypophysial system under long‐term salt‐loading of adult rats. Brain Res 925: 67-75. Agid Y. 1991. Parkinson’s disease pathophysiology. Lancet 337: 1321-1323. Agid Y, Javoy F, Glowinski J. 1973. Hyperactivity of remaining dopaminergic neurons after partial destruction of the nigrostriatal dopaminergic system in the rat. Nat New Biol 245: 150-151. Annunziato L, Leblanc P, Kordon C, Weiner RI. 1980. Differences in the kinetics of dopamine uptake in synaptosome preparations of the median eminence relative to other dopaminergically innervated brain regions. Neuroendocrinology 31: 316-320. Arai R, Karasawa N, Geffard M, Nagatsu I. 1995. L‐DOPA is converted to dopamine in serotonergic fibers of the striatum of the rat: A double‐labeling immunofluorescence study. Neurosci Lett 11: 195-198. Arai R, Karasawa M, Nagatsu I. 1996. Aromatic L‐amino acid decarboxylase is present in serotonergic fibers of the striatum of the rat. A double‐labeling immunofluorescence study. Brain Res 706: 177-179. Arbogast LA, Voogt JL. 1991. Hyperprolactinemia increases and hypoprolactinemia decreases tyrosine hydroxylase messenger ribonucleic acid levels in the arcuate nuclei, but not the substantia nigra or zona incerta. Endocrinology 128: 997-1005.

Arbogast LA, Voogt JL. 1997. Prolactin (PRL) receptors are colocalized in dopaminergic neurons in fetal hypothalamic cell cultures: Effect of PRL on tyrosine hydroxylase activity. Endocrinology 138: 3016-3023. Balan IS, Ugrumov MV, Calas A, Mailly P, Krieger M, et al. 2000. Tyrosine hydroxylase‐ and/or aromatic L‐amino acid decarboxylase‐expressing neurons in the mediobasal hypothalamus of perinatal rats: Differentiation and sexual dimorphism. J Comp Neurol 425: 167-176. Battaglia AA, Beltramo M, Thibault J, Krieger M, Calas A. 1995. A confocal approach to the morphofunctional characterization of the transient tyrosine hydroxylase system in the rat suprachiasmatic nucleus. Brain Res 696: 7-14. Beaudet A, Descarries L. 1979. Radioautographic characterization of a serotonin‐accumulating nerve cell group in the adult rat hypothalamus. Brain Res 160: 231-243. Beltramo M, Calas A, Chernigovskaya N, Borisova N, Polenova O, et al. 1994. Postnatal development of the suprachiasmatic nucleus in the rat. Morphofunctional characteristics and time course of tyrosine hydroxylase immunopositive fibers. Neuroscience 63: 603-610. Berger B, Verney C, Gaspar P, Febvret A. 1985. Transient expression of tyrosine hydroxylase immunoreactivity in some neurons of the rat neocortex during postnatal development. Brain Res 355: 141-144. Bergstrom BP, Garris PA. 2003. ‘‘Passive stabilization’’ of striatal extracellular dopamine across the lesion spectrum encompassing the presymptomatic phase of Parkinson’s disease: A voltammetric study in the 6‐OHDA‐lesioned rat. J Neurochem 87: 1224-1236. Bernheimer H, Birkmayer W, Hornykiewcz O, Jellinger K. Seitelberger R. 1973. Brain dopamine and the syndromes of Parkinson and Huntington. J Neurol Sci 20: 415-455.

Brain neurons partly expressing monoaminergic phenotype Betarbet R, Greenamyre JT. 1999. Differential expression of glutamate receptors by the dopaminergic neurons of the primate striatum. Exp Neurol 159: 401-408. Betarbet R, Turner R, Chockkan V, DeLong MR, Allers KA, et al. 1997. Dopaminergic neurons intrinsic to the primate striatum. J Neurosci 17: 6761-6768. Bezard E, Gross ChE, Brotchie JM. 2003. Presymptomatic compensation in Parkinson’s disease is not dopamine‐ mediated. Trends Neurosci 26: 215-221. Bjo¨rklund A, Lindvall O. 1984. Dopamine‐containing system in the CNS. Handbook of Chemical Neuroanatomy, Vol. 2., Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 55-122. Bosler O, Calas A. 1982. Radioautographic investigation of monoaminergic neurons: An evaluation. Brain Res Bull 9: 151-169. Bosse R, Fumagalli F, Jaber M, Giros B, Gainetdinov RR, et al. 1997. Anterior pituitary hypoplasia and dwarfism in mice lacking the dopamine transporter. Neuron 19: 127-138. Chaillou E, Tramu G, Thibault J, Tillet Y. 1998. Presence of galanin in dopaminergic neurons of the sheep infundibular nucleus: A double staining immunohistochemical study. J Chem Neuroanat 15: 251-259. Chen JC, Ramirez VD. 1989. Effects of prolactin on tyrosine hydroxylase activity of central dopaminergic neurons of male rats. Eur J Pharmacol 166: 473-479. Cossette M, Parent A, Levesque D. 2004. Tyrosine hydroxylasepositive neurons intrinsic to the human striatum express the transcription factor Nurr1. Eur J Neurosci 20: 20892095. Cossette M, Lecomte F, Parent A. 2005a. Morphology and distribution of dopaminergic neurons intrinsic to the human striatum. J Chem Neuroanat 29: 1-11. Cossette M, Le´vesque D, Parent A. 2005b. Neurochemical characterization of dopaminergic neurons in human striatum. Parkinsonism Relat Disord 11: 277-286. Coyle JT, Henry D. 1973. Catecholamines in fetal and newborn rat brain. J Neurochemistry 21: 61-67. Dahlstro¨m A, Fuxe K. 1964. Evidence for the existence of monoamine‐containing neurons in the central nervous system. I. Demonstration of monoamines in the cell bodies of brainstem neurons. Acta Physiol Scand Suppl 232: 1-55. Daikoku S, Kawano H, Okamura I, Tokuzen M, Nagatsu I. 1986. Ontogenesis of immunoreactive tyrosine hydroxylase‐ containing neurons in rat hypothalamus. Brain Res 393: 85-98. Dairman W. 1972. Catecholamine concentrations and the activity of tyrosine hydroxylase after an increase in the concentration of tyrosine in rat tissues. Br J Pharmacol 44: 307-310.

2

Demarest KT, Moore KE. 1980. Accumulation of L‐DOPA in the median eminence: An index of tuberoinfundibular dopaminergic nerve activity. Endocrinology 106: 463-468. De Vitry F, Hamon M, Catelon J, Dubois M, Thibault J. 1986. Serotonin initiates and autoamplifies its own synthesis during mouse central nervous system development. Proc Natl Acad Sci USA 83: 8629-8633. Du X, Iacovitti L. 1995. Synergy between growth factors and transmitters required for catecholamine differentiation in brain neurons. J Neurosci 15: 5420-5427. Dubach M, Schmidt R, Kunkel D, Bowden DM, Martin R, et al. 1987. Primate neostriatal neurons containing tyrosine hydroxylase: Immunohistochemical evidence. Neurosci Lett 75: 205-210. Dufourny L, Caraty A, Clarke IJ, Robinson JE, Skinner DC. 2005. Progesterone‐receptive dopaminergic and neuropeptide Y neurons project from the arcuate nucleus to gonadotropin‐releasing hormone‐rich regions of the ovine preoptic area. Neuroendocrinology 82: 21-31. Durbec P, Marcos‐Gutierrez CV, Kilkenny C, Grigoriou M, Suvanto P, et al. 1996. GDNF signaling through the Ret receptor tyrosine kinase. Nature 381: 789-793. Ershov PV, Ugrumov MV, Calas A, Krieger M, Thibault J. 2002a. Differentiation of tyrosine hydroxylase‐expressing and/or aromatic L‐amino acid decarboxylase‐expressing neurons in the rat mediobasal hypothalamus: Quantitative double‐immunofluorescence study. J Comp Neurol 446: 114-122. Ershov PV, Ugrumov MV, Calas A, Krieger M, Thibault J. 2005. Degeneration of dopaminergic neurons triggers an expression of individual enzymes of dopamine synthesis in non‐dopaminergic neurons of the arcuate nucleus in adult rats. J Chem Neuroanat 30: 27-33. Ershov PV, Ugrumov MV, Calas A, Makarenko IG, Krieger M, et al. 2002b. Neurons possessing enzymes of dopamine synthesis in the mediobasal hypothalamus of rats. Topographic relations and axonal projections to the median eminence in ontogenesis. J Chem Neuroanat 24: 95-107. Everitt B, Meister B, Ho¨kfelt T, Melander T, Terenins L, et al. 1986. The hypothalamic arcuate nucleus‐median eminence complex: Immunocytochemistry of transmitters, peptides and DARPP‐32 with special reference to coexistence in dopamine neurons. Brain Res Rev 11: 97-155. Fenske M, Wuttke W. 1976. Effects of intraventricular 6‐hydroxydopamine injections on serum prolactin and LH levels: Absence of stress‐induced pituitary prolactin release. Brain Res 104: 63-70. Ferna´ndez E, Torrents D, Zorzano A, Palacı´n M, Chillaro´n J. 2005. Identification and functional characterization of a novel low affinity aromatic‐preferring amino acid transporter (arpAT). One of the few proteins silenced during primate evolution. J Biol Chem 280: 19364-19372.

67

68

2

Brain neurons partly expressing monoaminergic phenotype

Fetisov S, Marsais F, Nicolandis S, Calas A. 1997. Expression of tyrosine hydroxylase in magnocellular hypothalamic neurons of obese (fa/fa) and lean heterozygous (Fa/fa) Zucker rats. Mol Brain Res 50: 314-318. Fisher A, Biggs CS, Eradiri O, Starr MS. 2000. Dual effects of L‐3,4‐Dihydroxyphenylalanine on aromatic L‐amino acid decarboxylase, dopamine release and motor stimulation in the reserpine‐treated rat: Evidence that behaviour is dopamine dependent. Neuroscience 95: 97-111. Frankfurt H, Azmitia E. 1983. The effect of intracerebral injections of 5,7‐dihydroxytryptamine and 6‐hydroxydopamine on the serotonin‐immunoreactive cell bodies and fibers in the adult rat hypothalamus. Brain Res 261: 91-99. Frankfurt H, Lauder J, Azmitia E. 1981. The immunocytochemical localization of serotoninergic neurons in the rat hypothalamus. Neurosci Lett 24: 227-232. Fuxe K, Ungerstedt U. 1968. Histochemical studies on the effect of (positive)‐amphetamine, drugs of the imipramine group and tryptamine on central catecholamine and 5‐hydroxytryptamine neurons after intraventricular injection of catecholamines and 5‐hydroxytryptamine. Eur J Pharmacol 4: 135-144. Garris PA, Walker QD, Wightman RM. 1997. Dopamine release and uptake rates both decrease in the partially denervated striatum in proportion to the loss of dopamine terminals. Brain Res 753: 225-234. Gaspar P, Berger B, Febvret A, Vigny A, Krieger‐Poulet M, et al. 1987. Tyrosine hydroxylase‐immunoreactive neurons in the human cerebral cortex: A novel catecholaminergic group? Neurosci Lett 80: 257-262. Gaspar P, Cases O, Maroteaux L. 2003. The developmental role of serotonin: News from mouse molecular genetics. Nat Rev Neurosci 4: 1002-1012. Goshima Y, Kubo T, Misu Y. 1988. Transmitter‐like release of endogenous 3,4‐dihydroxyphenylalanine from rat striatal slices. J Neurochem 50: 1725-1730. Goshima Y, Nakamura S, Ohno K, Misu Y. 1991. Picomolar concentrations of L‐dopa stereoselectively potentiate activities of presynaptic b‐adrenoreceptors to facilitate the release of endogenous noradrenaline from rat hypothalamic slices. Neurosci Lett 129: 214-216. Hardebo JE, Emson PC, Falck B, Owman Ch, Rosengren E. 1980. Enzymes related to monoamine transmitter metabolism in brain microvessels. J Neurochem 35: 1388-1393. Hoffman BJ, Hansson SR, Mezey E´, Palkovits M. 1998. Localization and dynamic regulation of biogenic amine transporters in the mammalian central nervous system. Front Neuroendocrinol 19: 187-231. Ho¨kfelt T, Johansson O, Goldstein M. 1984. Central catecholamine neurons as revealed by immunocytochemistry with special reference to adrenaline neurons. Handbook of

Chemical Neuroanatomy, Vol. 2., Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 157-276. Hou Y, Yang SP, Voogt JL. 2003. Changes in estrogen receptor‐a expression in hypothalamic dopaminergic neurons during proestrous prolactin surge. Endocrine 20: 131-138. Huot P, Le´vesque M, Parent A. 2007. The fate of striatal dopaminergic neurons in Parkinson’s disease and Huntington’s chorea. Brain 130: 222-232. Huot P, Parent A. 2007. Dopaminergic neurons intrinsic to the striatum. J Neurochem 101:1441-1447. Ibata Y, Fukui K, Obata HL, Tanaka M, Hisa Y, et al. 1982. Postnatal ontogeny of catecholamine and somatostatin neuron systems in the median eminence of the rat as revealed by a colocalization technique. Brain Res Bull 9: 407-415. Ikemoto K, Kitahama K, Jouvet A, Arai R, Nishimura A, et al. 1997. Demonstration of L‐dopa decarboxylating neurons specific to human striatum. Neurosci Lett 232: 111-114. Ikemoto K, Kitahama K, Nishimura A, Jouvet A, Nishi K, et al. 1999. Tyrosine hydroxylase and aromatic L‐amino acid decarboxylase do not coexist in neurons in the human anterior cingulate cortex. Neurosci Lett 269: 37-40. Ikemoto K, Nagatsu I, Kitahama K, Jouvet A, Nishimura A, et al. 1998a. A dopamine‐synthesizing cell group demonstrated in the human basal forebrain by dual labeling immunohistochemical technique of tyrosine hydroxylase and aromatic L‐amino acid decarboxylase. Neurosci Lett 243: 129-132. Ikemoto K, Nagatsu I, Nishimura A, Nishi K, Arai R. 1998b. Do all of human midbrain tyrosine hydroxylase neurons synthesize dopamine? Brain Res 805: 255-258. Ikemoto K, Nishimura A, Oda T, Nagatsu I, Nishi K. 2003. Number of striatal D‐neurons is reduced in autopsy brains of schizophrenics. Legal Medicine 5: 221-224. Ishida Y, Yokoyama C, Inatomi T, Yagita K, Dong X, et al. 2002. Circadian rhythm of aromatic L‐amino acid decarboxylase in the rat suprachiasmatic nucleus: Gene expression and decarboxylating activity in clock oscillating cells. Genes Cells 7: 447-459. Izvolskaia M, Duittoz AH, Ugrumov M, Tillet Y. 2006. Tyrosine hydroxylase expression in the olfactory/respiratory epithelium in early sheep fetuses (Ovis aries). Brain Res 1083: 29-38. Jaeger CB, Albert VR, Joh TH, Reis DJ. 1983. Aromatic L‐amino acid decarboxylase in the rat brain: Coexistence with vasopressin in small neurons of the suprachiasmatic nucleus. Brain Res 276: 362-366. Jaeger CB, Ruggiero DA, Albert VR, Park DH, Joh TH, et al. 1984. Immunocytochemical localization of aromatic‐ L‐amino acid decarboxylase. Handbook of Chemical

Brain neurons partly expressing monoaminergic phenotype Neuroanatomy, Vol. 2., Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp 387-408. Jaeger CB, Teitelman G. 1992. Immunocytochemical distribution of aromatic L‐amino acid decarboxylase (AADC) in rat embryos. Handbook of Chemical Neuroanatomy, Vol. 10: Ontogeny of Transmitters and Peptides in the CNS. Bjo¨rklund A, Ho¨kfelt T, Tohyama M, editors. Amsterdam: Elsevier. Jones EE, Naftolin F. 1990. Estrogen effects on the tuberoinfundibular dopaminergic system in the female rat brain. Brain Res 510: 84-91. Jonsson G. 1983. Chemical lesions techniques: Monoamine neurotoxins. Handbook of Chemical Neuroanatomy, Vol. 1: Methods in Chemical Neuroanatomy. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; Pages 463-507. Juorio AV, Li X‐M, Walz W, Paterson IA. 1993. Decarboxylation of L‐dopa by cultured mouse astrocytes. Brain Res 626: 306-309. Kannari K, Huo Shen H, Arai A, Tomiyama M, Baba M. 2006. Reuptake of L‐DOPA‐derived extracellular dopamine in the striatum with dopaminergic denervation via serotonin transporters. Neurosci Lett 402: 62-65. Karasawa N, Arai R, Isomura G, Nagatsu T, Nagatsu I. 1995. Chemical features of monoaminergic and non‐ monoaminergic neurons in the brain of laboratory shrew (Suncus murinus) are changed by systemic administration of monoamine precursors. Neurosci Res 24: 67-74. Karasawa N, Arai R, Isomura G, Nagatsu T, Nagatsu I. 1997. Coexistence of tyrosine hydroxylase and serotonin in the raphe nucleus of the laboratory shrew (Suncus murinus) during postnatal life. Brain Res Dev Brain Res 99: 121-125. Karasawa N, Arai R, Isomura G, Yamada K, Sakai K, et al. 1994. Phenotypic changes of AADC‐only immunopositive premamillary neurons in the brain of laboratory shrew Suncus murinus by systemic administration of monoamine precursors. Neurosci Lett 179: 65-70. Kiss JZ, Mezey E. 1986. Tyrosine hydroxylase in magnocellular neurosecretory neurons. Response to physiological manipulations. Neuroendocrinology 43: 519-525. Kitahama K, Be´rod A, Denoyer M, Jouvet M. 1987. Visualization of tyrosine hydroxylase‐immunoreactive neurons in the cat dorsal motor vagal cells after treatment with parachlorophenylalanine. Neurosci Lett 77: 155-160. Kitahama K, Ikemoto K, Jouvet A, Nagatsu I, Sakamoto N, et al. 1998. Aromatic L‐amino acid decarboxylase‐ and tyrosine hydroxylase‐immunohistochemistry in the adult human hypothalamus. J Chem Neuroanat 16: 43-55. Klein DC, Moore RY, Reppert SM. 1991. Suprachiasmatic Nucleus. The Mind’s Clock. New York: Oxford University Press.

2

Komori K, Fujii T, Nagatsu I. 1991. Do some tyrosine hydroxylase‐immunoreactive neurons in the human ventrolateral arcuate nucleus and globus pallidus produce only L‐dopa? Neurosci Lett 133: 203-206. Laihinen AO, Rinne JO, Na˚gren KA˚, Lehikoinen PK, Oikonen VJ, et al. 1995. PET studies on brain monoamine transporters with carbon‐11‐b‐CIT in Parkinson’s disease. J Nucl Med 36: 1263-1267. Lauder J M. 1993. Neurotransmitters as growth regulatory signals: Role of receptors and second messengers. Trends Neurosci 16: 233-240. Lebrand C, Cases O, Wehrle’ R, Blakely RD, Edwards RH, et al. 1998. Transient developmental expression of monoamine transporters in the rodent forebrain. J Comp Neurol 401: 506-524. Lerant A, Freeman ME. 1998. Ovarian steroids differentially regulate the expression of PRL‐R in neuroendocrine dopaminergic neuron populations: A double label confocal microscopic study. Brain Res 802: 141-154. Lopez A, Mun˜oz A, Guerra MJ, Labandeira‐Garcia JL. 2001. Mechanisms of the effects of exogenous levodopa on the dopamine‐denervated striatum. Neuroscience 103: 639-651. Lopez‐Real A, Rodriguez‐Pallares J, Guerra M.J, Labandeira‐ Garcia JL. 2003. Localization and functional significance of striatal neurons immunoreactive to aromatic L‐amino acid decarboxylase or tyrosine hydroxylase in rat Parkinsonian models. Brain Res 969: 135-146. Lorang D, Amara SG, Simerly RB. 1994. Cell‐type‐specific expression of catecholamine transporters in the rat brain. J Neurosci 14: 4903-4914. Maeda T, Nagata K, Yoshida Y, Kannan K. 2005. Serotonergic hyperinnervation into the dopaminergic denervated striatum compensates for dopamine conversion from exogenously administered L‐DOPA. Brain Res 1046: 230-233. Marsais F, Parmentier C, Terao E, Taxi J, Calas A. 2002. Expression of tyrosine hydroxylase and vasopressin in magnocellular neurons of salt‐loaded aged rats. Microsc Res Tech 56: 81-91. Maurer‐Spurej E. 2005. Serotonin reuptake inhibitors and cardiovascular diseases: A platelet connection. Cell Mol Life Sci 62: 159-170. Max SR, Bossio A, Iacovitti L. 1996. Co‐expression of tyrosine hydroxylase and glutamic acid decarboxylase in dopamine differentiation factor‐treated striatal neurons in culture. Brain Res Dev Brain Res 91: 140-142. McCann SM, Lumpkin MD, Mizunuma H, Khorram O, Ottlecz J, et al. 1984. Peptidergic and dopaminergic control of prolactin release. Trends Neurosci 7: 127-131. Meister B, Ho¨kfelt T, Steinbusch HWM, Skagerberg G, Lindvall O, et al. 1988. Do tyrosine hydroxylase‐immunoreactive neurons in the ventro‐lateral arcuate nucleus

69

70

2

Brain neurons partly expressing monoaminergic phenotype

produce dopamine or only L‐DOPA? J Chem Neuroanat 1: 59-64. Melnikova V, Orosco M, Calas A, Sapronova A, Gainetdinov R, et al. 1999. Dopamine turnover in the mediobasal hypothalamus in rat fetuses. Neuroscience 89: 235-241. Melnikova V, Orosco M, Rouch C, Calas A, Nicolaidis S, et al. 1998. Prolactin secretion and its dopamine inhibitory control in rat fetuses. Eur J Endocrinol 139: 337-342. Melnikova VI, Sapronova AY, Lavrentieva AV, Proshlyakova EV, Voronova SN, et al. 2006. Brain is on of the sources of L‐3,4‐dihydroxyphenylalanine in general circulation of fetal and neonatal rats (Russia). J Evol Physiol Biochem 42: 19-25. Melnikova VI, Ugrumov MV, Proshlyakova EV, Calas A, Thibault J. 2001. Tyrosine hydroxylase expression in differentiating neurons of the rat arcuate nucleus: Stimulatory influence of serotonin afferents. Neural Plast 8: 271-284. Meredith GE, Farrell T, Kellaghan P, Tan Y, Zahm DS, et al. 1999. Immunocytochemical characterization of catecholaminergic neurons in the rat striatum following dopamine‐ depleting lesions. Eur J Neurosci 11: 3585-3596. Mirochnik VV, Makarenko IG, Ugryumov MV. 2002. The source of transitory innervation of suprachiasmatic nucleus by tyrosine hydroxylase‐immunoreactive fibers during postnatal period in rats (Russia). Ontogenez 33: 182-186. Misu Y, Kitahama K, Goshima Y. 2003. L‐3,4‐Dihydroxyphenylalanine as a neurotransmitter candidate in the central nervous system. Pharmacol Ther 97: 117-137. Mitchell V, Loyens A, Spergel DJ, Flactif M, Poulain PM, et al. 2003. A confocal microscopic study of gonadotropin‐ releasing hormone (GnRH) neuron inputs to dopaminergic neurons containing estrogen receptor alpha in the arcuate nucleus of GnRH‐green fluorescent protein transgenic mice. Neuroendocrinology 77: 198-207. Mogi M, Harada M, Kiuchi K, Kojima K, Kondo T, et al. 1988. Homospecific activity (activity per enzyme protein) of tyrosine hydroxylase increases in Parkinson’s brain. J Neural Transm 72: 77-81. Mons N, Tison F, Geffard M. 1989. Identification of L‐dopa‐ dopamine and L‐dopa cell bodies in the rat mesencephalic dopaminergic cell systems. Synapse 4: 99-105. Montange M, Calas A. 1988. Serotonin and endocrinology – the pituitary. Neural Serotonin. Osborne NN, Hamo M, editors. Chichester: John Wiley & Sons; pp. 271-303. Moore KE, Riegle GD, Demarest KT. 1985. Regulation of tuberoinfundibular dopaminergic neurons: Prolactin and inhibitory neuronal influences. Catecholamines as Hormone Regulators. Raven Press; New York: pp. 31-48. Mura A, Jackson D, Manley MS, Young SJ, Groves PM. 1995. Aromatic L‐amino acid decarboxylase immunoreactive cells

in rat striatum: A possible site for the conversion of exogenous L‐DOPA to dopamine. Brain Res 704: 51-60. Mura A, Linder JC, Young SJ, Groves PM. 2000. Striatal cells containing aromatic L‐amino acid decarboxylase: An immunohistochemical comparison with other classes of striatal neurons. Neuroscience 98: 501-511. Nagatsu T. 1995. Tyrosine hydroxylase: Human isoforms, structure and regulation in physiology and pathology. Essays Biochem 30: 15-35. Nagatsu I, Komori K, Takeuchi T, Sakai M, Yamada K, et al. 1990. Transient tyrosine hydroxylase‐immunoreactive neurons in the region of the anterior olfactory nucleus of pre‐ and postnatal mice do not contain dopamine. Brain Res 511: 55-62. Nagatsu I, Sakai M, Takeuchi T, Arai R, Karasawa N, et al. 1997. Tyrosine hydroxylase (TH)‐only‐immunoreactive non‐catecholaminergic neurons in the brain of wild mice or the human TH transgenic mice do not contain GTP cyclohydrolase I. Neurosci Lett 228: 55-57. Nagatsu I, Takeuchi T, Sakai M, Karasawa N, Yamawaki Y, et al. 1996. Transient appearance of tyrosine hydroxylase‐ immunoreactive non‐catecholaminergic neurons in the medial geniculate nucleus of postnatal mice. Neurosci Lett 211: 183-186. Nakahara T, Yamamoto T, Endo K, Kayama H. 2001. Neuronal ectopic expression of tyrosine hydroxylase in the mouse striatum by combind administration of 1‐methyl‐4‐phenyl‐ 1,2,3,6‐tetrahydropyridine and 3‐nitropropionic acid. Neuroscience 108: 601-610. Nakajima K, Hida H, Shimano Y, Fujimoto I, Hashitani T, et al. 2001. GDNF is a major component of trophic activity in DA‐depleted striatum for survival and neurite extension of DAergic neurons. Brain Res 916: 76-84. Nakamura K, Ahmed M, Barr E, Leiden JM, Kang UJ. 2000. The localization and functional contribution of striatal aromatic L‐amino acid decarboxylase to L‐3,4‐dihydroxyphenylalanine decarboxylation in rodent parkinsonian models. Cell Transplant 9: 567-576. Nakamura S, Goshima Y, Yue JL, Misu Y. 1992. Transmitter‐ like basal and K(þ)‐evoked release of 3,4‐dihydroxyphenylalanine from the striatum in conscious rats studied by microdialysis. J Neurochem 58: 270-275. Novak CM, Nunez AA. 1998. Tyrosine hydroxylase‐ and/or aromatic L‐amino acid decarboxylase‐containing cells in the suprachiasmatic nucleus of the Syeian hamster (Mesocricetus auratus). J Chem Neuroanat 14: 87-94. Okamura H, Kitahama K, Mons N, Ibata Y, Jouver M, et al. 1988a. L‐DOPA‐immunoreactive neurons in the rat hypothalamic tuberal region. Neurosci Lett 95: 42-46. Okamura H, Kitahama K, Nagatsu I, Geffard M. 1988b. Comparative topography of dopamine‐ and tyrosine

Brain neurons partly expressing monoaminergic phenotype hydroxylase‐immunoreactive neurons in the rat arcuate nucleus. Neurosci Lett 95: 347-353. Okamura H, Kitahama K, Raynaud B, Nagatsu I, Borri‐ Volttatorni C, et al. 1988c. Aromatic L‐amino acid decarboxylase (AADC)‐immunoreactive cells in the tuberal region of the rat hypothalamus. Biomed Res 9: 261-267. Okamura H, Murakami S, Chihara K, Nagatsu I, Ibata Y. 1985. Coexistence of growth hormone releasing factor‐like and tyrosine hydroxylase‐like immunoreactivities in neurons of the rat arcuate nucleus. Neuroendocrinology 41: 177-179. Palfi S, Leventhal L, Chu Y, Ma SY, Emborg M, et al. 2002. Lentivirally delivered glial cell line‐derived neurotrophic factor increases the number of striatal dopaminergic neurons in primate models of nigrostriatal degeneration. J Neurosci 22: 4942-4954. Panayotacopoulou MT, Raadsheer FC, Swaab DF. 1994. Colocalization of tyrosine hydroxylase with oxytocin or vasopressin in neurons of the human paraventricular and supraoptic nucleus. Brain Res Dev Brain Res 83: 59-66. Porritt MJ, Batchelor PE, Hughes AJ, Kalnins R, Donnan GA, et al. 2000. New dopaminergic neurons in Parkinson’s disease striatum. Lancet 356: 44-45. Richards KS, Simon DJ, Pulver SR, Beltz BS, Marder E. 2003. Serotonin in the developing stomatogastric system of the lobster, Homarus americanus. J Neurobiol 54: 380-392. Robinson TE, Whishaw IQ. 1988. Normalization of extracellular dopamine in striatum following recovery from a partial unilateral 6‐OHDA lesion of the substantia nigra: A microdialysis study in freely moving rats. Brain Res 450: 209-244. Sakai M, Fujii T, Karasawa N, Arai R, Nagatsu I. 1995. Enhanced expression of tyrosine hydroxylase and aromatic L‐amino acid decarboxylase in cerebellar Purkinje cells of mouse after hyperosmotic stimuli. Neurosci Lett 194: 142-144. Saucedo‐Cardenas O, Quintana‐Hau JD, Le WD, Smidt MP, Cox JJ, et al. 1998. Nurr1 is essential for the induction of the dopaminergic phenotype and the survival of ventral mesencephalic late dopaminergic precursor neurons. Proc Natl Acad Sci USA 95: 4013-4018. Schneider JS, Rothblat DS, DiStefano L. 1994. Volume transmission of dopamine over large distances may contribute to recovery from experimental Parkinsonism. Brain Res 643: 86-91. Serri O, Chik CL, Ur E, Ezzat S. 2003. Diagnosis and management of hyperprolactinemia. CMAJ 169: 575-581. Sherman TG, Moody CA. 1995. Alterations in tyrosine hydroxylase expression following partial lesions of the nigrostriatal bundle. Brain Res Mol Brain Res 29: 285-296.

2

Skagerberg G, Meister B, Ho¨kfelt T, Lindvall O, Goldstein M, et al. 1988. Studies on dopamine‐, tyrosine hydroxylase‐ and aromatic L‐amino acid decarboxylase‐containing cells in the rat diencephalon: Comparison between formaldehyde‐induced histofluorescence and immunofluorescence. Neuroscience 24: 605-620. Smith GC, Helme RD. 1974. Ultrastructural and fluorescence histochemical studies on the effects of 6‐hydroxydopamine on the rat median eminence. Cell Tissue Res 152: 493-512. Squire LR, Bloom FE, McConnell SK, Roberts JL, Spitzer NC, et al. 2003. Fundamental Neuroscience. Amsterdam: Academic Press. Steinbusch HWM. 1984. Serotonin‐immunoreactive neurons and their projections. Handbook of Chemical Neuroanatomy, Vol. 3., Classical CNS Transmitters and Transmitter receptors in the CNS. II. Bjo¨rklund A, Ho¨kfelt T, Kuhar MJ, editors. Amsterdam: Elsevier; pp. 68-125. Sugaya Y, Sasaski Y, Goshima Y, Kitahama K, Kusakabe T, et al. 2001. Autoradiographic studies using L‐(14C)DOPA and 3 þ L‐( H)DOPA reveal regional Na ‐dependent uptake of the neurotransmitter candidate L‐DOPA in the CNS. Neuroscience 104: 1-14. Tanaka H, Kannari K, Maeda T, Tomiyama M, Suda T, et al. 1999. Role of serotonergic neurons in L‐DOPA‐derived extracellular dopamine in the striatum of 6‐OHDA‐ lesioned rats. Neuroreport 1: 631-634. Tande´ D, Ho¨glinger G, Debeir T, Freundlieb N, Hirsch EC, et al. 2006. New striatal dopamine neurons in MPTP‐ treated macaques result from a phenotypic shift and not neurogenesis. Brain 129: 1194-2000. Tashiro Y, Kaneko T, Nagatsu I, Kikuchi H, Mizuno N. 1990. Increase of tyrosine hydroxylase‐like immunoreactive neurons in the nucleus accumbens and the olfactory bulb in the rat with the lesion in the ventral tegmental area of the midbrain. Brain Res 531: 159-166. Tashiro Y, Kaneko T, Sugimoto T, Nagatsu I, Kikuchi H, et al. 1989a. Striatal neurons with aromatic L‐amino acid decarboxylase‐like immunoreactivity in the rat. Neurosci Lett 100: 29-34. Tashiro Y, Sugimoto T, Hattori T, Uemura Y, Nagatsu I, et al. 1989b. Tyrosine hydroxylase‐like immunoreactive neurons in the striatum of the rat. Neurosci Lett 97: 6-10. Tinner B, Fuxe K, Ko¨hler C, Hersh L, Andersson K, et al. 1989. Evidence for the existence of a population of arcuate neurons costoring choline acetyltransferase and tyrosine hydroxylase immunoreactivities in the male rat. Neurosci Lett 99: 44-49. Tomac A, Lindqvist E, Lin LF, Ogren SO, Young D, et al. 1995. Protection and repair of the nigrostriatal dopaminergic system by GDNF in vivo. Nature 373: 335-339.

71

72

2

Brain neurons partly expressing monoaminergic phenotype

Trupp M, Belluardo N, Funaksohi H, Ibanez CF. 1997. Complementary and overlapping expression of glial cell line‐derived neurotrophic factor (GDNF), c‐ret proto‐ oncogene, and GDNF receptor‐a indicates multiple mechanisms of trophic actions in the adult rat CNS. J Neurosci 17: 3554-3567. Ugrumov MV. 1997. Hypothalamic monoaminergic systems in ontogenesis: Development and functional significance. Int J Dev Biol 41: 809-816. Ugrumov MV. 2001. Neurotransplantation in treatment of Parkinson’s disease. Basal Ganglia and Thalamus in Health and Movement Disorders. Kultas‐Ilinsky K, Ilinsky IA, editors. New York: Kluwer Academic Publishers/Plenum; pp. 349-363. Ugrumov MV. 2002. Magnocellular vasopressin system in ontogenesis: Development and regulation. Microsc Res Tech 56: 164-171. Ugrumov MV, Ivanova IP, Mitskevich MS, Liposits Z, Setalo, et al. 1985a. Axovascular relationships in developing median eminence of perinatal rats with special reference to luteinizing hormone‐releasing hormone projections. Neuroscience 16: 897-906. Ugrumov M, Melnikova V, Ershov P, Balan I, Calas A. 2002. Tyrosine hydroxylase‐ and/or aromatic L‐amino acid decarboxylase‐expressing neurons in the rat arcuate nucleus: Ontogenesis and functional significance. Psychoneuroendocrinology 7: 533-548. Ugrumov MV, Melnikova VI, Lavrentyeva AV, Kudrin VS, Rayevsky KS. 2004. Dopamine synthesis by non‐dopaminergic neurons expressing individual complementary enzymes of the dopamine synthetic pathway in the arcuate nucleus of fetal rats. Neuroscience 124: 629-635. Ugrumov MV, Popov AP, Vladimirov SV, Kasmambetova S, Thibault J. 1994. Development of the suprachiasmatic nucleus in rats during ontogenesis: Tyrosine hydroxylase immunopositive cell bodies and fibers. Neuroscience 58: 151-160. Ugrumov MV, Taxi J, Mitskevich MS, Arluison M, Tramu G. 1985b. Immunocytochemical and radioautographic study of serotonin projections to cerebral ventricles of perinatal rats. Brain Res 350: 225-230. Ugrumov MV, Taxi J, Mitskevich MS, Tramu G. 1986. Development of the hypothalamic serotoninergic system during ontogenesis in rats. Immunocytochemical and radioautographic study. Brain Res 395: 75-84. Ugrumov MV, Taxi J, Steinbusch HWM, Tramu G, Mitskevich MS. 1989a. On the distribution and morpho‐functional characteristics of 5‐HT‐immunoreactive cells in the hypothalamus of fetuses and neonatal rats. Brain Res Dev Brain Res 46: 233-241

Ugrumov M, Taxi J, Tixier‐Vidal A, Thibault J, Mitskevich MS. 1989b. Ontogenesis of tyrosine hydroxylase‐immunopositive structures in the rat hypothalamus. An atlas of neuronal cell bodies. Neuroscience 29: 135-156. Ugrumov M, Taxi J, Tixier‐Vidal A, Thibault J, Mitskevich MS. 1989c. Ontogenesis of tyrosine hydroxylase‐immunopositive structures in the rat hypothalamus. Fiber pathways and terminal fields. Neuroscience 29: 157-166. Uhl GR, Walther D, Mash D, Faucheux B, Javoy‐Agid F. 1994. Dopamine transporter messenger RNA in Parkinson’s disease and control substantia nigra neurons. Ann Neurol 35: 494-498. Van den Pol AN, Herbst RS, Powell JF. 1984. Tyrosine hydroxylase‐immunoreactive neurons of the hypothalamus: A light and electron microscopic study. Neuroscience 13: 1117-1156. Verney C, Gaspar P, Febvret A, Berger B. 1988. Transient tyrosine hydroxylase‐like immunoreactive neurons contain somatostatin and substance P in the developing amygdala and bed nucleus of the stria terminalis of the rat. Brain Res 470: 45-58. Verney C, El Amraoui A, Zecevic N. 1996. Comigration of tyrosine hydroxylase and gonadotropin‐releasing hormone‐ immunoreactive neurons in the nasal area of human embryos. Brain Res Dev Brain Res 97: 251-259. Verney C, Lebrand C, Gaspar P. 2002. Changing distribution of monoaminergic markers in the developing human cerebral cortex with special emphasis on the serotonin transporter. Anat Rec 267: 87-93. Vincent SR, Hope BT. 1990. Tyrosine hydroxylase containing neurons lacking aromatic amino acid decarboxylase in the hamster brain. J Comp Neurol 295: 290-298. Vizi ES. 2000. Role of high‐affinity receptors and membrane transporters in nonsynaptic communication and drug action in the central nervous system. Pharmacol Rev 52: 63-89. Warembourg M, Varoqueaux F, Poulain P. 1993. Le re´cepteur de la progeste´rone dans les neurones hypothalamiques du cobayae: Colocalisation avec la TH et projections des neurones vers l’e´minence me´deiane. Ann Endocrinol 54: 34 Weihe E, Depboylu C, Schu¨tz B, Scha¨fer MKH, Eiden LE. 2006. Three types of tyrosine hydroxylase‐positive CNS neurons distinguished by dopa decarboxylase and VMAT2 co‐expression. Cell Mol Neurobiol 26: 659-678. Weihe E, Eiden LE. 2000. Chemical neuroanatomy of the vesicular amine transporters. FASEB J 14: 2435-2449. Weiner RI, Findell PR, Kordon C. 1988. Role of classic and peptide neuromediators in the neuroendocrine regulation of LH and prolactin. The Physiology of Reproduction. Knobil E, Neill J, et al. editors. New York: Raven Press; pp. 1235-1281.

Brain neurons partly expressing monoaminergic phenotype Yagita K, Okamura H, Ibata Y. 1994. Rehydration process from salt‐loading: Recovery of vasopressin and its coexisting galanin, dynorphin and tyroisine hydroxylase immunoreactivities in the supraoptic and paraventricular nuclei. Brain Res 667: 13-23. Zhang WQ, Tilson HA, Nanry KP, Hudson PM, Hong JS, et al. 1988. Increased dopamine release from striata of rats after unilateral nigrostriatal bundle damage. Brain Res 461: 335-342.

2

Zoli M, Agnati LF, Tinner B, Steinbusch H, Fuxe K. 1993. Distribution of dopamine‐immunoreactive neurons and their relationship to transmitter and hypothalamic hormone‐immunoreactive neuronal systems in the rat mediobasal hypothalamus. A morphometric and microdensitometric analysis. J Chem Neuroanat 6: 293-310. Zoli M, Torri C, Ferrari R, Jansson A, Zini I, et al. 1998. The emergence of the volume transmission concept. Brain Res Brain Res Rev 26: 136-147.

73

3

In Vivo Imaging of Neurotransmitter Systems with PET

B. Gulya´s . C. Halldin . B. Mazie`re

1

Introduction: Neurotransmitter and Neuroreceptor Systems and In Vivo Neuroimaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76

2

Positron Emission Tomography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77

3

Labeling Tracers and Ligands with PET Bioisotopes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79

4

Preliminary Steps in the Development of Radioligands for Human CNS Receptors . . . . . . . . . . . . 82

5

Measuring Radioligand Effects in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83

6

Modeling Ligand Effects in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85

7 Two Approaches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 7.1 Direct Approach: Radiolabeling of Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 7.2 Indirect Approach: Using Radiolabeled Ligands and Drug Candidates . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 8 8.1 8.2 8.3 8.4 8.5 8.6 8.7 8.8 8.9

Radioligands for Mapping Neurotransmitter Systems: Some Examples . . . . . . . . . . . . . . . . . . . . . . . . . 87 Dopamine Receptor and Transporter Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89 Serotoninergic Neurotransmission Radioligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 Norepinephrine Neurotransmission Radioligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Acetylcholine Radioligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Central Benzodiazepine‐Binding Site Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Peripheral Benzodiazepine Receptor or Benzodiazepine‐Binding Site Receptor Ligands . . . . . . . . . . 94 Glutamate Neurotransmission Radioligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94 Cannabinoid Neurotransmission Radioligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Opioid Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95

9

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95

#

2008 Springer ScienceþBusiness Media, LLC.

76

3

In vivo imaging of neurotransmitter systems with PET

Abstract: The advent of functional neuroimaging techniques has significantly widened the methodological repertoire of neurochemistry. Using positron emission tomography PET, the human and non-human primate brain’s neurotransmitter and neuroreceptor systems can be studied in vivo. With the help of PET the distribution of the various neurotransmitter and neuroreceptor systems can be localized in precise anatomical context and several parameters of these systems can be measured in a quantitative manner. The basics of the technique, development of radiolabelled ligands, modeling and measuring radioligand effects in the brain are, among others, those key issues that are discussed concisely in the present chapter. List of Abbreviations: CT, Computed tomography; FDG, fluoro-deoxy-glucose; HPLC, high performance liquid chromatography; MRI, magnetic resonance imaging; NMSP, N‐methylspiperone; PBBS, peripheral benzodiazepine‐binding site; PET, positron emission tomography; SA, specific activity; SPECT, single‐ photon‐emission computed tomography; SR, specific radioactivity

1

Introduction: Neurotransmitter and Neuroreceptor Systems and In Vivo Neuroimaging

In addition to ex vivo–in vitro biochemical probes and postmortem brain imaging techniques, functional neuroimaging techniques, including single‐photon‐emission computed tomography or SPECT and positron emission tomography or PET, have in recent decades become inevitable parts of the methodological armory for the exploration of the human and nonhuman primate brain’s neurotransmitter and neuroreceptor systems. The present chapter focuses on PET research. And as the recent advances in PET radiochemistry result in the emergence of newer and newer PET radioligands with an increasing pace, in the present chapter the major emphasis is on the basic principles of PET radioligand development, and no emphasis is laid on a comprehensive summary of all available radioligands. For this reason, in the first part of this chapter a somewhat detailed overview on the bases of PET neuroreceptor research is given, whereas in the second part of the chapter we provide the reader with a short survey of those established PET radioligands, which have gained application grounds in a number of PET laboratories. During the past decades, well over a hundred different neurotransmitter subsystems have been identified and described in detail. In parallel with this, radiochemistry research has focused on the development of labeled PET radioligands to cover as wide as possible a range of biological markers usable for in vivo brain mapping with PET. As several neurotransmitter and receptor systems have been demonstrated to be directly involved in neurological or psychiatric diseases, drug addiction, or personality disorders, the importance of PET in receptor‐system‐related drug research has increased tremendously in recent years. Take an example! > Figure 3-1 is a schematic overview of the presynaptic generation and synaptic transmission of one of the key monoamine neurotransmitters, dopamine. Dopamine, similarly to several other central neurotransmitters, in addition to its extrasynaptic effects, predominantly acts as a synaptic transmitter (and this is displayed in the figure). Until now, two major families and five subtypes of the dopamine receptor have been identified (the dopamine D1 and D5 receptors in one family, and the D2, D3, and D4 receptors in the other). Most of these receptor subtypes can be targeted with PET radioligands. These receptors are, at the same time, the major targets of neuropsychiatric drugs used e.g., in schizophrenia or other psychiatric diseases. In addition to the receptor systems, the transporter molecule, responsible for the reuptake of the intrasynaptic dopamine molecules, is also a major target of neuropsychiatric drug research, since this molecule is predominantly responsible for the regulation of the intrasynaptic dopamine concentration. Consequently, the transporter system can also be a prime target for PET radioligand development. As exemplified by the figure, in vivo neuroimaging with PET using labeled ligands or ‘‘radioligands’’ can ‘‘visualize’’ the various receptor and transporter systems and measure in quantitative terms their densities, binding and occupancy status, and other parameters. Naturally, for the visualization of these systems other imaging techniques, for example postmortem autoradiography, can also be used. Consequently, the application of radioligands can also help us understand the role of various neurotransmitter systems and their concerted behavior (‘‘receptor fingerprint’’) in the normal and pathological functioning of the human brain. Furthermore, PET imaging of central neuroreceptor systems can greatly contribute to our efforts in developing novel drugs targeting dedicated receptor sites in the human brain.

In vivo imaging of neurotransmitter systems with PET . Figure 3-1 A schematic overview of synaptic dopamine neurotransmission. A number of relevant 3 H‐ or 125I‐labeled autoradiography radioligands are indicated in the figure

2

11

3

C‐labeled PET and

Positron Emission Tomography

PET is a noninvasive biological imaging technique that can quantitatively measure the distribution of radiolabeled molecules in the living body. Although the spatial resolution of the technique is lagging behind that of dedicated anatomical imaging techniques, in combination with anatomical/morphological imaging methods (CT or MRI), PET can map ‘‘quantitatively’’ biochemical and physiological parameters in a proper anatomical context. On the other hand, the temporal resolution of the method allows us to follow the biodynamics of the labeled molecules in the brain and body. A PET laboratory consists of expensive and sophisticated instrumentation and its activities require concerted actions from a number of experts of various disciplines (> Figure 3-2). The basis of the technique is the detectability of g photons, generated during the annihilation of positrons and electrons (> Figure 3-3). Positron‐emitting radionuclides, including those which are most commonly used in PET (> Table 3-1), decay by emitting a positively charged electron, a positron. The positron is the counterpart of the electron: its mass is identical with that of the electron, but it has a positive charge. Within a short distance (1–2 mm) from the place of the decay, the emitted positron encounters an electron. During the encounter, the two annihilate each other by releasing two 511 keV g photons in opposite directions along an axis. These two g photons can be detected by a pair of scintillation crystals (e.g., NaI or BiGe crystals), which transform the g photons into photons in the range of the visible light spectrum. These scintillation photons are amplified in photomultipliers, and the resulting signals can be detected by a coincidence circuit (> Figure 3-3). In the PET scanner, the detectors are built into detector rings (> Figure 3-4a). Inside a detector ring, with the help of the functional logic of coincidence circuits, one detector can be ‘‘functionally coupled’’ to several other detectors, i.e., a large number of detector channels can be formed (> Figure 3-4b). Several

77

78

3

In vivo imaging of neurotransmitter systems with PET

. Figure 3-2 The components and main functions of a complex PET center

. Figure 3-3 The physical basis of positron emission and positron–electron annihilation, and coincidence detection with the help of a functionally coupled detector system

. Table 3-1 Positron‐emitting radionuclides commonly used in PET Nuclide T½ (min) Target Nuclear reaction Radioactive decay Maximal energy (MeV) Specific radioactivity (Ci/mmol) Effective dose equivalent per 100 MBq radiotracer (mSv) Common forms Critical organ

11

C 20.3 NþO 14 N(p,a)!11C 11 11 0 6C ! 5B þ þ1e 0.97 9
106 0.4

13

N 9.98 H2O 12 C(d,n)!13N

15

O 2.05 NþO 14 N(d,n)!15O

18

1.20 19
106 0.25

1.74 90
106 0.1

0.64 1.7
106 2.5

11

13

N, 13NH3 Liver

15

O2, C15O Lung

H18F, 18F2 Bladder

CO, 11CO2 Liver

F 110 H218O 18 O(p,n)!18F

In vivo imaging of neurotransmitter systems with PET

3

. Figure 3-4 (a) Detector rings inside the scanner. (b) The possible arrangements of detector channels, related to one given detector, in a detector ring. (c) A PET scanner

rings can form the gantry of the scanner so that a larger section of the body can fully be covered by the field of view of the system (> Figure 3-4c). The detected radioactivity distributions inside the body can be reconstructed with the help of filtered backprojection algorithms (> Figure 3-5). Using additional physiological information and mathematical models, from the radioactivity distribution maps, obtained by the PETscanner, a large number of biochemical parameters can be estimated (> Table 3-2).

3

Labeling Tracers and Ligands with PET Bioisotopes

Among the PET radionuclides, the frequently utilized 18F has earlier been used in one single form: FDG. More recently, a burgeoning variety of radioligands is available with this radionuclide. The use of 13N and 15 O is more limited, mainly owing to the shorter half‐times of these radionuclides. 15O‐Labeled water or butanol is used in blood flow studies, whereas 13N‐labeled ammonia can be used in cardiac PET studies. Definitely, the most widely used PET radionuclide is 11C. For its ‘‘comfortable’’ half‐time (20 min) and carbon’s central role in organic chemistry, 11C has for long been used in the widest variety of PET radiochemistry applications. An important prerequisite for radioligand development is that the molecule maintains its properties after labeling. This is one further reason for the common use of the short‐lived

79

80

3

In vivo imaging of neurotransmitter systems with PET

. Figure 3-5 Schematic diagram showing the procedure of filtered backprojection. (a) The original biological target has a radioactivity distribution pattern inside the scanner’s gantry. (b) The main question of data sampling and reconstruction centers around the reliability of the generated image. (c) In the special case of a point source centered in the gantry, all projections, regardless of angle, will be the same. (d) If these projections were to be backprojected, the reconstructed image would be smeared. (e) Instead, a convolution between the profile and the reconstruction filter is applied, creating a filtered profile. (f) The filtered profile can then be used to successfully reconstruct the original image

. Table 3-2 Physiological–biochemical parameters measurable with PET Blood flow Blood volume Protein synthesis Molecular diffusion Tissue pH Metabolism of oxygen, glucose, amino acids, fatty acids, fluor, etc. Receptor and transporter systems: uptake, distribution, binding, occupancy Pharmacodynamics and pharmacokinetics of labeled drugs

positron‐emitting radionuclide 11C, for the substitution of naturally occurring 12C with 11C does not change the biochemistry or the pharmacology of the ligand molecule. The range and number of the applications of 11C‐labeled receptor ligands and drugs have steadily been increasing. The most widely used approach for labeling ligands using 11C is [11C]methylation. A typical total synthesis time for a 11C‐labeled radioligand including HPLC purification is 30 min. The reaction may require the presence of base for the generation of the nucleophile. Moreover, [11C]acylations with [11C]acyl

In vivo imaging of neurotransmitter systems with PET

3

chlorides and [11C]cyanation with [11C]cyanide are two common types of reactions, but, for instance, [11C] ethylation and other operations can also be used in [11C] radiochemistry. Nowadays, 18F is used more and more often for labeling central nervous system (CNS) ligands. The unique advantage of 18F lies in its relatively longer half‐time: 110 min, which provides us with the opportunity of transporting the labeled ligands to larger distances, i.e., distributing [18F]‐labeled radioligands to PET scanners not adjacent to a cyclotron facility. The production and application of PET radioligands, used for human studies, requires several consecutive steps, the timing of which is of vital importance due to the decay of the radionuclide and the loss of specific activity (SA) of the labeled radioligand (> Figure 3-6). A general rule of thumb is that the radiochemical synthesis time should start without a delay after the production of the radionuclide and it should be completed within 4–5 half‐life times of the radionuclide, whereas the end of batch efficacy

. Figure 3-6 The flowchart of radioligand production and its use in PET measurements

(radioactivity in the end‐product versus initial radioactivity upon the start of the radiochemistry process) should be over 20%. As in most cases, the radioligand’s SA (the proportion of labeled versus unlabeled molecules in the batch) is an essential parameter and the highest possible SA values are favored; the radioligand should be administered without any significant delay. These rules are less strict in the case of 18 F ligands, as compared with 11C ligands. (Fowler et al. (1999); Guly´as et al. (1998); Haldin et al. (1991, 2001, 2004); Heiss et al. (2001)).

81

82

3 4

In vivo imaging of neurotransmitter systems with PET

Preliminary Steps in the Development of Radioligands for Human CNS Receptors

When a potential candidate ligand has been identified and a radiochemical labeling technique developed, some preclinical evaluation, before PET in humans, needs to be performed. In the first instance, useful information may be obtained by studies in rodents. Typically, the radioligand is injected intravenously into a series of rodents. These are then sacrificed at known times after injection and either the animal’s different organs are removed and radioactivity is counted, thus providing the distribution of the radiotracer in different organs over time (tissue distribution study), or the body is sliced, the slices are mounted on glass, and exposed to radiosensitive film (ex vivo autoradiography). In addition to this, clearance of radioactivity from plasma and information on the appearance of labeled metabolites in plasma can be obtained. Specificity of binding may be demonstrated by using selective and potent unlabeled compounds that act in a competing way at the site of interest. A complementary tool in early receptor radioligand development may be autoradiography experiments, wherein frozen slices of tissue obtained from the organ of interest (e.g., brain) are mounted on glass slides and incubated for a given time with a buffered radioligand solution. These sections are exposed to radiation‐sensitive film. Autoradiography may provide information about a radioligand’s suitability for PET studies with special regard to its affinity, selectivity, and nonspecific binding. An advantage compared with in vitro homogenate binding assays is the use of intact tissue, which provides information in an anatomical context. However, no data regarding the in vivo pharmacokinetics of a novel ligand can be deduced from such an experiment. This lack of information can be compensated for by ex vivo autoradiography approaches, where, as mentioned earlier, the analysis is done in vitro after in vivo administration of the radioligand into small animals. Traditionally, autoradiography experiments are performed with 3H‐ or 125 I‐labeled compounds, but the use of b‐sensitive film makes the procedure also suitable for molecules labeled with positron emitters, even though this lowers the achievable spatial resolution. An useful variant

. Table 3-3 Important receptor-mapping parameters in in vivo imaging of neurotrasmitter systems with PET Parameter In vitro target affinity Selectivity (relative affinity to competing binding sites) Concentration in binding sites Binding affinity Reversibility of binding Dissociation rate constant Equilibrium dissociation constant Target affinity/nonspecific affinity ratio Potency Toxicity Specific binding Nonspecific binding Plasma protein binding Blood–brain barrier permeability Equilibrium Tissue clearance Distribution volume Binding potential Occupancy Receptor saturation

Kd/Ki/IC50/Km

Unit nM

Bmax

pM/ml

k–1/koff/k4 koff Kd

sec–1, min–1

EC50 LD50

mM mg/kg Bmax

log P log P

BP %

In vivo imaging of neurotransmitter systems with PET

3

of this method is whole‐hemisphere autoradiography of the postmortem human brain. A comparison between this technique and PET imaging is shown in > Table 3-4.

. Table 3-4 A comparison of postmortem autoradiography (PMAR) and PET PMAR

PET

Advantage High spatial resolution Quantifiable Relatively easy pharmacology Easy receptor discrimination Behavior, disease versus distribution, binding

Disadvantage Postmortem Ante finem effects? (Drug treatment) Postmortem changes/deterioration? Static (snapshot) Limited availability Lower resolution In vivo limiting factors (blood–brain barrier, in vivo metabolism, etc.)

Quantifiable Test–retest possible Kinetic follow‐up Larger populations possible

Ligand‐dependent selectivity

The next step in receptor radioligand development is normally PET investigations in animals. For brain‐ imaging studies, nonhuman primates—such as cynomolgus monkey (Macaca fascicularis)—are preferred. Analysis of labeled metabolites from venous blood samples provides useful information regarding clearance and metabolic pathways. Administration of potent and selective competing compounds before radioligand injection (pretreatment) or during the time course of the PET experiment (displacement) can demonstrate the specificity and reversibility of radioligand binding (see later). It should be noted, however, that species differences may be encountered and lead to different results between animals and human subjects.

5

Measuring Radioligand Effects in the Brain

The great advantage of the PET technique is that it is capable of obtaining absolute measurements of regional radioactivity concentrations which, in turn, with the help of appropriate kinetic models, can be transformed into quantitative parametric maps of related receptor parameters. What is relevant from our point of view is that several parameters related to the distribution and density of receptor systems and the ligands’ interaction with the receptors can also be measured (> Table 3-5). The initial selection of radioligands for neurotransmitter binding sites, such as receptors, neuronal uptake systems, and vesicular uptake systems, is often guided by data obtained in vitro by using tritiated or iodinated radioligands or by displacing a reference radioligand with the unlabeled molecule. In vitro binding normally provides information regarding ligand ‘‘affinity’’ (e.g., the dissociation equilibrium constants Kd or Ki) and ‘‘selectivity’’ (i.e., the relative affinity to competing binding sites) as well as regarding the ‘‘concentration’’ of binding sites (Bmax). The optimum affinity is closely related to the expected Bmax. It is preferable if the Bmax clearly exceeds the Kd of a ligand, i.e., if a binding site exists in vivo at nanomolar concentrations, a potentially successful radioligand ideally should have a subnanomolar affinity. Binding affinity is an important factor that determines the ratio of specific binding to nonspecific binding. The higher the ratio the more sensitive the signal is likely to be to changes in available binding site concentration, caused by disease or drug occupancy. Binding affinity (i.e., the fraction of dissociation rate constant, koff, and association rate constant, kon) usually governs the approach to be taken in the biomathematical modeling of the ligand–receptor

83

84

3

In vivo imaging of neurotransmitter systems with PET

. Table 3-5 Optimal criteria of PET ligands for neuroreceptors Aspect Radiochemical development

Biochemical properties

Favorable brain exposure

Key criteria  Should be radiolabeled with 11C or 18F  Radiolabeling should yield high specific radioactivity  Should be radiolabeled in a position not subject to formation of radioactive metabolites that pass the BBB  Sufficient affinity to the target receptor system (0.01–5 nM; the ideal affinity is dependent on target expression level)  High selectivity for the target receptor system (ideally 100‐fold as compared with any other binding sites)  Low nonspecific binding  As few as possible labeled metabolites in the brain; if they occur, their concentrations should be low with rapid clearance  Good permeation through the blood–brain barrier  Lipophilic (log P = 1–4)  Not a substrate for P‐glycoproteins  Low volume of distribution (Vd)  Low protein binding in plasma  No trapping in peripheral compartments

interaction. If the binding of a radioligand is reversible over the timescale of a PET experiment, ‘‘equilibrium’’ approaches toward quantification can be utilized. On the contrary, irreversible ligands normally demand ‘‘kinetic’’ modeling, wherein the transfer of radioligand between pharmacological compartments (e.g., plasma, tissue, receptors) is described in terms of rate constants. This approach requires in most cases the determination of an input function (i.e., the time course of free radioligand in plasma), which makes the measurement of radioligand metabolites in arterial plasma necessary. Very high binding affinity of a radioligand in combination with a comparatively slow clearance from tissue can restrict its usefulness for PET, as the rate‐limiting step of tracer retention may become the delivery instead of the binding process (flow‐limited conditions). A further important criterion for a radioligand is binding selectivity. Ideally, the affinity of a radioligand should be highest for the site of interest by more than one order of magnitude. However, lack of selectivity may be acceptable if nontarget sites are separated anatomically from the target‐binding sites. In the light of advances in molecular biology and pharmacology the term selectivity often needs to be revised. Most neurotransmitter receptors have now been found to exhibit multiple subtypes, and ligands that were initially thought to bind to a single class of receptors, truly display affinity toward several subtypes. This fact is also reflected by different ‘‘research philosophies’’ in drug development: whether ‘‘pharmacologically clean’’ molecules need to be developed with extremely high affinity to a given receptor system and low to other systems, therefore exhibiting high selectivity; or molecules with ‘‘rich pharmacology’’ are preferred, with an ‘‘optimal’’ mixture of affinities to a number of different receptor systems. Another substantial consideration in the development of a new radioligand is estimation of nonspecific binding. This is an essentially nonsaturable component of the total tissue uptake of a radioligand, usually attributed to adhesion to proteins and lipids. Nonspecific binding and its clearance in vivo are difficult to predict with absolute confidence. However, within a class of structurally related compounds, nonspecific interactions with tissue generally increase with increased lipophilicity. The logarithm of the partition coefficient between water (or preferably buffer to account for ionization at physiological pH) and octanol (log P) is often taken as a useful index for the lipophilicity of a compound in the context of biological systems. Conversely, some degree of lipid solubility is needed for good passage over the blood–brain barrier, which is a prerequisite for satisfactory counting statistics. However, the lipophilic nature of a molecule might also favor binding to plasma proteins, thus reducing the available ‘‘free fraction’’ in blood that is

In vivo imaging of neurotransmitter systems with PET

3

capable of diffusing through membranes. Moreover, lipophilic molecules can be extracted and metabolized in lung tissue when passing through the lung circulation, which prevents them from reaching their sites of action. Taken together, it appears that there is an optimal—but rather narrow—‘‘window’’ of lipophilicity for brain radioligands, wherein brain uptake is high and nonspecific binding comparatively weak. PET can quantitatively measure the regional radioactivity concentration without being able to distinguish the chemical forms or environments in which the radioactivity resides. For a clearly interpretable signal, it is therefore necessary that radiolabeled metabolites do not contribute to specific binding. Thus, radioligands should be preferably resistant to rapid metabolism over the period of data acquisition. Furthermore, radiolabeled metabolites should not be taken up and/or retained in the target area. This requirement may have important consequences concerning the elaboration of a radiolabeling strategy. A very important consideration in the context of radiochemistry is specific radioactivity (SR) of the radioligand. Too low SR may result in pharmacological effects or toxicity of the radiotracer. Moreover, low SR may saturate the biological system of interest, thus abolishing the mandatory tracer conditions. For low‐density binding sites very high SR is essential in order to exclude a substantial occupation of target sites by unlabeled ligand. The controlled administration of high SR radioligands versus low SR ligands may help estimate the binding profile of the labeled molecule as well as the general nature or actual status of the binding site. A major consideration is related to the intrinsic activity and efficacy of the prospective ligand as well as the developed radioligand on a given receptor system. The ligands have an intrinsic activity which is (1) 0 for antagonists fully blocking a receptor system; (2) vary between 0 and 1 for agonists (partial or full agonists); and (3) vary between 0 and –1 for inverse agonists (partial or full inverse agonists). The intrinsic efficacy of a ligand on a receptor can vary between a maximal effect and a minimal or no effect. An example for these properties is given in > Figure 3-7.

6

Modeling Ligand Effects in the Brain

The great advantage of the PET technique, in contrast to other functional imaging modalities, is that it is capable of obtaining absolute measurements of regional radioactivity concentrations which, in turn, with the help of appropriate tracer kinetic models can be transformed into quantitative parametric maps of various biochemical, physiological, or neurotransmitter/receptor system‐related parameters. The quantitative measurements with PET require the determination of tissue and blood/plasma radioactivity concentrations and the a priori knowledge of a number of experimental parameters, including basic features related to the tracer, the biochemical and physiological characteristics, metabolic stability of the tracer/ligand, and those of the scanner. In the next step, a multicompartmental model describing the distribution and metabolism of the ligand in the brain and, eventually, in other body compartments, is developed, tested, and validated. With the help of appropriate tracer kinetic models the requested biological variables, e.g., receptor occupancy data, can be described in quantitative terms in precise anatomical context in the organ covered by the PET scan (> Figure 3-8).

7

Two Approaches

The PET technique is sensitive for determinations of concentrations as low as the sub‐picomolar range (10–12 mol/l). Effective radiochemical labeling provides a radioligand with high SR, i.e., with a high ratio of radiolabeled to unlabeled drug molecules. A consequence is that i.v. injection of less than a microgram (mg) of the radiolabeled drug is sufficient for a PET study in man. The concept ‘‘tracer dose’’ is often used to emphasize the low mass, which does not induce drug effects. Similarly to neuropsychopharmacological drug development studies with PET, there are two possible experimental designs usable in mapping neuroreceptor systems. 1. The direct approach: to radiolabel the prospective ligand and measure its uptake, anatomical distribution, and binding in the brain. 2. The indirect approach: to study how an unlabeled molecule inhibits the specific binding of a well‐ characterized selective PET radioligand.

85

86

3

In vivo imaging of neurotransmitter systems with PET

. Figure 3-7 Intrinsic activity and effect of compounds labeled as existing or prospective PET radioligands for GABAA– benzodiazepine receptors. The ligands have an intrinsic activity which may change for agonists between 0 and 1, for inverse agonists between 0 and 1, and 0 for antagonists that can fully block a receptor. The intrinsic efficacy of a ligand on a receptor can also vary between maximal effect and no effect. In the case of benzodiazepine receptors, those ligands have positive intrinsic efficacy that increase GABAergic neurotransmission. The lower part of the figure shows recently developed compounds, a part of which are already used as antipsychotic drugs, whereas others are used as radioligands

7.1 Direct Approach: Radiolabeling of Ligands In this case the ligand can be radiolabeled and administered intravenously. The ligand’s brain uptake and distribution can be visualized and, alongside with a number of binding parameters, can be measured in a quantitative manner with PET (> Figure 3-9).

7.2 Indirect Approach: Using Radiolabeled Ligands and Drug Candidates In this case the unlabeled drug (ligand, drug, or drug candidate molecule) is competing with a well‐ characterized radioligand for occupying a receptor system (> Figure 3-10). The competition between the two molecules is not really a ‘‘fair competition’’, as the amount of the unlabeled drug exceeds that of the labeled drug. The unlabeled drug is given in pharmacological dose (mg range) whereas the labeled ligand is given in tracer dose (pico‐nano‐microgram range), the two doses being different in several orders of

In vivo imaging of neurotransmitter systems with PET

3

. Figure 3-8 The principles of quantitative PET imaging. The datasets required for acquiring quantitative data needed for modeling are: tissue radioactivity measurements, obtained with a PET scanner, and blood (and, eventually, plasma) radioactivity measurements, obtained by way of regular blood sampling. Quantitative models are usually compartmental models with the help of which biochemical parameters can be estimated in a qualitative manner

magnitude. This is done in order to occupy a given receptor system as completely as possible by the unlabeled drug and test the occupancy or ‘‘blockade’’ of the system. This can be achieved either by giving the unlabeled drug before the administration of the labeled drug (pretreatment) (see > Figure 3-11a) or after its administration (displacement) (> Figure 3-11b).

8

Radioligands for Mapping Neurotransmitter Systems: Some Examples

A short overview of the central neuroreceptor systems with useful radioligands is given in > Table 3-6 and PET images, obtained in humans, of a few representative PET radioligands are shown in > Figure 3-12. Despite the fact that the monoamine neurotransmitter systems represent only a fraction of the central neuroreceptors, the most useful and best‐characterized PET radioligands are available for these systems; more specifically: the dopamine and serotonin systems. Intense studies have been focused in recent years to target other neuroreceptor systems, as well. Furthermore, PET radioligand development has also been focusing nowadays on the development of useful radiolabeled biomarkers for other than receptor binding sites in the brain, including amyloid plaques.

87

88

3

In vivo imaging of neurotransmitter systems with PET

. Figure 3-9 The direct approach involves the radiolabeling of a potential novel ligand and to trace its anatomical distribution and measure its binding in the brain

. Figure 3-10 A basic principle used in PET radioligand development. A well‐characterized labeled radioligand (used in tracer dose) is competing with a drug (used in therapeutic dose) for the same receptor system

In vivo imaging of neurotransmitter systems with PET

3

. Figure 3-11 The principles of pretreatment and displacement. The baseline curve (indicated with rhomboids) displays the normal kinetic behavior of a radioligand in the brain or a brain structure. If a drug, having affinity for the same receptor system and thereby blocking it, is administered before the radioligand injection in pharmacological dose, the radiolabeled ligand cannot bind to the blocked receptors in the same amount as in the baseline condition. This is called pretreatment condition (indicated with squares). If a drug, having affinity for the same receptor system and thereby blocking it, is administered after the radioligand injection in pharmacological dose (injection time is indicated by the arrow), the radiolabeled ligand, already bound to the receptors, will compete with the unlabeled drug. Because of the concentration differences between the labeled ligand and the unlabeled drug (tracer dose versus pharmacological dose; concentration differences usually over four orders of magnitude), the unlabeled drug will displace the radioligands bound to the receptors. This is called displacement condition (indicated with triangles)

8.1 Dopamine Receptor and Transporter Ligands Dopamine exerts its signaling effect by binding to specific membrane bound receptors, which belong to the family of G protein‐coupled receptors. Detailed molecular genetic studies have shown that the five types of dopamine receptors belong to two receptor families: D2, D3, and D4 receptors form one family, whereas D1 and D5 receptors form the other (> Table 3-7). Until now, only D1 and D2 receptors have been successfully visualized in vivo in humans, whereas there exist no useful selective radioligands for the dopamine D3 and D4 subtypes. There is a need to develop highly selective dopamine receptor radioligands for all the five subtypes, with special regard to the D3, D4, and D5 subtypes. The available dopamine radioligands labeled with b‐emitters have been extensively used in investigation of the dopamine receptors in physiology, neurological and mental disorders, and clinical pharmacology. Despite the fact that of the several billions of neurons in the human brain only a very small fraction (approximately 0.0003–0.0004%) use dopamine as a neurotransmitter; dopamine signaling plays a cardinal role in several brain functions, including locomotor control, positive reinforcement, cognitive functions, personality traits, and neuroendocrine regulation. Alterations of dopaminergic neurotransmission have been implicated in the pathophysiology of several neuropsychiatric disorders, such as Parkinson’s disease, Huntington’s disease, schizophrenia, ADHD, and drug abuse. The recently available highly useful PET radioligands of the dopamine receptor system use a few leads: the benzazepines such as [11C]SCH 23390 and [11C]NNC 112 bind to both D1 and D5 and the benzamides such as [11C]raclopride bind to both D2 and D3.

89

90

3

In vivo imaging of neurotransmitter systems with PET

. Table 3-6 Some representative PET radioligands used to study neuroreceptors in man Neurotransmitter systems Dopamine D1 Dopamine D2

Dopamine D3 Dopamine transporter

Serotonin 5‐HT1A

Serotonin 5‐HT2A

Serotonin transporter

Norepinephrine transporter

Glutamate Opiate

Muscarinic

Nicotinic

Radioligand [11C]‐SCH 23390 [11C]‐NNC 112 [11C]‐Raclopride [11C]‐NMSP [11C]‐FLB 457 [11C]‐MNPA [18F]‐Fallypride [18F]‐Fluoroethylspiperone [11C]‐RGH1756 [11C]‐MMC [11C]‐Methylphenidate [11C]‐PE2I [11C]‐b‐CIT‐FE [11C]‐b‐CFT [11C]‐Altropane [11C]‐Cocaine [18F]‐b‐CIT‐FP [Carbonyl‐11C]‐WAY‐100635 [11C]‐DWAY [11C]‐FCWAY [11C]‐NAD299 [11C]‐NMSP [11C]‐MDL 100907 [18F]‐Altanserin [18F]‐Setoperone [11C]‐DASB [11C]‐MADAM [11C]‐McN5652 [11C]‐nor‐b‐CIT [11C]‐MeNER

Selected references Halldin et al. (1986); Farde et al. (1987) Halldin et al. (1998a); Abi‐Dargham et al. (2000) Farde et al. (1986); Halldin et al. (1991) Wagner et al. (1983); Burns et al. (1984) Halldin et al. (1995); Olsson et al. (1999) Finnema et al. (2005); Seneca et al. (2005) Mukherjee et al. (1999); Christian et al. (2000) Jovkar et al. (1990); Wienhard et al. (1990) So´va´go´ et al. (2004, 2005) Gao et al. (2005) Ding et al. (1994); Volkow et al. (1998) Halldin et al. (1998a); Dolle´ et al. (2000) Halldin et al. (1996); Farde et al. (2000) Laakso et al. (1998a); Tsukada et al. (2001a) Madras et al. (1998); Fischman et al. (2001) Fowler et al. (1989); Volkow et al. (1999) Chaly et al. (1996); Lundkvist et al. (1997) Pike et al. (1996); Farde et al. (1998) Pike et al. (1998); Marchais‐Oberwinkler et al. (2005) Lang et al. (1999); Carson et al. (2000, 2002) Sandell et al. (1999, 2002); Andree et al. (2003) Burns et al. (1984); Andree et al. (1998) Lundkvist et al. (1996); Ito et al. (1998) Crouzel et al. (1992); Biver et al. (1994) Blin et al. (1990); Crouzel et al. (1992) Houle et al. (2000); Wilson et al. (2002) Halldin et al. (2005); Lundberg et al. (2005) Szabo´ et al. (1995, 1996) Mu¨ller et al. (1993); Bergstro¨m et al. (1997) Schou et al. (2003)

[18F]‐FMeNER [18F]‐FD2MeNER [11C]‐Desipramine [11C]‐Talopram [11C]‐Talsupram [11C]‐MPEP [11C]‐JNJ‐16567083) [11C]‐Diprenorphine [11C]‐Carfentanil [18F]‐Cyclofoxy [11C]‐NMPD [11C]‐3‐MPB [11C]‐Benztropine [11C]‐Nicotine [11C]‐Mecamylamine

Schou et al. (2004) Schou et al. (2005); Seneca et al. (2005) Van Dort et al. (1997); Schou et al. (2006) McConathy et al. (2004); Schou et al. (2006) Schou et al. (2004); McConathy et al. (2004) Yu et al. (2005) Huang et al. (2005) Mayberg et al. (1991); Jones et al. (1994) Dannals et al. (1985); Frost et al. (1989) Theodore et al. (1992); Cohen et al. (1998) Mulholland et al. (1995); Zubieta et al. (2001) Takahashi et al. (1999); Tsukada et al. (2001a) Dewey et al. (1990); Fujiwara et al. (1996) Nordberg et al. (1989) Sobrio et al. (2005)

In vivo imaging of neurotransmitter systems with PET

3

. Table 3-6 (continued) Neurotransmitter systems Histamine Adenosine Neurokinin‐1 GABA/benzodiazepine Peripheral benzodiazepine

Radioligand 2‐[18F]Fluoro‐A‐85380 [11C]‐Doxepin [11C]‐Pyrilamine [11C]‐MDPX [18F]‐CPFPX [18C]‐SPA‐RQ [11C]‐Flumazenil [11C]‐RO 5‐4513 [11C]‐PK11195 [11C]‐Vinpocetine [11C]‐DAA1106

Selected references Gallezot et al. (2004); Obrzut et al. (2005) Tashiro et al. (2004); Iwabuchi et al. (2005) Szabo´ et al. (1993); Kim et al. (1999) Fukumitsu et al. (2003, 2005); Kimura et al. (2004) Bauer et al. (2003); Meyer et al. (2006) Bergstro¨m et al. (2004); Solin et al. (2004); Hietala et al. (2005) Mazie`re et al. (1984); Persson et al. (1989) Halldin et al. (1992); Pike et al. (1993) Hashimoto et al. (1989); Banati et al. (1999) Gulya´s et al. (2005); Vas and Gulya´s, (2005) Okuyama et al. (1999); Maeda et al. (2004)

A benzazepine derivative, SCH 23390, has been described as the first high‐affinity selective dopamine D1 receptor antagonist and labeled with positron‐emitting 11C. An improved benzazepine compound [11C]NNC 112 has also been developed and is now used worldwide. The gold standard of dopamine D2 receptor ligands is 11C‐raclopride, which has permitted the selective analysis by PET of central D2 receptors in primates and humans. Radiolabeled benzamides of picomolar affinities, which make them suitable to examine the extrastriatal D2 receptors in the human brain, have been developed during the recent years, such as [11C]FLB 457. The butyrophenon N‐[11C]methylspiperone ([11C]NMSP) is also showing promising features. Whereas these ligands are antagonists, the newly developed agonist radioligand, [11C]MNPA, shows high binding affinity to the D2 receptor system. An 18F‐labeled benzamide, also suitable for investigation of extrastriatal D2 receptors, is 18F‐fallypride. The labeling of dopamine D3 receptors is a challenge, and whereas a few candidates have been tested (e.g., [11C]‐RGH1756), no selective radioligand is available at this time. The dopamine reuptake sites play an effective role in the regulation of the intrasynaptic dopamine concentration. Several attempts have been made to find appropriate PET ligands for the labeling of this system. The recently used ligands include [11C]‐PE2I (N‐(3‐iodoprop‐2E‐enyl)‐2b‐carbomethoxy‐3b‐(40 ‐ methylphenyl)nortropane) and [11C]‐FE‐CIT (N‐(2‐fluoroethyl)‐2 b‐carbomethoxy‐3b‐(4‐iodophenyl) nortropane). An overview of the useful dopamine receptor and transporter radioligands are shown in > Figure 3‐12a.

8.2 Serotoninergic Neurotransmission Radioligands In addition to the dopamine system, the serotonin (5‐hydroxytryptamine, 5‐HT) system is the other monoaminergic system for which a number of PET radioligands are available. The serotonin system consists of a large number of postsynaptic 5‐HT receptors, belonging to seven major families (5‐HT1–5‐HT7), each containing several subtypes, and the serotonin transporter, regulating the reuptake of the intrasynaptic serotonin into the presynaptic neuron. Within the 5HT1 group there are subtypes 5HT1A, 5HT1B, 5HT1D, 5HT1E, and 5HT1F. There are three 5HT2 subtypes, 5HT2A, 5HT2B, and 5HT2C as well as two 5HT5 subtypes, 5HT5B and 5HT5B. Most of these receptors are metabotropic; however, the 5HT3 class is, for instance, ionotropic. The most commonly studied serotonin receptor subsystem, the 5‐HT1A, is present in high densities in the hippocampus, septum, amygdala, hypothalamus, and neocortex. The other commonly studied receptor subsystem, the 5‐HT2A receptors, are present in the neocortex, followed by, in the hippocampus, basal ganglia, and thalamus. The transporter has high densities in midbrain structures, with special regard to the raphe nuclei, the thalamus, and the striatum.

91

92

3

In vivo imaging of neurotransmitter systems with PET

. Figure 3-12 Horizontal PET images of the human brain showing the cerebral uptake of some of the most commonly used PET radioligands. For details. see text and > Table 3-6

Radioligand development to all receptor subtypes is a challenge and a number of PET laboratories are working on this challenge. The recently available radioligands include the 5‐HT1A antagonist WAY‐100635 (N‐(2‐(4‐(2‐methoxyphenyl)‐1‐piperazinyl)ethyl)‐N‐(2‐pyridyl) cyclohexane‐carboxamide), the 5‐HT2A antagonists MDL100907 ((R)‐(þ)‐a‐(2,3‐dimethoxyphenyl)‐1‐[2‐(4‐fluoro‐phenyl)ethyl]‐4‐piperidine methanol] and N‐methylspiperone (NMSP), and the transporter ligands MADAM (11C‐N,N‐Dimethyl‐2‐ (2‐amino‐4‐methylphenylthio)benzylamine) and DASP (3‐amino‐4‐(2‐dimethylaminomethyl‐phenylsulfanyl)benzonitrile) (see > Figure 3‐12b).

In vivo imaging of neurotransmitter systems with PET

3

. Table 3-7 Human dopamine receptor subtypes Amino acid

D1 446

D2 D2A: 443 D2B: 414 11q22–q23 Yes Gi, Go cAMP Ca2þ channel þKþ channel IP3? Caudatus

Chromosome Introns G‐protein Coupling

5q31–34 No Gs þcAMP þIP3

Main localization in the CNS

Nucleus Caudatus Putamen

Putamen

Accumbens

Accumbens

Substantia nigra Cortex

Substantia nigra

D3 400

D4 387

D5 477

3q13.3 Yes Gi? cAMP?

11p15.5 Yes Gi cAMP?

4p15.1–16.1 No Gs þcAMP

Island of Calleja Accumbens

Entorhinal cortex

Hippocampus

Bed nucleus Stria terminalis

Hippocampus

Lateral mamillary bodies

Amygdala Medulla

8.3 Norepinephrine Neurotransmission Radioligands The third major family of CNS monoaminergic receptors is the noradrenaline or norepinephrine system. The radiochemistry of norepinephrine transporter ligands is, at this day, more developed that that of the receptor ligands. Among others, [11C]]MRB ([11C]O‐methylreboxetine), [11C]desipramine, and the 11C and 18F versions of MeNER ((S,S)‐2‐(a‐(2‐fluoromethoxyphenoxy)benzyl)morpholine), an O‐methyl analog of reboxetine, have recently been introduced to the PET community (see > Figure 3-12c).

8.4 Acetylcholine Radioligands The acetylcholine receptors are usually subdivided into nicotinic and muscarinic receptors, the former belonging to the ligand‐gated ion channels, whereas the latter being metabotropic receptors. Both systems have been implicated in several neurological functions and, consequently, in neurological or psychiatric diseases, including Alzheimer’s disease, Parkinson’s disease, and schizophrenia. Intensive radioligand development activities aim at targeting both systems. Despite this fact, no highly useful and widely used PET ligands exist for these two systems, despite the fact that a number of radioligands have recently been advocated, including [11C]NMPB, [11C]benztropin, [11C]scopolamine, [11C]IQNP, and [11C]IQNB for the muscarinic and [18F]2‐F‐A85380 and [11C]A84543 for the nicotinic systems.

8.5 Central Benzodiazepine‐Binding Site Ligands In the mammalian brain, two different benzodiazepine‐binding sites, the central and the peripheral types, have been characterized. The central type benzodiazepine‐binding site is part of the GABA/benzodiazepine/ Cl supramolecular receptor complex. This specific site mediates all pharmacological properties of

93

94

3

In vivo imaging of neurotransmitter systems with PET

the classical benzodiazepines (sedative, anxiolytic, anticonvulsant, and myorelaxant). The chloride channel‐ gating function of GABAA receptors in the CNS can be allosterically modulated by ligands acting at the benzodiazepine receptor. The compounds which have been labeled for visualizing central benzodiazepine receptors belong to three different structural chemical families: the classical benzodiazepines, the cyclopyrrolones, and the imidazobenzodiazepine derivatives. Several benzodiazepine agonists (flunitrazepam, diazepam, suriclone) have been prepared and used as radioligands. [11C]‐Flunitrazepam and [11C]‐diazepam led to the first visualization of benzodiazepine receptors in baboon and human brains. [11C]‐Suriclone, a cyclopyrrolone, has also been proposed for in vivo purposes. A search for a more specific central type benzodiazepine ligands has led to the labeling of antagonists such as oxoquazepam, a selective BZ1 subtype ligand, and flumazenil (RO151788), an imidazobenzodiazepine. Flumazenil has been labeled with 11C either by N‐methylation with [11C]‐methyl iodide or by esterification with [11C]‐ethyl iodide. An analogue of flumazenil labeled with 18F, [18F]‐fluoroethylflumazenil, appears to be a suitable PET ligand despite its lower affinity and more rapid metabolism and kinetics. [11C]‐Flumazenil is considered now as the reference tracer used for pharmacological and clinical PET studies of the central benzodiazepine receptor.

8.6 Peripheral Benzodiazepine Receptor or Benzodiazepine‐Binding Site Receptor Ligands Unlike the central benzodiazepine receptors which are located on the cell membrane, the peripheral benzodiazepine receptor (PBR) is localized in the mitochondrial and nuclear subcellular fractions. For this and other characteristics of the protein complex, it is different from the classical membrane receptors and is often termed as the peripheral benzodiazepine‐binding site (PBBS). The peripheral benzodiazepine receptor plays a cardinal role in cellular respiration, oxidative metabolism and ion transport, neurosteroid biosynthesis, porphyrin transport and heme synthesis, regulation of calcium flow, apoptosis, and several other cellular processes. In the brain, the PBR can be visualized in activated microglia and astrocytes. The isoquinoline PK11195, a ligand for the PBBS, binds with relative cellular selectivity to activated, but not resting, microglia. [11C]‐PK11195 has then be used to study inflammatory and neurodegenerative brain diseases in vivo using PET. Moreover, [11C]‐PK11195 appears to be a biomarker of neuronal injury not only at the primary lesion site but also at the antero‐ and retrograde projection areas of the lesioned neurons. More recently, other candidate radiomarkers have also been tested, including [11C]‐DAA1106 (N‐5‐fluoro‐2‐phenoxyphenyl)‐N‐(2‐hydroxy‐5‐methoxybenzyl)acetamide and [11C]‐vinpocetine (cis‐ ethyl‐apovincaminate). Vinpocetine enters the brain in larger proportions that PK11195, and with the help of it even the age‐related physiological increase of PBR densities in the human brain can be demonstrated (> Figure 3-12d).

8.7 Glutamate Neurotransmission Radioligands Metabotropic glutamate (mGlu) receptors play a vital role in normal brain functions, and consequently, also in neurological and psychiatric disorders. The precise functions of these receptors are still undefined. Progress toward understanding their functions has been hampered by the lack of selective ligands with appropriate pharmacokinetic properties. However, the glutamate system is a conundrum for PET radiochemistry. Despite years of intensive search for useful radioligands, no breakthrough has yet been reported in this field. A few promising approaches were published, using, among others, 2‐methyl‐6‐(phenylethynyl)‐ pyridine (MPEP) and its analogues, M‐MPEP and M‐PEPy, [11C]‐ABP688 (3‐(6‐methyl‐pyridin‐2‐ylethynyl)‐cyclohex‐2‐enone‐O‐[11C]‐methyl‐oxime), bis(phenylalkyl)amines, [11C]‐3‐[2‐[(3‐methoxyphenylamino)‐carbonyl]‐ethenyl]‐4,6‐dichloroindole‐2‐carboxylic acid (3MPICA), and N,N0 ‐diphenyl and

In vivo imaging of neurotransmitter systems with PET

3

N‐naphthyl‐N0 ‐phenyl guanidine derivatives. As none of these approaches have until now resulted in a usable PET radiomarker, this particular field is one of the greatest challenges nowadays of PET radioligand chemistry.

8.8 Cannabinoid Neurotransmission Radioligands In the endocannabinoid system two receptor subtypes, CB1 and CB2, are recognized. Both receptors belong to the G protein‐coupled superclass of receptors. CB1 receptors are located throughout the body including within the CNS at presynaptic nerve terminals. CB2 receptors are mainly associated with cells of the immune system. Brain CB1 receptors represent an interesting target for the treatment of several psychiatric (e.g., anxiety, addiction, depression) and neurodegenerative disorders (e.g., Huntington’s disease and Tourette’s syndrome). The role of the CB1 receptors in these disorders is not well understood. The search for adequate PET radioligands is recently centered around a few structures, including 1,5‐diarylpyrazole, 3‐(4‐fluoronaphthoyl)‐1‐(N‐methylpiperidin‐2‐ylmethyl)indole, N‐([18F]fluorophenyl)‐ 5‐(4‐bromophenyl)‐1‐(2,4‐dichlorophenyl)‐1H‐pyrazole‐3‐carboxamide, and O‐methyl‐[11C]‐1‐(2‐chlorophenyl)‐5‐(4‐methoxyphenyl)‐4‐methyl‐1H‐pyrazole‐3‐carboxylic acid piperidin‐1‐ylamide ([11C]‐1).

8.9 Opioid Receptors The opioid receptors belong to the G protein‐coupled receptor families. The opioid receptors, m, d and k, are the mediators of the pharmacological effects of opioid drugs. The receptor subtypes are broadly expressed in the CNS. Their activation by opioid compounds (including morphine, codeine, heroin) is intimately involved with reward, tolerance, and withdrawal. Their significant role in analgesia, antinociception, and drug addiction has been demonstrated extensively. Among the recently available PET radioligands, [11C]‐carfentanil, a m‐receptor agonist, has been studied most extensively. Furthermore, [11C]‐diprenorphine, a nonspecific antagonist, [18F]‐diprenorphine, a nonspecific antagonist, and [18F]‐cyclofoxy, a m‐ and k‐receptor antagonist, are worth mentioning. Ongoing research focuses on other structures, as well, including cyclohexyl piperazine, diprenorphine, benzamide analogues, and 4‐anilidopiperidines.

9

Conclusion

PET radioligands play an important role in the qualitative visualization and quantitative exploration and mapping of neuroreceptor systems in the primate brain. With the help of PET imaging using adequate radioligands, the distribution of various receptor systems in the brain, their densities, and occupancy levels can be quantitatively measured in precise anatomical context. Physiological situations can be explored in normal conditions and under various pharmacological challenges, as well as alterations related to disease conditions or long‐lasting drug therapies or addiction can be studied. PET will present for the coming years the ‘‘par excellence’’ research tool for studies aiming at the in vivo quantitative exploration and anatomical mapping of neuroreceptor systems in the human brain.

References Abi‐Dargham A, Martinez D, Mawlawi O, Simpson N, Hwang DR, et al. 2000. Measurement of striatal and extrastriatal dopamine D1 receptor binding potential with [11C]NNC 112 in humans: Validation and reproducibility. J Cereb Blood Flow Metab 20(2): 225-243.

Andree B, Nyberg S, Ito H, Ginovart N, Brunner F, et al. 1998. Positron emission tomographic analysis of dose-dependent MDL 100,907 binding to 5-hydroxytryptamine-2A receptors in the human brain. J Clin Psychopharmacol 18: 317-323.

95

96

3

In vivo imaging of neurotransmitter systems with PET

Andree B, Hedman A, Thorberg SO, Nilsson D, Halldin C, et al. 2003. Positron emission tomographic analysis of dose‐ dependent NAD‐299 binding to 5‐hydroxytryptamine‐1A receptors in the human brain. Psychopharmacology (Berl) 167(1): 37-45. Banati RB, Goerres GW, Myers R, Gunn RN, Turkheimer FE, et al. 1999. [11C](R)‐PK11195 positron emission tomography imaging of activated microglia in vivo in Rasmussen’s encephalitis. Neurology 53(9): 2199-2203. Bauer A, Holschbach MH, Meyer PT, Boy C, Herzog H, et al. 2003. In vivo imaging of adenosine A1 receptors in the human brain with [18F]CPFPX and positron emission tomography. Neuroimage 19(4): 1760-1769. Bergstro¨m KA, Halldin C, Hall H, Lundkvist C, Ginovart N, et al. 1997. In vitro and in vivo characterisation of nor‐b‐ CIT: A potential radioligand for visualisation of the serotonin transporter in the brain. Eur J Nucl Med 24(6): 596-601. Bergstro¨m M, Hargreaves RJ, Burns HD, Goldberg MR, Sciberras D, et al. 2004. Human positron emission tomography studies of brain neurokinin 1 receptor occupancy by aprepitant. Biol Psychiatry 55(10): 1007-1012. Biver F, Goldman S, Luxen A, et al. 1994. Multicompartmental study of fluorine‐18 altanserin binding to brain 5HT2 receptors in humans using positron emission tomography. Eur J Nucl Med 21: 937-946. Blin J, Sette G, Fiorelli M, Bletry O, Elghozi JL, et al. 1990. A method for the in vivo investigation of the serotonergic 5‐HT2 receptors in the human cerebral cortex using positron emission tomography and 18F‐labeled setoperone. J Neurochem 54(5): 1744-1754. Burns HD, Dannals RF, Langstro¨m B, et al. 1984. (3‐N‐[11C]‐ methyl)spiperone—a ligand binding to dopamine receptors: Radiochemical synthesis and biodistribution studies in mice. J Nucl Med 25: 1222-1227. Burns HD, Dannals RF, Langstrom B, Ravert HT, Zemyan SE, et al. 1984. (3‐N‐[11C]methyl)spiperone, a ligand binding to dopamine receptors: Radiochemical synthesis and biodistribution studies in mice. J Nucl Med 25(11): 1222-1227. Carson RE, Toczek MT, Lang L, et al. 2002. Human functional imaging with the 5‐HT1A ligand 18F‐FCWAY. J Nucl Med 43: 55. Carson RE, Lang L, Watabe H, Der MG, Adams HR, et al. 2000. PET evaluation of [(18)F]FCWAY, an analog of the 5-HT(1A) receptor antagonist, WAY-100635. Nucl Med Biol 27: 493-497. Chaly T, Dhawan V, Kazumata K, et al. 1996. Radiosynthesis of [18F]N‐3‐fluoropropyl‐2‐b‐carbomethoxy‐3‐b‐(4iodophenyl)nortropane and the first human study with positron emission tomography. Nucl Med Biol 23: 999-1004. Christian BT, Shi B, Mukherjee J. 2000. Quantification of striatal and extrastriatal D2 dopamine receptors using

PET imaging of [18F]fallypride in nonhuman primates. Synapse 38: 71-79. Cohen RM, Carson RE, Channing M, et al. 1998. [18F]‐cyclofoxy PET imaging in man. J Nucl Med 29: 796-807. Crouzel C, Guillaume M, Barre L, Lemaire C, Pike VW. 1992. Ligands and tracers for PET studies of the 5‐HT system— Current status. Nucl Med Biol‐Int J Rad Appl B 19: 857-870. Dannals RF, Ravert HT, Frost JJ, Wilson AA, Burns HD, et al. 1985. Radiosynthesis of an opiate receptor binging radiotracer: (11C) Carfentanil. Int J Appl Radiat Isot 36: 303-306. Dewey SL, Mac Gregor RR, Brodie JD, et al. 1990. Mapping muscarinic receptors in human and baboon brain using [N‐11C‐methyl]‐benztropine. Synapse 5: 213-223. Ding YS, Fowler JS, Volkow ND, Gatley SJ, Logan J, et al. 1994. Pharmacokinetics and in vivo specificity of [11C]DL‐threo‐ methylphenidate for the presynaptic dopaminergic neuron. Synapse 18(2): 152-160. Dolle F, Bottlaender M, Demphel S, et al. 2000. High efficient synthesis of [11C]PE2I, a selective radioliogand for the quantification of the dopamine transporter using PET. J Label Compd Radiopharm 43: 997-1004. Farde L, Ginovart N, Halldin C, Chou YH, Olsson H, et al. 2000. A PET study of [11C]b‐CIT‐FE binding to the dopamine transporter in the monkey and human brain. Int J Neuropsychopharm 2: 203-214. Farde L, Hall H, Ehrin E, Sedvall G. 1986. Quantitative analysis of dopamine D2 receptor binding in the human brain by PET. Science 231: 258-261. Farde L, Halldin C, Stone‐Elander S, Sedvall G. 1987. PET analysis of human dopamine receptor subtypes using 11 C‐SCH 23390 and 11C‐raclopride. Psychopharmacology 92: 278-284. Farde L, Ito H, Swahn C‐G, Pike VW, Halldin C. 1998. Quantitative analyses of carbonyl‐carbon‐11‐WAY‐100635 binding to central 5‐hydroxytryptamine‐1A receptors in man. J Nucl Med 39: 1965-1971. Finnema SJ, Seneca N, Farde L, Shchukin E, Sovago J, et al. 2005. A preliminary PET evaluation of the new dopamine D2 receptor agonist [11C]MNPA in cynomolgus monkey. Nucl Med Biol 32(4): 353-360. Fischman AJ, Bonab AA, Babich JW, Livni E, Alpert NM, et al. 2001. [11C, 127I] Altropane: A highly selective ligand for PET imaging of dopamine transporter sites. Synapse 39(4): 332-342. Fowler JS, Volkow ND, Wang GJ, Ding YS, Dewey SL. 1999. PET and drug research and development. J Nucl Med 40(7): 1154-1163. Fowler JS, Volkow ND, Wolf AP, Dewey SL, Schlyer DJ, et al. 1989. Mapping cocaine binding sites in human and baboon brain in vivo. Synapse4(4): 371-377. Frost JJ, Douglass KH, Mayberg HS, et al. 1989. Multicompartmental analysis of [11C]‐carfentanil binding to opiate

In vivo imaging of neurotransmitter systems with PET receptors in humans measured by positron emission tomography. J Cereb Blood Flow Metab 9: 398-409. Fujiwara T, Meija M, Itoh M, Anai K, Meguro K, et al. 1996. Quantitative imaging of [11C]benztropine in the human brain with graphic analysis and spectral analysis. Quantification of Brain Function Using PET. Myers R, Cunningham MG, Bailey D, Jones T, editors. San Diego: Academic Press. Fukumitsu N, Ishii K, Kimura Y, Oda K, Sasaki T, et al. 2003. Imaging of adenosine A1 receptors in the human brain by positron emission tomography with [11C]MPDX. Ann Nucl Med 17(6): 511-515. Fukumitsu N, Ishii K, Kimura Y, Oda K, Sasaki T, et al. 2005. Adenosine A1 receptor mapping of the human brain by PET with 8‐dicyclopropylmethyl‐1‐11C‐methyl‐3‐propylxanthine. J Nucl Med 46(1): 32-37. Gallezot JD, Bottlaender M, Gregoire MC, Roumenov D, Deverre JR, et al. 2004. In vivo imaging of human cerebral nicotinic acetylcholine receptors with 2‐18F‐fluoro‐A‐85380 and PET. J Nucl Med 46(2): 240-247. Gao M, Mock BH, Hutchins GD, Zheng QH. 2005. Synthesis and initial PET imaging of new potential dopamine D3 receptor radioligands (E)‐4,3,2‐[11C]methoxy‐N‐4‐(4‐(2‐methoxyphenyl)piperazin‐1‐yl)butyl‐cinnamoylamides. Bioorg Med Chem 13(22): 6233-6243. Gulya´s B, Halldin C, Vas A, Banati RB, Shchukin E, et al. 2005. [11C]Vinpocetine: A prospective peripheral benzodiazepine receptor ligand for primate PET studies. J Neurol Sci 229–230: 219-223. Halldin C, Farde L, Hogberg T, et al. 1991. A comparative PET‐study of five carbon‐11 or fluorine‐18 labelled salicylamides. Preparation and in vitro dopamine D2 receptor binding. Nucl Med Biol 18: 871-881. Halldin C, Farde L, Hogberg T, et al. 1995. Carbon‐11‐FLB 457: A radioligand for extrastriatal D2 dopamine receptors. J Nucl Med 36: 1275-1281. Halldin C, Farde L, Litton JE, Hall H, Sedvall G. 1992. (C‐11) Ro 15‐4513, a ligand for visualization of benzodiazepine receptor binding—Preparation, autoradiography and positron emission tomography. Psychopharmacology 108: 16-22. Halldin C, Farde L, Lundkvist C, et al. 1996. (11C)‐b‐CIT‐FE, a radioligand for quantitation of the dopamine transporter in the living brain using positron emission tomography. Synapse 22: 386-390. Halldin C, Foged C, Chou YH, et al. 1998a. Carbon‐11 NNC 112: A radioligand for PET examination of striatal and neocortical D1‐dopamine receptors. J Nucl Med 39: 2064-2068. Halldin C, Guilloteau D, Okubo Y, 1998b. PET‐characterization of [C‐11]PE2I binding to dopamine transporters in the monkey brain. J Nucl Med 39: 118.

3

Halldin C, Gulya´s B, Farde L. 2001. PET studies with carbon‐ 11 radioligands in neuropsychological drug development. Curr Radiopharm Design 7: 1907–1929. Halldin C, Gulya´s B, Farde L. 2004. PET for drug development. From Morphological Imaging to Molecular Targeting. Berlin Heidelberg: Springer Verlag; pp. 95–110. Halldin C, Gulya´s B, Langer O, Farde L. 2001. Brain radioligands—State of the art and new trends. Q J Nucl Med 2: 139-152. Halldin C, Ho¨gberg T. 1991. Radiotracers: Synthesis and use in imaging. A Textbook of Drug Design and Development. Kroogsgaard‐Larsen P, Liljefors T, Madsen U, editors. Amsterdam: Harwood; pp. 174–205. Halldin C, Lundberg J, Sovago J, Gulya´s B, Guilloteau D, et al. 2005. [11C]MADAM, a new serotonin transporter radioligand characterized in the monkey brain by PET. Synapse 58(3): 173-183. Halldin C, Stone‐Elander S, Farde L, et al. 1986. Preparation of 11C‐labelled SCH 23390 for the in vivo study of dopamine D1 receptors using positron emission tomography. Appl Radiat Isot‐Int J Rad A 37: 1039-1043. Hashimoto K, Inoue O, Suzuki K, Yamasaki T, Kojima M. 1989. Synthesis and evaluation of 11C‐PK 11195 for in vivo study of peripheral‐type benzodiazepine receptors using positron emission tomography. Ann Nucl Med 3(2): 63-71. Heiss WD, Herholz K. 2006. Brain receptor imaging. J Nucl Med 47(2): 302-312. Hietala J, Nyman MJ, Eskola O, Laakso A, Gronroos T, et al. 2005. Visualization and quantification of neurokinin‐1 (NK1) receptors in the human brain. Mol Imaging Biol 7(4): 262-272. Houle S, Ginovart N, Hussey D, Meyer JH, Wilson AA. 2000. Imaging the serotonin transporter with positron emission tomography: Initial human studies with [11C]DAPP and [11C]DASB. Eur J Nucl Med 27(11): 1719-1722. Huang Y, Narendran R, Bischoff F, Guo N, Zhu Z, et al. 2005. A positron emission tomography radioligand for the in vivo labeling of metabotropic glutamate 1 receptor: (3‐ethyl‐2‐[ 11C]methyl‐6‐quinolinyl)(cis‐4‐methoxycyclohexyl)methanone. J Med Chem 48(16): 5096-5099. Ito H, Nyberg S, Halldin C, Lundkvist C, Farde L. 1998. PET imaging of central 5‐HT2A receptors with carbon‐11‐MDL 100,907. J Nucl Med 39(1): 208-214. Iwabuchi K, Ito C, Tashiro M, Kato M, Kano M, et al. 2005. Histamine H1 receptors in schizophrenic patients measured by positron emission tomography. Eur Neuropsychopharmacol 15(2): 185-191. Jones AK, Cunningham VJ, Ha‐Kawa SK, Fujiwara T, Liyii Q, et al. 1994. Quantitation of [11C]diprenorphine cerebral kinetics in man acquired by PET using presaturation, pulse‐chase and tracer‐only protocols. J Neurosci Methods 51(2): 123-134.

97

98

3

In vivo imaging of neurotransmitter systems with PET

Jovkar S, Wienhard K, Pawlik G, Coenen HH. 1990. The quantitative analysis of D2‐dopamine receptors in baboon striatum in vivo with 3‐N‐[20 ‐18F]fluoroethylspiperone using positron emission tomography. J Cereb Blood Flow Metab 10(5): 720-726. Kim SE, Szabo´ Z, Seki C, Ravert HT, Scheffel U, et al. 1999. Effect of tracer metabolism on PET measurement of [11C] pyrilamine binding to histamine H1 receptors. Ann Nucl Med 13(2): 101-107. Kimura Y, Ishii K, Fukumitsu N, Oda K, Sasaki T, et al. 2004. Quantitative analysis of adenosine A1 receptors in human brain using positron emission tomography and [1‐methyl‐ 11C]8‐dicyclopropylmethyl‐1‐methyl‐3‐propylxanthine. Nucl Med Biol 31(8): 975-981. Laakso A, Bergman J, Haaparanta M, Vilkman H, Solin O, et al. 1998. [F‐18]CFT ([F‐18]WIN 35,428), a radioligand to study the dopamine transporter with PET: Characterization in human subjects. Synapse 28: 244-250. Lang L, Jagoda E, Schmall B, Vuong BK, Adams HR, et al. 1999. Development of fluorine‐18‐labeled 5‐HT1A antagonists. J Med Chem 42(9): 1576-1586. Lundberg J, Odano I, Olsson H, Halldin C, Farde L. 2005. Quantification of 11C‐MADAM binding to the serotonin transporter in the human brain. J Nucl Med 46(9): 1505-1515. Lundkvist C, Halldin C, Ginovart N, et al. 1996. [C‐11]MDL 100907, a radioligand for selective imaging of 5‐HT2A receptors with positron emission tomography. Life Sci 58: PL187-PL192. Lundkvist C, Halldin C, Ginovart N, Swahn C‐G, Farde L. 1997. [18F]b‐CIT‐FP is superior to [11C]b‐CIT‐FP for quantitation of the dopamine transporter. Nucl Med Biol 24: 621-627. Madras BK, Meltzer PC, Liang AY, Elmaleh DR, Babich J, et al. 1998. Altropane, a SPECT or PET imaging probe for dopamine neurons: I. Dopamine transporter binding in primate brain. Synapse 29: 93-104. Maeda J, Suhara T, Zhang MR, Okauchi T, Yasuno F, et al. 2004. Novel peripheral benzodiazepine receptor ligand [11C]DAA1106 for PET: An imaging tool for glial cells in the brain. Synapse 52(4): 283-291. Marchais‐Oberwinkler S, Nowicki B, Pike VW, Halldin C, Sandell J, et al. 2005. N‐oxide analogs of WAY‐100635: New high affinity 5‐HT1A receptor antagonists. Bioorg Med Chem 13(3): 883-893. Mayberg HS, Sadzot B, Meltzer CC, Fisher RS, Lesser RP, et al. 1991. Quantification of mu and non‐mu opiate receptors in temporal lobe epilepsy using positron emission tomography. Ann Neurol 30(1): 3-11. Mazie`re M, Hantraye P, Prenant C, Sastre J, Comar D. 1984. Synthesis of ethyl 8‐fluoro‐5,6‐dihydro‐5‐[11C]methyl‐6‐ oxo‐4H‐imidazo [1,5‐a] [1,4]benzodiazepine‐3‐carboxylate

(RO 15.1788‐11C): A specific radioligand for the in vivo study of central benzodiazepine receptors by positron emission tomography. Int J Appl Radiat Isot 35: 973-976. McConathy J, Owens MJ, Kilts CD, Malveaux EJ, Camp VM, et al. 2004. Synthesis and biological evaluation of [11C] talopram and [11C]talsupram: Candidate PET ligands for the norepinephrine transporter. Nucl Med Biol 31(6): 705-718. Meyer PT, Elmenhorst D, Matusch A, Winz O, Zilles K, et al. 2006. 18F‐CPFPX PET: On the generation of parametric images and the effect of scan duration. J Nucl Med 47(2): 200-207. Mukherjee J, Yang ZY, Brown T, Lew R, Wernick M, et al. 1999. Preliminary assessment of extrastriatal dopamine D-2 receptor binding in the rodent and nonhuman primate brains using the high affinity radioligand, 18F-fallypride. Nucl Med Biol 26: 519-527. Mulholland GK, Kilbourn MR, Sherman P, et al. 1995. Synthesis, in vivo biodistribution and dosimetry of [C‐11]N‐ methylpiperidyl benzilate ([C‐11]NMPB), a muscarinic acetylcholine receptor antagonist. Nucl Med Biol 22: 13-17. Mu¨ller L, Halldin C, Farde L, Karlsson P, Hall H, et al. 1993. [11C] b‐CIT, a cocaine analogue. Preparation, autoradiography and preliminary PET investigations. Nucl Med Biol 20(3): 249-255. Nordberg A, Hartvig P, Lundqvist H, Antoni G, Ulin J, et al. 1989. Uptake and regional distribution of (þ)‐(R)‐ and ()‐(S)‐N‐[methyl‐11C]‐nicotine in the brains of rhesus monkey. An attempt to study nicotinic receptors in vivo. J Neural Transm Park Dis Dement Sect 1(3): 195-205. Obrzut SL, Koren AO, Mandelkern MA, Brody AL, Hoh CK, et al. 2005. Whole‐body radiation dosimetry of 2‐[18F] Fluoro‐A‐85380 in human PET imaging studies. Nucl Med Biol 32(8): 869-874. Okuyama S, Chaki S, Yoshikawa R, Ogawa S, Suzuki Y, et al. 1999. Neuropharmacological profile of peripheral benzodiazepine receptor agonists, DAA1097 and DAA1106. Life Sci 64(16): 1455-1464. Olsson H, Halldin C, Swahn C‐G, Farde L. 1993. Quantification of [C‐11]FLB 457 binding to extrastriatal dopamine receptors in the human brain. J Cerebr Blood Flow Metab 19: 1164-1173. Persson A, Pauli S, Halldin C, et al. 1989. Saturation analysis of specific 11C‐Ro 15–1788 binding to the human neocortex using positron emission tomography. Hum Psychopharmacol 4: 21-31. Pike VW, Halldin C, Crouzel C, et al. 1993. Radioligands for PET studies of central benzodiazepine receptors and PK (peripheral benzodiazepine) binding sites—Current status. Nucl Med Biol 20: 503-525. Pike VW, Halldin C, McCarron JA, Lundkvist C, Hirani E, et al. 1998. [Carbonyl‐11C]Desmethyl‐WAY‐100635 (DWAY)

In vivo imaging of neurotransmitter systems with PET is a potent and selective radioligand for central 5‐HT1A receptors in vitro and in vivo. Eur J Nucl Med 25(4): 338-346. Pike VW, Mccarron JA, Lammertsma AA, et al. 1996. Exquisite delineation of 5‐HT1A receptors in human brain with PET and [carbonyl‐C‐11]WAY‐100635. Eur J Pharmacol 301: R5-R7. Sandell J, Halldin C, Chou YH, Swahn CG, Thorberg SO, et al. 2002. PET‐examination and metabolite evaluation in monkey of [11C]NAD‐299, a radioligand for visualisation of the 5‐HT1A receptor. Nucl Med Biol 29(1): 39-45. Sandell J, Halldin C, Hall H, et al. 1999. Radiosynthesis and autoradiographic evaluation of [C‐11]NAD‐299, a radioligand for visualization of the 5‐HT1A receptor. Nucl Med Biol 26: 159-164. Schou M, Halldin C, Pike VW, Mozley PD, Dobson D, et al. 2005. Post‐mortem human brain autoradiography of the norepinephrine transporter using (S,S)‐[18F]FMeNER‐D2. Eur Neuropsychopharmacol 15(5): 517-520. Schou M, Halldin C, Sovago J, Pike VW, Gulyas B, et al. 2003. Specific in vivo binding to the norepinephrine transporter demonstrated with the PET radioligand, (S,S)‐[11C] MeNER. Nucl Med Biol 30(7): 707-714. Schou M, Halldin C, Sovago J, Pike VW, Hall H, et al. 2004. PET evaluation of novel radiofluorinated reboxetine analogs as norepinephrine transporter probes in the monkey brain. Synapse 53(2): 57-67. Schou M, So´va´go´ J, Pike VW, Gulya´s B, Bogeso KP, et al. 2006. Synthesis and positron emission tomography evaluation of three norepinephrine transporter radioligands: [C‐11]desipramine, [C‐11]talopram and [C‐11]talsupram. Mol Imaging Biol 8(1): 1-8. Seneca N, Andree B, Sjoholm N, Schou M, Pauli S, et al. 2005. Whole‐body biodistribution, radiation dosimetry estimates for the PET norepinephrine transporter probe (S,S)‐[18F] FMeNER‐D2 in non‐human primates. Nucl Med Commun 26(8): 695-700. Sobrio F, Debruyne D, Dhilly M, Chazalviel L, Camsonne R, et al. 2005. Evaluation in rats and primates of [11C]‐mecamylamine, a potential nicotinic acetylcholine receptor radioligand for positron emission tomography. Neurochem Int 46(6): 479-488. Solin O, Eskola O, Hamill TG, Bergman J, Lehikoinen P, et al. 2004. Synthesis and characterization of a potent, selective, radiolabeled substance‐P antagonist for NK1 receptor quantitation: ([18F]SPA‐RQ). Mol Imaging Biol 6(6): 373-384. So´va´go´ J, Farde L, Halldin C, Langer O, Laszlovszky I, et al. 2004. Positron emission tomographic evaluation of the putative dopamine‐D3 receptor ligand, [11C]RGH‐1756 in the monkey brain. Neurochem Int 45(5): 609-617. So´va´go´ J, Farde L, Halldin C, Schukin E, Schou M, et al. 2005. Lack of effect of reserpine‐induced dopamine depletion on

3

the binding of the dopamine‐D3 selective radioligand, [11C] RGH‐1756. Brain Res Bull 67(3): 219-224. Szabo´ Z, Kao PF, Mathews WB, Ravert HT, Musachio JL, et al. 1996. Positron emission tomography of 5‐HT reuptake sites in the human brain with C‐11 McN5652 extraction of characteristic images by artificial neural network analysis. Behav Brain Res 73(1–2): 221-224. Szabo´ Z, Ravert HT, Gozukara I, Geckle W, Seki C, et al. 1993. Noncompartmental and compartmental modeling of the kinetics of carbon‐11 labeled pyrilamine in the human brain. Synapse 15(4): 263-275. Szabo´ Z, Scheffel U, Suehiro M, Dannals RF, Kim SE, et al. 1995. Positron emission tomography of 5‐HT transporter sites in the baboon brain with [11C]McN5652. J Cereb Blood Flow Metab 15(5): 798-805. Takahashi K, Murakami M, Miura S, Iida H, Kanno I, et al. 1999. Synthesis and autoradiographic localization of muscarinic cholinergic antagonist (þ)N‐[11C]methyl‐3piperidyl benzylate as a potent radioligand for positron emission tomography. Appl Radiat Isot 5: 521-525. Tashiro M, Sakurada Y, Iwabuchi K, Mochizuki H, Kato M, et al. 2004. Central effects of fexofenadine and cetirizine: Measurement of psychomotor performance, subjective sleepiness, and brain histamine H1‐receptor occupancy using 11C‐doxepin positron emission tomography. J Clin Pharmacol 44(8): 890-900. Theodore WH, Carson RE, Andreasen P, et al. 1992. PET imaging of opiate receptor binding in human epilepsy using [F‐18]cyclofoxy. Epilepsy Res 13: 129-139. Tsukada H, Nishiyama S, Kakiuchi T, Ohba H, Sato K, et al. 2001a. Ketamine alters the availability of striatal dopamine transporter as measured by [11C]b‐CFT and [11C]b‐CIT‐FE in the monkey brain. Synapse 42(4): 273-280. Tsukada H, Takahashi K, Miura S, Nishiyama S, Kakiuchi T, 2001b. Evaluation of novel PET ligands (þ)N‐[11C]methyl‐ 3‐piperidyl benzilate ([11C](þ)3‐MPB) and its stereoisomer [11C](–)3‐MPB for muscarinic cholinergic receptors in the conscious monkey brain: A PET study in comparison with. Synapse 39(2): 182-192. Van Dort ME, Kim JH, Tluczek L, Wieland DM. 1997. Synthesis of 11C‐labeled desipramine and its metabolite 2‐hydroxydesipramine: Potential radiotracers for PET studies of the norepinephrine transporter. Nucl Med Biol 24(8): 707-711. Vas A, Gulya´s B. 2005. Eburnamine derivatives and the brain. Med Res Rev 25(6): 737-757. Volkow ND, Fowler JS, Ding YS, Wang GJ, Gatley SJ. 1998. Positron emission tomography radioligands for dopamine transporters and studies in human and nonhuman primates. Adv Pharmacol 42: 211-214. Volkow ND, Fowler JS, Ding YS, Wang GJ, Gatley SJ. 1999. Imaging the neurochemistry of nicotine actions: Studies

99

100

3

In vivo imaging of neurotransmitter systems with PET

with positron emission tomography. Nicotine Tob Res 1: S127-S132. Wagner HN, Burns HD, Dannals RF, et al. 1983. Imaging dopamine receptors in the human brain by positron tomography. Science 221: 1264-1266. Wienhard K, Coenen HH, Pawlik G, Rudolf J, Laufer P, et al. 1990. PET studies of dopamine receptor distribution using [18F]fluoroethylspiperone: Findings in disorders related to the dopaminergic system. J Neural Transm Gen Sect 81(3): 195-213. Wilson AA, Ginovart N, Hussey D, Meyer J, Houle S. 2002. In vitro and in vivo characterisation of [11C]‐DASB: A probe

for in vivo measurements of the serotonin transporter by positron emission tomography. Nucl Med Biol 29(5): 509-515. Yu M, Tueckmantel W, Wang X, Zhu A, Kozikowski AP, et al. 2005. Methoxyphenylethynyl, methoxypyridylethynyl and phenylethynyl derivatives of pyridine: Synthesis, radiolabeling and evaluation of new PET ligands for metabotropic glutamate subtype 5 receptors. Nucl Med Biol 32(6): 631-640. Zubieta JK, Koeppe RA, Frey KA, Kilbourn MR, Mangner T, et al. 2001. Assessment of muscarinic receptor concentrations in aging and Alzheimer disease with [11C]NMPB and PET. Synapse 39: 275-287.

4

Synaptic and Nonsynaptic Release of Transmitters

E. S. Vizi . B. Lendvai

1

Historical Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102

2 Communication Between Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 2.1 Synaptic Interaction Between Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 2.2 Nonsynaptic Interaction Between Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 3 Release of Transmitters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 3.1 Nonsynaptic Release of Transmitter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 3.2 Spillover of Transmitters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 4 4.1 4.2 4.3 4.4

What Influences the Concentration of Transmitter in the Extracellular Space? . . . . . . . . . . . . . . . 106 Amount of Transmitter Released . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 Uptake of Transmitters by Plasma Membrane Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 Volume of Extracellular Space . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107 Effect of Drugs on Targets Located Intrasynaptically and Extrasynaptically: Law of Mass Action . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107

5

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108

#

2008 Springer ScienceþBusiness Media, LLC.

102

4

Synaptic and nonsynaptic release of transmitters

Abstract: Nonclassical receptorial functions represent revolutionary possibilities at the cellular level for some less-nderstood features of neural and cerebral activities. Although different forms of nonsynaptic communication often appear in different studies, their difference from synaptic actions is generally not recognized. The corner stones of interneuronal nonsynaptic communication include the release of transmitters into the extracellular space and the extrasynaptic receptors and transporters. Transmitters can be released from nonsynaptic varicosities without being coupled to frequency‐coded neuronal activity and from synapses following high presynaptic activity via spillover. The released substances are able to diffuse over large distances to reach remote tissue. Extrasynaptic receptors may occur at all possible membrane surfaces in various systems. These receptors are of high affinity, providing targets for low-dose drugs in many instances of medical therapy. List of Abbreviations: ACh, acetylcholine; AMPA, alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid; CNS, central nervous system; DA, dopamine; 5-HT, serotonin; GABA, gamma-aminobutyric acid; NA, noradrenaline; NMDA, N-methyl-D-aspartate; nAChR, nicotinic acetylcholine receptor; VTA, ventral tegmental area

1

Historical Background

Of all the cells in the body, only nerve cells are able to communicate regularly with one another. In the nineteenth century, it was believed that ‘‘nerve centers’’ were made up of a continuous intermediary network between the motor nerves and the sensitive and sensory nerves (Cajal, 1937). Ramon y Cajal in his experiments applied Golgi staining to discover that neurons are independent units, and not entities fused to each other. This was an important step that changed the way of thinking of many scientists. Ramon y Cajal received the Nobel Prize in 1906 for this discovery. The question arose of how neurons communicate, if they do not fuse to one another with anastomosis, forming large neural nets of an intermediary network. Bernard demonstrated the dissociation of nerve and muscle activity by curarizing frogs (Bernard, 1857). This is regarded as one of the key experiments in the development of the concept of chemical transmission. The idea that nerves are able to release chemicals to communicate with other cells was first explicitly proposed for sympathetic nerves when Elliott, a young medical student in Cambridge, suggested in 1904, ‘‘adrenalin might. . .be the chemical stimulant liberated on each occasion when the impulse arrives at the periphery.’’ This brilliant hypothesis was confirmed by Loewi (1921), who showed that the stimulation of sympathetic nerves in frog heart is mediated by Acceleransstoff (adrenaline). The observation that the action of adrenaline and sympathetic stimulation are similar (Elliot, 1905) was further supported by the finding that the action of acetylcholine (ACh) and parasympathetic stimulation are also similar (Loewi, 1922; Dale, 1956). In spite of strong evidence suggesting otherwise, the alternative view that transmission is electrical enjoyed rather wide support during the first half of the twentieth century. This was mainly due to the fact that the electrical properties of what is now called conduction and transmission were seen to be similar (Erlanger, 1939; Gasser, 1939; Eccles, 1946). This belief gradually gained currency; a large number of neuroscientists believed that findings obtained in the neuromuscular junction were relevant not only to autonomic, but also to central synaptic transmission. Eccles in 1946 wrote ‘‘The original hypothesis was made as general as possible by applying it to the neuromuscular functions of skeletal muscle, and to the synapses of the sympathetic ganglia as well as of the central nervous system.’’ In fact, Eccles believed that the primary transmitter was electrical, but that chemical transmitters could be responsible only for slower and longer responses, detectable as a tail to the transmitter action on the postsynaptic site. This assumption was accepted almost universally by scientific society. Although different scientists (Dale, Feldberg, Kuffler, Uvna¨s, etc.) provided rather strong evidence to support and confirm the hypothesis of chemical transmission, Eccles resisted until 1948, when he acknowledged that even the fast response is due to acetylcholine at the neuromuscular junction and accepted the idea of chemical transmission. After his Pauline conversion from electrical to chemical transmission, the community of neuroscientists generally accepted the theory of chemical transmission for example, the communication between nerves and between nerve endings and target cells is chemical. It means that chemicals are released from nerve terminals in response to electrical depolarization followed by Ca2þ‐influx transmit messages between pre‐ and postsynaptic sites.

Synaptic and nonsynaptic release of transmitters

2

4

Communication Between Cells

2.1 Synaptic Interaction Between Cells From Sherrington’s classical work on ‘‘Integrative action of the nervous system,’’ it has been generally accepted that the synapse, the ‘‘surface of separation’’ between neurons, is the primary site of neuronal information processing. Transmitter is released into the synaptic cleft in quantal packages. The average neuron forms about 1,000 synapses and it receives about 10,000 inputs. Since the human brain contains about 1011 nerve cells, it means that in the brain there are at least 1014 synapses for information processing. The generally accepted form of chemical communication between nerves and between nerve endings and target cells is that the transmitter is released into the synaptic cleft in quantal packages as a result of action potentials arriving at the terminals. The transmitter acts on receptors located on the postsynaptic site and either opens or closes ion channels, thereby establishing chemical communication between pre‐ and postsynaptic sites. The effect of transmitters is terminated by either enzymatic degradation (e.g., in case of ACh) or by active reuptake into nerve terminals by transporters (cf. Amara and Kuhar, 1993; Raiteri et al., 2002). Nevertheless, our current knowledge of how information is conveyed chemically from one cell to another is derived from and heavily influenced by the textbook data regarding the neuromuscular junction (cf. Katz, 1969), where the transmitter is released in quanta. This system is adopted for very fast signaling; the information transfer occurs within millisecond time intervals and is able to transmit messages at a rate of several hundred impulses per second.

2.2 Nonsynaptic Interaction Between Neurons In addition to transmitter substances acting at close range in chemical synaptic neurotransmission, chemical interaction exists and information processing occurs between neurons and between neurons and target cells without any close synaptic contact; there is a nonsynaptic communication system which operates over some distance in the extracellular spaces (cf. Vizi, 1974, 1979, 1980, 1984, 2000; Vizi et al., 1985; Agnati et al., 1986, 1995; Fuxe and Agnati, 1991; Bach‐y‐Rita, 1993; Vizi and Kiss, 1998). In the past few years, several neurochemical, anatomical, pharmacological, and neurophysiological observations have been made which suggest that chemical interaction between cells does not only take place across the synaptic gap between pre‐ and postsynaptic membranes but may also occur in the absence of such specialized contacts, i.e., nonsynaptically (> Figure 4-1). Neurochemical evidence has been obtained suggesting that noradrenaline (NA) released from axon terminals, which do not make synaptic contact with cholinergic terminals in the gut (Furness and Costa, 1974; Gordon‐Weeks, 1982), inhibits the release of acetylcholine from cholinergic varicosities of the Auerbach plexus (Vizi, 1968; Paton and Vizi, 1969; Knoll and Vizi, 1970; Vizi and Knoll, 1971). A very similar observation was first made in the cerebral cortex (Vizi, 1974, 1979, 1980) where the majority of noradrenergic varicosities do not make synaptic contacts (Descarries et al., 1977). This new concept of information processing, now known as nonsynaptic chemical transmission (Vizi, 1980, 1984), has been shown to be a rule rather than an exemption in the CNS (central nervous system) and has gained widespread acceptance (since 1986, it has also been called volume transmission, Agnati et al., 1986; Fuxe and Agnati, 1991; Agnati et al., 1995, paracrine release, spillover, nonconventional release, etc). Compelling neurochemical, functional, and pharmacological evidence (cf. Vizi, 2000) has accumulated suggesting that transmitters released from axon terminals are able to diffuse far away from the release site and have an effect on receptors located nonsynaptically.

3

Release of Transmitters

3.1 Nonsynaptic Release of Transmitter The idea that transmitters can be released from nonsynaptic areas was first suggested for transmitters in the autonomic nervous system, where the axon terminals rarely make synaptic contact with the target cells.

103

104

4

Synaptic and nonsynaptic release of transmitters

. Figure 4-1 Nonsynaptic chemical transmission. A proportional diagram of two synaptic clefts: a typical synapse with a gap of 20 nm and a free axon terminal with its remote target cell (e.g., the vegetative nervous system). Note the difference in volume. Let us assume that both transmission sites are cholinergic. The average diameter of a vesicle containing acetylcholine (ACh) is 50 nm, therefore its volume is about 65,000 nm3 and its ACh concentration is about 0.1 M (cf. Marchbanks, 1979). If we assume that its content is completely discharged into a small synaptic cleft whose volume is about 200,000 nm3 (20 x 100 x 100 nm), the final concentration of ACh in the cleft is 30 mM (0.1/(200,000/65,000)), which is an extremely high concentration. Let us suppose that, for example, in Auerbach’s plexus, where the target smooth muscle cell is far from the varicose axon terminals (100–1,000 nm), only one vesicle is released. The volume in which the ACh released is 109 nm3 (1,000 x 1,000 x 1,000), i.e., a volume 104 times larger than that of a vesicle. Therefore, the ACh released from the vesicle is diluted by a factor of 10,000. If the cholinesterase is not active, the final concentration of ACh which reaches the muscarinic receptors of the smooth muscle is about 10
4 M. In this calculation, the cytoplasmic release has not been taken into account. Note the concentration of ACh in the cleft

Since then, a large body of evidence has shown that transmitters/modulators can be released from regions other than the nerve‐ending. Electron microscope and histochemical studies of the relationship between nerve terminals and target cells have shown that there are wide varieties of normal distances. The minimum width of the cleft between nerve varicosities and effector cells varies considerably in different tissues. In the vas deferens and sphincter pupillae, the separation is about 15–20nm. In blood vessels, the smallest space between varicosities in the perivascular plexus at the advential‐medial border and smooth muscle cells varies from about 50 nm to 2 mm (small muscular arteries, large arterioles, large elastic arteries) (see Burnstock, 1979). In these cases, no postjunctional specializations have been found with any consistency for wider neuromuscular junctions. Release from axonal varicosities devoid of synaptic membrane specialization has recently been suggested to be the function of the central monoamine terminals (Descarries et al., 1977; Beaudet and Descarries, 1978). Beaudet and Descarries (1978) claimed that the release of biogenic amines solely from varicosities making synaptic contact could hardly account for the total amount released from axon terminals. As the number of locus coeruleus cells in the rat is only about 1,400 (Descarries and Saucier, 1972) and the density of noradrenergic varicosities is 2 million/mm3, each cell body has an average of about 140,000 varicosities in the hippocampus. It means that the excitatory inputs to a noradrenergic cell body might activate a neuron whose transmitter, noradrenaline, released from varicosities might control a rather large field. It also means that a transmitter or modulator released from nonsynaptic varicosities could affect very large neuronal assemblies.

Synaptic and nonsynaptic release of transmitters

4

There is an interesting difference in the localization of receptors on effector cells where the transmitter is released into a small synaptic gap compared with those where it is released into a large extraneuronal space. In the former case, there is a small area of the cell where the receptors are concentrated. At the neuromuscular junction, for example the extrasynaptic area is relatively insensitive to the transmitter, because only a few receptors are there. However, when the transmitter release site and the target cells (e.g., Auerbach’s plexus varicose axon terminals and smooth muscle cells) are widely separated from each other (100–1,000 nm), there is no specific subsynaptic arrangement, and the receptors are evenly distributed along the whole surface of the smooth muscle cell. This morphological arrangement accommodates any type of diffusion‐mediated transmission, where the advantages of quantal release cannot be used. In the CNS, there are large amount of receptors located extrasynaptically and they are of high affinity. Even the cholinergic neurons fail to make synaptic contact in the hippocampus. Jones and Wonnacott (2004) provided evidence that in the ventral tegmental area (VTA), 27% of presynaptic a7 nAChRs (nicotinic acetylcholine receptors) are located extrasynaptically. The absence of axo‐axonic synapses (Descarries et al., 1997), i.e., direct cholinergic synaptic input to presynaptic a7 nAChRs, indicates that these receptors are likely to be activated by choline or ACh released from cholinergic varicosities (boutons) that are far away. Electron microscopy studies revealed that cholinergic varicosities in the hippocampal CA1 region are largely (93%) nonsynaptic compared with another transmitter system (Umbriaco et al., 1995), e.g., GABAergic and glutamatergic neurons, which make exclusively synaptic contacts. The nonsynaptic control of chemical neurotransmission by different modulators released from axonal varicosities lacking junctions might play a physiological role both in the CNS and in the neurovegetative system in shaping emotion, behavior, or learning processes, or in controlling the balance between the parasympathetic and the sympathetic nervous systems.

3.2 Spillover of Transmitters Though glutamate is the major excitatory transmitter, GABA is the most important inhibitory transmitter in the brain and spinal cord. Both glutamatergic and GABAergic terminals make exclusively synaptic contacts with other neurons. In this respect, they are different from monoaminergic nerve terminals which in the majority do not make synaptic contact (cf. Vizi, 2000). In recent years, it has become increasingly clear that receptors sensitive to glutamate and GABA besides their subsynaptic localization, they are also expressed extrasynaptically. The question that arises is where glutamate and GABA come from to signal these extrasynaptic receptors if they are only released into the synaptic cleft. The plausibility of the spillover (> Figure 4-2) depends on how much glutamate or GABA is released and how easily it can diffuse out of the synaptic cleft and how transporters terminate it to diffuse away. Glutamate, for example, the major excitatory transmitter of the brain, participates mainly in synaptic interactions between glutamatergic release sites predominantly located within synapses (Umbriaco et al., 1995) and AMPA and NMDA (N‐methyl‐D‐aspartate) receptors. Some synaptic spillover of glutamate has been observed and its effect on extrasynaptic NMDA (but not on AMPA) receptors was shown (Asztely et al., 1997; Kullmann and Asztely, 1998; Semyanov and Kullmann, 2000). The probability of spillover depends on how much glutamate is released, how easily it can diffuse out of the synaptic cleft, and how intra‐ and extrasynaptic transporters terminate it to diffuse away. The diffusion of glutamate away from synapses is therefore very limited because of effective neuronal and glial uptake processes. As far as the functional role of spillover is concerned, it has been shown (Mitchell and Silver, 2000) that spillover of glutamate released from excitatory mossy fibers is able to inhibit GABA release from neighboring Golgi cell terminals by activating presynaptic mGluRs. This heteroreceptor‐mediated inhibition of inhibitory fibers, in fact, boosts the efficacy of excitatory fibers. Kaneda and colleagues (1995) showed in cerebellar granule cells using voltage‐clamp experiments that in response to application of GABAA receptor antagonists there is a reduction in the ‘‘holding current.’’ A similar observation was made by Nusser and Mody (2002) on granule cells of the dentate gyrus. The inhibitory role of ambient GABA concentration was also shown (Semyanov et al., 2003, 2004) in hippocampal interneurons of CA1 region of the hippocampus, but interestingly not in adult pyramidal cells of hippocampus (Demarque et al., 2002). Nevertheless, there is a significant tonic GABAA

105

106

4

Synaptic and nonsynaptic release of transmitters

. Figure 4-2 Spillover of synaptic transmitters in the central nervous system. In this scheme, a presynaptic axon terminal (dark gray) makes a synapse on a dendritic spine (light gray). Transmitters (black dots) in the vesicles are released into the synaptic cleft where it reaches the synaptic receptors (pentameric structures in this case). Arrows indicate the spillover of the released transmitters from the synapse

receptor mediated current in pyramidal cells when the GABA uptake is inhibited (Bai et al., 2001). During development, the tonic activation of GABAA receptors is induced by GABA released in a [Ca2þ]o‐dependent way, but in adult rats the tonic activation of GABAA receptors is produced by nonvesicular transmitter release (Rossi et al., 2003). This fact indicates that high‐affinity GABAA receptors are activated by extrasynaptic ambient GABA concentrations (> Figure 4-2).

4

What Influences the Concentration of Transmitter in the Extracellular Space?

4.1 Amount of Transmitter Released The amount of transmitter released is determined by the amplitude of the depolarization of the nerve terminals, which, in turn, is determined by the number and frequency of the action potentials in the axons. Transmitters are released in the form of packets, in quanta, that correspond to synaptic vesicles. There seems to be no evidence that quantal release can occur solely at varicosities with synaptic contact. In the varicosities of nerves, transmitter release occurs intermittently following the stimulation of the parent axon (Cunnane and Stja¨rne, 1982; Blakely et al., 1986) and is facilitated by high‐frequency train stimulation (Cunnane and Stja¨rne, 1984). There is a low probability of release in any varicosity invaded by a nerve action potential (Cunnane and Stja¨rne, 1982). The intermittency might be due to a failure of conduction of nerve action potentials within the varicose terminals (Stja¨rne, 1978; Morita and North, 1981) so that large parts of the distal region of arborization could be intermittently excluded from transmitter secretion. In places where the gap is large, where there is no synaptic specialization, or where the transmitter must cross distances of micrometers to reach the target cell, the transmitter released from the cytoplasm also plays a critical role in chemical transmission (Vizi et al., 1982). In fact, there is a large body of available evidence showing that the release of transmitter of cytoplasmic origin is also involved (carrier‐mediated release).

4.2 Uptake of Transmitters by Plasma Membrane Transporters Once a transmitter is released into extrasynaptic space, its effect on receptors is terminated by its reuptake into the surrounding nerve terminals and glia, a process mediated by plasma membrane transporters. These nonsynaptic transporters also terminate the overspill of the synaptically released transmitter, and thereby they play an important role in influencing the concentration of transmitters in the extraneuronal space. Monoamine uptake carriers belong to the family of Naþ/Cl
‐dependent membrane transporters containing

Synaptic and nonsynaptic release of transmitters

4

12 transmembrane domains (Amara and Arriza, 1993). The cloning and sequencing of monoamine transporters in the early 1990s (Blakely et al., 1991; Giros et al., 1991, 1992; Pacholczyk et al., 1991; Ramamoorthy et al., 1993) revealed that these proteins show a very high degree of structural homology. Extrasynaptic glutamate spillover was shown in the hippocampus (Asztely et al., 1997). It was also shown that glutamate transporters play a critical role in terminating the nonsynaptic diffusion of glutamate, thereby limiting cross talk between neighboring excitatory synapses. The activity of transporters is temperature‐ dependent (Amara and Arriza, 1993). Voltammetry and microdialysis techniques provided temporally resolved information concerning the concentration of transmitters in the extrasynaptic space. It turns out that in clinical practice, the chemicals are able to reach 0.1–23 mM in the extraneuronal space of the brain (> Table 4-1). . Table 4-1 A few examples of presynaptic autoreceptors and heteroreceptors able to inhibit or facilitate the release of neurotransmitters Neurotransmitter Acetylcholine Noradrenaline Dopamine 5‐HT GABA Glutamate

Inhibitory autoreceptor M2 a2A/D

Facilitatory autoreceptor nAChR b2

D2/D3 5‐HT1D GABAB Metabotropic, CB1

– 5‐HT – –

Inhibitory heteroreceptors a2, D2/D3, 5‐HT1B Opiate, H3, M2, D2, PGE2, GABAB M2 a2 GABAB, CB1, M2 –

Facilitatory heteroreceptors NMDA, nAChR Angiotensin II, nAChR, NMDA, GABAA, P2x7 nAChR, NMDA – – nAChR

Abbreviations: M2, muscarinic acetylcholine receptor; nAChR, nicotinic acetylcholine receptor; NMDA, N‐methyl‐D‐aspartate; PG, Prostaglandin. For literature, see Vizi and Kiss (1998), Starke (2001), Go¨bel et al. (2000)

The high degree of homology may explain the accumulating observations that functional segregation of monoaminergic pathways is not as marked as it was assumed previously. Accumulating evidence indicates the promiscuity of nonsynaptically located monoamine transporters. Using selective uptake blockers and specific pathway lesions, it was proved that (i) [3H]DA could be taken up by noradrenergic and serotonergic neurons (Descarries et al., 1987), (ii) dopaminergic terminals take up and release [3H]5‐HT in the striatum, (iii) serotonergic varicosities take up and release [3H]DA in the hippocampus of rabbit (Feuerstein et al., 1986), (iv) serotonergic transporters take up NA, and (v) serotonergic varicosities can release NA (Vizi et al., 2004). These findings may provide a better understanding of the functional properties of monoaminergic systems and the mechanism of action of antidepressant drugs.

4.3 Volume of Extracellular Space It has been shown that all the transmitters in the CNS are present in the extracellular space, which is about 12– 25% of the brain volume (Nicholson, 1985). This space has been called ‘‘communication channel’’ (Nicholson et al., 1979), because the migration of chemical signals by diffusion plays a very important role in nonsynaptic transmission. Tortuosity and volume fraction of the extracellular space modifies the diffusion (Nicholson, 2005).

4.4 Effect of Drugs on Targets Located Intrasynaptically and Extrasynaptically: Law of Mass Action According to pharmacological textbooks, most drugs produce their effects by binding to protein molecules (receptors, transporter molecules, enzymes, and ion channels). It is generally accepted that the magnitude of the biological response produced by an endogenous ligand is related to the number of receptors (target proteins)

107

108

4

Synaptic and nonsynaptic release of transmitters

occupied (Rang, 2006); the receptor can bind only one drug molecule at a time. The first step in the action of drugs on target proteins (receptors) is the formation of a reversible drug‐target protein (receptor) complex. AðdrugÞ þ Rðfree receptorÞ ! ARðcomplexÞ: This step is governed by the Law of Mass Action; therefore, the actual concentrations of agonist (endogenous ligand, e.g., noradrenaline) and antagonist (e.g., drug applied) and the affinity of target proteins (receptors) play very important roles in the effect. Suppose that the intrasynaptic concentration of transmitter released into the synaptic cleft is between 1 and 10 mM, and that the orally administered drug (e.g., 10 mg/70 kg) can reach a concentration of about 0.05–5 mM in the extracellular space (> Table 4-2).

. Table 4-2 Concentration of drugs in the extracellular space Concentration (mM) Drug

Plasma

Nicotine Imipramine Citalopram Desmethylimipramine Fluoxetine

0.4–4.5 1–2 1 2 1

Cerebrospinal fluid

0.1

References Zevin et al. (1998) Besret et al. (1996) Hyttel (1982) Muscettola et al. (1978) Pato et al. (1991)

It is expected that a similar concentration would be found in the synapse. Therefore, taking into account the Law of Mass Action, the drug effect is marginal, if any. Therefore, the site of action of most drugs is those binding proteins (receptors, transporters, ion channels, and enzymes), which are located nonsynaptically, i.e., outside the synapse, in the extracellular space where the receptors and transporters are of high affinity (Vizi, 2000).

5

Conclusions

The nonsynaptic control of chemical neurotransmission by different modulators released from axonal varicosities lacking junctions might play a physiological role both in the CNS and in the neurovegetative system in shaping emotion, behavior, or learning processes, or in controlling the balance between the parasympathetic and sympathetic nervous system. Are the receptors located outside the postsynaptic density of the synapse functionally a part of chemical transmission, or are they promiscuous and accessible to chemicals released from different boutons with (in case of spillover) and without synaptic arrangements? The answer is yes. Presynaptic release modulating receptors represent suitable targets for pharmacological intervention by exogenous compounds acting as agonists, partial agonists, or antagonists. It is thus possible that presynaptic release modulating autoreceptors and heteroreceptors (> Figure 4-3) of high affinity may become the target of action for a new generation of drugs which can produce the desired therapeutic actions through the modulation or fine tuning of the release of neurotransmitters or cotransmitters. This novel mechanism differs from the well‐ established approach of using agonists or antagonists to directly stimulate or block postsynaptic receptors. An important way to control the activation of nonsynaptic receptors is to regulate the levels of transmitters in the extracellular space. Plasma membrane transporters play a very important role in terminating the levels of the transmitters released into the extraneuronal space. Clinically applied antidepressants reaching concentrations of 0.5–13 mM in the extraneuronal space may exert their effects on high‐affinity nonsynaptic transporters (Vizi, 2000). Because of the major impact of tonic inhibition or stimulation by endogenous transmitters on neuronal activity, this form of influence, besides its physiological importance, could be an

Synaptic and nonsynaptic release of transmitters

4

. Figure 4-3 Role of autoreceptors and heteroreceptors in modulation of transmitter release evoked by neuronal activity. This type of release is [Ca2+]‐dependent. Heteroreceptor is sensitive to a transmitter which is not produced by the neuron on which the receptor is expressed. Autoreceptors are the receptors sensitive to the neurons’ own transmitter substance. Scheme shows an example: noradrenaline (NA) inhibits its own release via the stimulation of presynaptic a2‐autoreceptors. Noradrenaline inhibits the release of acetylcholine (ACh) via the activation of presynaptic a2‐heteroreceptors. The effect of NA is terminated by its reuptake

important novel pharmacological target for the treatment of a wide range of disorders. Nonsynaptic and high‐ affinity GABAA receptors responsible for the tonic inhibitory conductance may be of clinical importance as targets for anesthetics and sedative drugs (Bai et al., 2001). It seems very likely that GABA and glutamate receptors need not be restricted to synapses to serve physiological functions.

References Agnati LF, Zoli M, Stromberg I, Fuxe K. 1995. Intercellular communication in the brain: Wiring versus volume transmission. Neuroscience 69: 711-726. Agnati LF, Fuxe K, Zoli M, Ozini I, Toffano G, et al. 1986. A correlation analysis of the regional distribution of central enkephalin and beta‐endorphin immunoreactive terminals and of opiate receptors in adult and old male rats. Evidence for the existence of two main types of communication in the central nervous system: The volume transmission and the wiring transmission. Acta Physiol Scand 128: 201-207. Amara SG, Arriza JL. 1993. Neurotransmitter transporters three distinct gene families. Curr Opin Neurobiol 3: 337-344. Amara SG, Kuhar MJ. 1993. Neurotransmitters: Recent progress. Annu Res Neurosci 16: 73-93. Asztely F, Erdemli G, Kullmann DM. 1997. Extrasynaptic glutamate spillover in the hippocampus: Dependence on temperature and the role of active glutamate uptake. Neuron 18: 281-293. Bach‐y‐Rita P. 1993. Neurotransmission in the brain by diffusion through the extracellular fluid: A review. Neuroreport 4: 343-350. Bai D, Zhu G, Pennefather P, Jackson MF, MacDonald JF, Orser BA. 2001. Distinct functional and pharmacological properties of tonic and quantal inhibitory postsynaptic currents mediated by gamma‐aminobutyric acid (A)

receptors in hippocampal neurons. Mol Pharmacol 59: 814-824. Beaudet A, Descarries L. 1978. The monoamine innervation of rat cerebral cortex: Synaptic and non‐synaptic axon terminals. Neuroscience 3: 851-860. Bernard C. 1857. Lecons sur les Effects des Substances Toxiques et Medicamenteuses. Paris: Bailliere; pp. 351-353. Blakeley AGH, Cunnane TC, Muir TC. 1986. Effects of calcium channel antagonists on action potential conduction and transmitter release in the guinea‐pig vas deferens. Br J Pharmacol 89: 235-244. Blakely RD, Berson HE, Fremeau RT, Jr Caron MG, Peek MM, et al. 1991. Cloning and expression of a functional serotonin transporter from rat brain. Nature 354: 66-70. Burnstock G. 1979. The ultrastructure of autonomic cholinergic nerves and Junctions. Prog Brain Res 79: 3-21. Cajal SR. 1937. Recollections of my life. Philadelphia: American Philosophical Society. Cunnane TC, Stja¨rne L. 1982. Secretion of transmitter from individual varicosities of guinea‐pig and mouse vas deferens. J Physiol 399: 607-632. Cunnane TC, Stja¨rne L. 1984. Frequency dependent intermittency and ionic basis of impulse conduction in postganglionic sympathetic fibres of guinea‐pig deferens. Neuroscience 11: 211-229.

109

110

4

Synaptic and nonsynaptic release of transmitters

Dale HH. 1956. Published in, Nobel Lectures in Physiology or Medicine, 1922–1941. Elsevier; Amsterdam: pp. 402-413. Demarque M, Represa A, Becq H, Khalilov I, Ben‐Ari Y, Aniksztejn L. 2002. Paracrine intercellular communicationby a Ca2þ and SNARE‐independent release of GABA and Glutamate prior to synapse formation. Neuron 36: 1051-1061. Descarries L, Saucier G. 1972. Disappearance of the locus coeruleus in the rat after intraventricular 6‐hydroxdopamine. Brain Res 37: 310-316. Descarries L, Gisiger V, Steriade M. 1997. Diffuse transmission by acetylcholine in the CNS. Prog Neurobiol 70(3): 603-625. Descarries L, Watkins KC, Lapierre Y. 1977. Noradrenergic axon terminals in the cerebral cortex of the rat. III Topometric ultrastructural analyses. Brain Res 133: 197-222. Descarries L, Lemay B, Doucet G, Berger B. 1987. Regional and laminar density of the dopamine innervation in adult rat cerebral cortex. Brain Res 441: 807-824. Eccles JC. 1946. An electrical hypothesis of synaptic and neuromuscular transmission. Ann NY Acad Sci 47: 429-455. Elliot TR. 1905. The action of adrenaline. J Physiol Lond 32: 401-467. Erlanger J. 1939. The initiation of impulses in axons. J Neurophysiol 2: 370-379. Feuerstein TJ, Hertting G, Lupp A, Neufang B. 1986. False labeling of dopaminergic terminals in the rabbit caudate nucleus: Uptake and release of [3H]‐5hydroxytryptamine. Br J Pharmacol 88: 677-684. Furness JB, Costa M. 1974. The adrenergic innervation of the gastrointestinal tract. Rev Physiol 69: 1-51. Fuxe K, Agnati LF. 1991. Two principal modes of electrochemical communication in the brain: Volume versus wiring transmission. Volume transmission in the brain: Novel mechanisms for neural transmission. Fuxe K, Agnati LF, editors. New York: Raven Press; pp. 1‐10. Gasser HS. 1939. Axons as samples of nervous tissue. J Neurophysiol 2: 361-369. Giros B, el Mestikawy S, Bertrand L, Caron MG. 1991. Cloning and functional characterization of a cocaine‐sensitive dopamine transporter. FEBS Lett 295: 149-154. Giros B, el Mestikawy S, Godinot N, Zheng K, Han H, et al. 1992. Cloning, pharmacological characterization, and chromosome assignment of the human dopamine transporter. Mol Pharmacol 42: 383-390. Gordon‐Weeks PR. 1982. Noradrenergic and non‐noradrenergic nerves containing small granular vesicles in Auerbach’s plexus of the guinea pig. Neuroscience 11: 2925-2936. Jones IW, Wonnacott S. 2004. Precise localization of alpha 7 nicotinic acetylcholine receptors on glutamatergic axon terminals in the rat ventral tegmental area. J Neurosci 24: 11244-11252.

Kaneda M, Farrant M, Cull‐Candy SG. 1995. Whole‐cell and single channel currents activated by GABA and glycine in granule cells of the rat cerebellum. J Physiol 485: 419-435. Katz B. 1969. The release of neural transmitter substances, Sherrington lecture. Liverpool: Liverpool University Press. Knoll J, Vizi ES. 1970. Presynaptic inhibition of acetylcholine release by endogeneous and exogeneous noradrenaline at high rate of stimulation. Br J Pharmacol 40: 554-555. Kullmann DM, Asztely F. 1998. Extrasynaptic glutamate spillover in the hippocampus: Evidence and implications. Trends Neurosci 21: 8-14. ¨ ber humorale U ¨ bertragbarkeit der Loewi O. 1921. U Herznervenwirkung. Pflu¨gers Arch 189: 239-242. ¨ ber humorale U ¨ bertragbarkeit der Loewi O. 1922. U Herznervenwirkung II. Mitteilung, Pflugers Arch 193: 201-213. Mitchell SJ, Silver RA. 2000. Glutamate spillover suppresses inhibition by activating presynaptic mGluRs. Nature 404: 498-502. Morita K, North RA. 1981. Opiates and encephalin reduce the excitability of neuronal processes. Neuroscience 6: 1943-1951. Nicholson C. 1985. Diffusion from an injected volume of a substance in brain tissue with arbitrary volume fraction and tortuosity. Brain Res 333: 325-329. Nicholson C. 2005. Factors governing diffusing molecular signals in brain extracellular space. J Neural Transm 112: 29-44. Nicholson C, Phillips JM, Gardner‐Medwin AR. 1979. Diffusion from an iontophoretic point source in the brain: Role of tortuosity and volume fraction. Brain Res 169: 580-584. Nusser Z, Mody I. 2002. Selective modulation of tonic and phasic inhibitions in dentate gyrus granule cells. J Neurophysiol 87: 2624-2628. Pacholczyk T, Blakely RD, Amara SG. 1991. Expression cloning of a cocaine‐ and antidepressant‐sensitive human noradrenaline transporter. Nature 350: 350-354. Paton WDM, Vizi ES. 1969. Non‐synaptic modulation of transmitter release: Pharmacological implication. Br J Pharmacol 35: 10-28. Raiteri L, Raiteri M, Bonanno G. 2002. Coexistence and function of different neurotransmitter transporters in the plasma membrane of CNS neurons. Prog Neurobiol 68: 287-309. Ramamoorthy S, Bauman AL, Moore KR, Han H, Yang‐Feng T, et al. 1993. Antidepressant‐ and cocaine‐sensitive human serotonin transporter. Molecular cloning, expression, and chromosomal localization. Proc Natl Acad Sci USA 90: 2542-2546. Rang HP. 2006. The receptor concept: Pharmacology’s big idea. Br J Pharmacol 147 (Suppl 1): S9-S16.

Synaptic and nonsynaptic release of transmitters Rossi DJ, Hamann M, Attwell D. 2003. Multiple modes of GABAergic inhibition of rat cerebellar granule cells. J Physiol 548: 97-110. Semyanov A, Kullmann DM. 2000. Modulation of GABAergic signaling among interneurons by metabotropic glutamate receptors. Neuron 25: 663-672. Semyanov A, Walker MC, Kullmann DM. 2003. GABA uptake regulates cortical excitability via cell type‐specific tonic inhibition. Nat Neurosci 6: 484-490. Semyanov A, Walker MC, Kullmann DM, Silver RA. 2004. Tonically active GABA A receptors: Modulating gain and maintaining the tone. Trends Neurosci 34(2): 262-269. Stja¨rne L. 1978. Facilitation and receptor mediated regulation of noradrenaline secretion by control of recruitment of varicosities as well as by control of electrosecretory coupling. Neuroscience 3: 1147-1155. Umbriaco D, Garcia S, Beaulieu C, Descarries L. 1995. Relational features of acetylcholine, noradrenaline, serotonin and GABA axon terminals in the stratum radiatum of adult rat hippocampus (CA1). Hippocampus 5: 605-620. Vizi ES. 1968. The inhibitory action of noradrenaline and adrenaline on release of acetylcholine from guinea‐pig ileum longitudinal strips. Arch Exp Path Pharmacol 259: 199-200. Vizi ES. 1974. Interaction between adrenergic and cholinergic systems presynaptic inhibitory effect of noradrenaline on: Acetylcholine release. J Neural Transm Suppl 11: 61-78. Vizi ES. 1979. Presynaptic modulation of neurochemical transmission. Prog Neurobiol 12: 181-290.

4

Vizi ES. 1980. Non‐synaptic modulation of transmitter release: Pharmacological implication. Trends Pharmacol Sci 1: 172-175. Vizi ES. 1984. Non‐synaptic interactions between neurons: Modulation of neurochemical transmission. London: Wiley. Vizi ES. 2000. Role of high‐affinity receptors and membrane transporters in nonsynaptic communication and drug action in the central nervous system. Pharmacol Rev 52: 63-90. Vizi ES, Kiss JP. 1998. Neurochemistry and pharmacology of the major hippocampal transmitter systems: Synaptic and nonsynaptic interactions. Hippocampus 8: 566-607. Vizi ES, Knoll J. 1971. The effects of sympathetic nerve stimulation and guanethidine on parasympathetic neuroeffector transmission: The inhibition of acetylcholine release. J Pharm Pharmacol 23: 918-925. Vizi ES, Ha´rsing LG Jr, Zima´nyi I, Gaa´l G. 1985. Release and turnover of noradrenaline in isolated median eminence: Lack of negative feedback modulation. Neuroscience 16: 907-916. Vizi ES, Zsilla G, Caron MG, Kiss JP. 2004. Uptake and release of norepinephrine by serotonergic terminals in norepinephrine transporter knock‐out mice: Implications for the action of selective serotonin reuptake inhibitors. J Neurosci 24: 7888-7994. Vizi ES, To¨ro¨k T, Seregi A, Serfo˝zo˝ p, Adam‐Vizi V. 1982. Na– K‐activated ATPase and the release of acetylcholine and noradrenaline. J Physiol (Paris) 78: 399-406.

111

5

Cholinergic Transmission

B. Lendvai

1

Historical Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 114

2

Synthesis, Storage, and Release of Acetylcholine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 114

3

Breakdown of ACh . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115

4

Structure of the Central Cholinergic System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116

5

Synaptic Versus Nonsynaptic Release of ACh . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116

6 6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.8 6.9

Nicotinic ACh Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Subunits and Subtypes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Selective Nicotinic Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Movement of Ions After nAChR Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Desensitization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Presynaptic Nicotinic Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118 Postsynaptic Nicotinic Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118 Role of Nicotinic Receptors in Reward Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 Special Role of Nicotinic Receptors in Neural Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 Nicotinic Receptors in Synaptic and Nonsynaptic Transmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120

7 7.1 7.2 7.3 7.4

Muscarinic ACh Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 Subunits and Subtypes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 Subcellular Action Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121 Pharmacology of Muscarinic Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122 Muscarinic Receptor Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122

8

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 124

#

2008 Springer ScienceþBusiness Media, LLC.

114

5

Cholinergic transmission

Abstract: The cholinergic system can modulate cognitive functions efficiently in the brain acting on a rich assembly of metabotropic and ionotropic receptors. The cholinergic system operates through the cooperation of the muscarinic and the nicotinic subsystems. While muscarinic ACh receptors mediate slow responses with considerable delay, nicotinic facilitation, following activation of nicotinic ACh receptors, evokes relatively fast responses. In some cases muscarinic and nicotinic ACh receptors form a dual control striatal on certain cell types, such as the spiny interneurons of the striatum. On important aspect of nicotinic transmission is that it modulates, rather than mediate, fast synaptic transmission. Desensitization of these receptors leads to a loss of function that is a key factor in the effect of nicotine during smoking. Desensitization extends the possible states of cholinergic transmission and increases the computational power of the neuron. As most nicotinic receptors are found in nonsynaptic localizations, especially on axons. They can directly release transmitters from presynaptic boutons. Importance of studies on nicotinic and muscarinic effects is highlighted by the fact that cholinergic therapy is the mainstay treatment for Alzheimer’s disease. The current view that nonsynaptic communication is dominant in cholinergic transmission also support the future perspective of drug therapy targeting high affinity nonsynaptic receptors. List of Abbreviations: ACh, acetylcholine; AChE, acetylcholine-esterase; ChAT, choline-acetyltranspherase; CNS, central nervous system; GABA, Gamma-aminobutyric acid; mAChR, muscarinic acetylcholine receptors; nAChR, nicotinic acetylcholine receptors; NMDA, N-methyl-D-aspartate

1

Historical Background

Cholinergic transmission has a unique position in the mind of neurochemists as being the first identified neurotransmitter during the first half of the twentieth century based on studies of Henry Dale, Otto Loewi, Feldberg and others. The very first discovery was the observation that a chemical substance (termed vagusstoff) is linked to the vagus action in the autonomic nervous system. Nicotinic receptors were the first receptors to be named. Today, studies focusing on the cholinergic system of the brain have received particular attention as the loss of cholinergic function is thought to underlie the age‐related learning impairments and memory loss that accompanies Alzheimer’s disease, one of the major health care problems in the world.

2

Synthesis, Storage, and Release of Acetylcholine

Acetylcholine (ACh) is synthesized in the boutons of cholinergic axons. Choline is taken up from the extracellular space by its specific transporter (> Figure 5-1). Although low‐affinity choline uptake is present in most tissues, cholinergic cells are equipped with a high‐affinity (Km
1–5 mM) choline uptake that provides stable choline supply under physiological conditions (choline is present in the plasma at about 10 mM). Hemicholinium, a plasma membrane choline transport inhibitor, causes disruption of ACh release especially during prolonged stimulation. Most of the choline derives from recycling of released ACh following hydrolysis. Another important source of choline is the breakdown of phosphatidylcholine. ACh is synthesized by the choline‐acetyltranspherase (ChAT), which transfers an acetyl group from acetyl coenzyme A to choline. Brain ChAT has a KD for choline of approximately 1 mM and a KD for acetyl coenzyme A of approximately 10 mM. There is an excess capacity of ChAT to produce ACh; the in vitro activity of the enzyme was found higher than that in vivo. The rate‐limiting step for ACh synthesis is likely the transport of acetyl coenzyme A, which comes from the inner membrane of mitochondria following the glucose– pyruvate transformation. The newly synthesized ACh is transported from the soma to the vesicles in the cholinergic axon terminal where it is stored. The transport of ACh can be prevented by incubation with vesamicol, which induces ACh accumulation in the plasma and the depletion of the vesicle pool of ACh. Most ACh molecules in the synaptosome, a neuronal preparation containing only the axon terminals, have been found associated with vesicles, as revealed by electron microscopy. ACh is typically released to the extracellular space from these vesicles following a fusion with the plasma membrane of the cell in

Cholinergic transmission

5

. Figure 5-1 Synthesis and nonsynaptic release of ACh. Cholinergic innervation arises from different loci where the cell body of the cholinergic projection neuron is located. Most of the ACh is released to the nonsynaptic, extracellular space (arrows). In contrast, ACh can be released in a few synaptic specializations for cholinergic transmission where ACh activates postsynaptic cell (gray shaded) in a one‐to‐one manner (small circle). Excitation of the postsynaptic cell, either by nonsynaptically or synaptically released ACh, occurs after activation postsynaptic/ dendritic nicotinic or muscarinic receptors. ACh is synthesized in the axon terminals where the high‐affinity choline uptake and the acetyl coenzyme A production provide the source molecules by ChAT. AChE is responsible for the breakdown of the released ACh

nonsynaptic sites (> Figure 5-1). From the functional aspect, it seems there are two distinct pools of ACh within the terminal: one ‘‘readily available’’ pool and a ‘‘reserve’’ or ‘‘depot’’ pool of ACh.

3

Breakdown of ACh

In the extracellular space, acetylcholine‐esterase (AChE) is responsible for the elimination of the effect of ACh by degrading the released ACh. AChE exists in several molecular forms (monomers, dimers, tetramers) with different solubility and subcellular localization. Cholinesterase genes encode a peptide without obvious membrane‐spanning regions making them available for secretion. Indeed, the soluble form of AChE can be released from nerve terminals. AChE shares structural similarity with neuroligins. During development the level of AChE regulates synaptogenesis for glutamatergic transmission (Dong et al., 2004). The distribution of AChE is relatively uniform in the central nervous system (CNS); it can be found in the cytoplasm and in the extracellular space even at large distances from cholinergic boutons. There are various ways to influence cholinergic transmission by modulating ACh release. Inhibitors of AChE, called anticholinesterases, induce accumulation of ACh in the extracellular space, and therefore prolong the action of the released ACh at the receptor. As a result, the decay of the postsynaptic current or potential by the released ACh is prolonged from 1–2 to 5–30 msec. On the system level, fasciculation and muscle twitching are initially observed after AChE inhibition, followed by flaccid paralysis. Some reversible inhibitors, such as gallamine and propidium, bind to a peripherial site, while other reversible inhibitors (neostigmine and physostigmine) act at the active site with an approximately 4‐h duration. The latter two inhibitors are used to treat glaucoma, myasthenia gravis, and dysfunction of smooth muscle. Another important application area of anticholinesterases is the pharmacotherapy of Alzheimer’s disease. Tacrine and donezepil are used to enhance cholinergic transmission in Alzheimer’s patients. Irreversible inhibitors completely inhibit the

115

116

5

Cholinergic transmission

breakdown of ACh, and therefore are dangerous for life. These molecules are used as insecticides. Nerve gases (sarin, soman) can cause death within 5 min of exposure. On the receptive side of cholinergic transmission there are two major receptor types: nicotinic (nAChRs) and muscarinic ACh receptors (mAChRs). All subtypes of nAChRs mediate excitatory conductances through the receptor ion channel. Muscarinic receptors have metabotropic functions and they can be excitatory or inhibitory depending on the subtype and the downstream subcellular mechanism.

4

Structure of the Central Cholinergic System

The cholinergic system is built up by two main cell types: cholinergic interneurons and projection neurons. ACh‐containing interneurons occur in the striatum, where the large aspiny neurons provide the cholinergic tone of the striatum. These cells are under the tonic inhibition of the nigrostriatal dopaminergic system through D2 receptors. In Parkinson’s disease the loss of this inhibitory influence results in high ACh concentration in the striatum. Prevention of the effect of ACh in Parkinson’s disease is an important part of the clinical therapy. Cholinergic interneurons were also identified in the hippocampus. Projection neurons include the cholinergic neurons of the medial septum, which provide an important innervation to the hippocampus, and the cholinergic neurons of the Meynert nucleus in the basal forebrain. The cholinergic neurons of the Meynert nucleus innervate the cerebral cortex. The cerebral cortex is diffusely innervated by cholinergic axons from the basal forebrain and axons from cholinergic interneurons (Eckenstein and Baughman, 1984). Cholinergic neurons of the medial habenula project to the interpeduncular nucleus. There are some other less investigated cholinergic loci in the CNS: vertical nucleus of the diagonal band projects to the hippocampus, neurons in the horizontal limb of the diagonal band innervate the olfactory bulb, the midbrain pedunculopontine and laterodorsal tegmental nucleus gives cholinergic input to the thalamus, and the cholinergic cells in the parabigeminal nucleus, which provide input to the superior colliculus.

5

Synaptic Versus Nonsynaptic Release of ACh

Ultrastructural morphometric studies demonstrated that the normal cholinergic innervation of adult rat parietal cortex (Umbriaco et al., 1994) and hippocampus (Umbriaco et al., 1995) predominantly does not result in synapses on other neurons (>85% of varicosities are without synaptic contact). These observations supported the idea that ACh participates primarily in nonsynaptic interactions (Descarries and Mechawar, 2000; Vizi, 2000). Cholinergic axons can be identified by staining for vesicular ACh transporter immunoreactivity, providing the strongest labeling in the stratum oriens of the CA1 region within the hippocampus (Towart et al., 2003). This observation indicates that the cholinergic input predominantly targets the basal dendrites of pyramidal neurons and the local interneurons. Another important evidence for the nonsynaptic nature of the cholinergic system is the somewhat surprising localization of AChE. The fact that AChE can be found in distant areas from cholinergic axon terminals and not restricted to the area of cholinergic release sites strongly suggests that the enzyme must sense and degrade ACh molecules that have traveled far in the extracellular space. This assumption is consistent with the nonsynaptic nature of most cholinergic boutons. ACh, released from the terminals without synaptic content, diffuses in the extracellular space to reach remote receptors where extracellular AChE terminates the cholinergic action. Neostigmine, a cholinesterase inhibitor, also enhances the extracellular level of noradrenaline measured by microdialysis (Kiss et al., 1999). During microdialysis, the samples are taken from the extracellular space, therefore, this method provides data for the nonsynaptic release of transmitters. It seems ACh is released from cholinergic boutons tonically; neostriatal cholinergic interneurons produce spontaneous tonic firing in the absence of synaptic input that results in a tonic release of ACh (Bennett and Wilson, 1999). The released ACh keeps the striatal DA terminals under a tonic control (Zhou et al., 2001). The release process can be pharmacologically manipulated; botulinum toxin A and tetanus toxin are known to block the release of ACh causing paralysis.

Cholinergic transmission

6

5

Nicotinic ACh Receptors

6.1 Subunits and Subtypes Genes encoding neural nAChR subunits express nine a (a2–a10) and three b subunits (b2–b4), which can coassemble to form pentameric functional receptors (Role and Berg, 1996). Pharmacological studies on transgenic mice have shown that the different subunits vary in their distribution and channel properties. The a4b2 and the a7 subtypes are predominant in the CNS while the a3b4 subunit‐containing nAChR is predominant in the peripherial nervous system (Role and Berg, 1996). Because of the diversity of subunits and minimal requirement of five subunits to form the channel of nAChRs, one cannot predict nAChR compositions based solely on the set of genes expressed by the neuron. There is evidence indicating the existence of multiple functional subtypes of nAChRs in the same neuron. For example, activation of nAChRs on hippocampal stratum radiatum interneurons involves both a7 and non‐a7 subunits and causes depolarization and action potential generation (Frazier et al., 1998b; McQuiston and Madison, 1999).

6.2 Selective Nicotinic Ligands There are a few selective agonists for nAChRs. Choline seems to be a selective activator of a7 nAChRs and a partial agonist on the a3b4 nAChRs (Alkondon et al., 1997). The antagonists methyllycaconitine (MLA) and a‐bungarotoxin selectively block the homomeric a7 nAChRs. At higher concentration (mM), MLA blocks other nAChRs, such as a4b2. Mecamylamine is a non‐selective channel blocker of all nAChRs with the highest potency on a4b4 and with relatively weak potency at a7 receptors. Nicotine‐induced increase in mEPSC frequency can be fully antagonized by a‐bungarotoxin, while mecamylamine causes only a 70% inhibition (Sharma and Vijayaraghavan, 2003). Mecamylamine, at higher (100 mM) concentration, can block other ligand‐gated ion channels, such as N‐methyl‐D‐aspartate (NMDA) receptors. The antagonist dihydro‐b‐ erithroidine (DHbE) has relatively low affinity for the a7 nAChRs and very high affinity to the a4bb2 and the a4b4 nAChRs.

6.3 Movement of Ions After nAChR Activation Nicotinic receptors are highly permeable to Naþ, Kþ, and Ca2þ. The Ca2þ/Naþ permeability ratio for the a7 receptor is >10 (the average ratio for all nAChRs is >1) (Seguela et al., 1993). In general, central nAChRs differ from the muscle nAChRs receptors in that the neuronal types are more permeable to Ca2þ (Vernino et al., 1992). Indeed, in hippocampal interneurons and cerebellar granule cells, nAChR stimulation induces Ca2þ transients via a7 nAChRs (Didier et al., 1995; Khiroug et al., 2003). Although nicotine can indeed cause large influx of Ca2þ into the cell, it is usually corroborated by various amplification mechanisms including voltage‐sensitive Ca2þ channels (VSCCs) and intracellular Ca2þ stores. In rat chromaffin cells, a part of the nAChR stimulation‐evoked Ca2þ response is mediated by VSCCs (Khiroug et al., 1997). Voltage‐sensitive Naþ channels can amplify the action of VSCCs in different tissue preparations (Mulle et al., 1992; Vijayaraghavan et al., 1992; Soliakov and Wonnacott, 1996). Nevertheless, in certain cells, Ca2þ rises by nicotinic stimulation can be exclusively mediated by the nAChR ion channel.

6.4 Desensitization Resting nAChR channels open in response to agonist binding to allow passage of ions. Prolonged presence of an agonist produces a desensitized state that no longer permits ion movements through the channel. Recovery from this state occurs after the agonist has been removed. For a long time, the main reason for the failures to detect nAChRs has been the rapid desensitization. Ca2þ accumulation due to receptor activation takes longer and so it was somewhat easier to detect rises in intracellular free Ca2þ; that is why most of the

117

118

5

Cholinergic transmission

first reports on cellular functions of central nAChRs emerged from optical studies (Vijayaraghavan et al., 1992). In outside‐out membrane patches of acutely isolated habenula neurons, applications of 100 mM nicotine produces macroscopic currents due to the opening of a large number of channels. During the continuous perfusion of the agonist, the number of open nAChR channels decreases exponentially because of receptor desensitization. A progressive loss in the number of channels with time is considered as receptor rundown (Lester and Dani, 1994). Desensitization expands the variability of nAChR‐mediated functions providing an extra computational power of the neuron. The findings that nicotine can cause similar effects to perfusion with nicotinic antagonists suggest a physiological role of desensitization of nAChRs during cigarette smoking (Chiodini et al., 1999; Zhou et al., 2001). Desensitization varies across cell types. In neurons, desensitization of nicotine‐induced currents becomes complete within a few seconds. Another important feature of the nicotinic desensitization is that the size of the intracellular Ca2þ transients can determine the rate of nAChR desensitization. In addition to the rapid desensitization, there is a long‐lasting or persistent inactivation of nAChRs, which, in contrast to desensitization, occurs immediately after the drug application and remains from 60 minutes to several hours (Lukas, 1991; Rowell and Duggan, 1998). The rates of recovery from desensitization depend on the time of agonist exposure and on the amount of agonist used to induce desensitization. The recovery time is about 10–30 sec after desensitization of nAChRs.

6.5 Presynaptic Nicotinic Receptors Presynaptic nAChRs have been described in various brain regions. These receptors that reside on the axon varicosity but far from the release site have been assigned the term preterminal receptors (Wonnacott, 1997). Why presynaptic (preterminal) nAChRs are so important for understanding cholinergic transmission? Activation of presynaptic nAChRs that precedes or coincides the arrival of an action potential into the terminal region of the neuron can increase the probability of release via the integrative capabilities of nAChR‐induced Ca2þ influx. The releasing action of presynaptic nAChRs are supported by several lines of experimental evidence: (1) the enhancement of the frequency and the size of excitatory synaptic potentials (EPSP) by nicotinic agonists leads to synchronization of the release process; this increase in the activity is sufficient to drive the postsynaptic cell above the firing threshold (Sharma and Vijayaraghavan, 2003) and (2) biochemical measurement of extracellular level of transmitters revealed that presynaptic nAChRs play an important role in regulating the release of different neurotransmitters (Vizi et al., 1995; Lendvai et al., 1996; Sershen et al., 1997; Vizi and Lendvai, 1999; Kofalvi et al., 2000). Different subtypes of nAChRs can be involved in the presynaptic regulation of transmitter release; a3b2 subtype composition has been suggested for the hippocampal noradrenaline release (Vizi et al., 1995; Sershen et al., 1997) and the striatal dopamine release (Kulak et al., 1997). b2 subunit‐containing nAChRs are involved in the regulation of GABA release in the thalamus (Lena and Changeux, 1997; Lena et al., 1993). In the superior cervical ganglion, activation of nAChRs enhanced the electrical field stimulation‐ induced release of ACh most likely via a3a7b2 nAChRs (Liang and Vizi, 1997). Activation of nAChRs could induce an increase in intracellular Ca2þ level of presynaptic neurits via a7 nAChRs (McGehee et al., 1995; Gray et al., 1996). Presynaptic nAChRs may combine their actions with an effect on the uptake, which results in a compound release mechanism (Szasz et al., 2005). This could occur because of the relatively high Ca2þ entry compared with Naþ influx that shifts the Ca2þ/Naþ exchanger to pump out Ca2þ leading to an increased Naþ entry and an opposite drive of the transporter. We may assume that if the Naþ concentration is high enough around the transporter it may change the direction of transport to release the transmitter instead of the uptake.

6.6 Postsynaptic Nicotinic Receptors There is evidence that a7‐ and b2‐containing nAChRs exist in both synaptic and nonsynaptic localizations in hippocampal neurons (Hill et al., 1993; Frazier et al., 1998b; Adams et al., 2001; Fabian‐Fine et al., 2001; Kawai et al., 2002; Graham et al., 2003). Most synapses containing nAChRs are not cholinergic but rather

Cholinergic transmission

5

belong to glutamatergic or GABAergic presynaptic partners (Fabian‐Fine et al., 2001). For these receptors, ACh must diffuse in the synaptic cleft from the extracellular space. Thus the transmission in which they are involved can be taken as a specific form of nonsynaptic communication, nonsynaptic modulation of the postsynaptic area. Besides the studies on the hippocampus, there are a number of observations for the existence of nAChRs at nonsynaptic sites (Hill et al., 1993; Horch and Sargent, 1995; Ullian and Sargent, 1995; Williams et al., 1998; Fabian‐Fine et al., 2001). In chicken, ciliary ganglion nAChRs contribute to both synaptic and nonsynaptic transmission; a3/a5 nAChR mediate synaptic responses, while a7 nAChRs appear in perisynaptic locations (Horch and Sargent, 1995; Williams et al., 1998). Most nAChRs, which receive ACh message from presynaptic cholinergic boutons, are not true ‘‘postsynaptic’’ receptors, but are mostly located on extrasynaptic membranes. Given the nonsynaptic nature of nicotinic transmission in the CNS, it is more precise to define them as somatodendritic receptors. Hippocampal GABAergic inhibitory interneurons can be excited by activation of nAChRs (Alkondon et al., 1997; Jones and Yakel, 1997; Frazier et al., 1998b; McQuiston and Madison, 1999). These stratum radiatum interneurons likely mediate feed‐forward inhibition primarily because they receive inputs from fibers entering the CA1 and inhibit mostly dendrites of pyramidal neurons. There is heterogeneity in the types of interneurons regarding the nicotine sensitivity; all interneurons in the stratum radiatum and the stratum lacunosum moleculare can be excited by nicotinic ligands but many interneurons in the pyramidal cell layer do not respond to nicotine (McQuiston and Madison, 1999). Functional dendritic nAChRs exist in pyramidal neurons of the hippocampus and cause excitatory actions (Ji et al., 2001; Ge and Dani, 2005). Nicotinic excitation in interneurons can be amplified by other factors; simultaneous activation of AMPA and NMDA receptors boosts the postsynaptic nicotinic current in interneurons of the hippocampus (Alkondon et al., 2003). Not only are hippocampal interneurons highly sensitive to nicotinic stimulation; different types of cortical layer 5 interneurons can be excited by stimulation of nAChRs (Xiang et al., 1998).

6.7 Role of Nicotinic Receptors in Reward Mechanisms Tobacco use is driven by the rewarding effects of nicotine in the brain. Nicotine is an addictive drug that reinforces self‐administration and increases locomotion. Nicotine elevates the level of dopamine in the nucleus accumbens (NAc), which in turn reinforces the drug use particularly during the acquisition phase (Dani et al., 2001). Chronic exposure to nicotine desensitizes nAChRs, and over the long term, nAChRs enter long‐lasting inactive states (Pidoplichko et al., 1997). Meanwhile, an increase in number of nAChRs can be observed with the possible purpose to maintain the level of excitability (Dani and Heinemann, 1996). Nicotine‐induced limbic dopamine release may drive tobacco use, while inactivation of nAChRs due to the sustained low‐dose nicotine plays a role in tolerance and withdrawal. The addictive power of nicotine during smoking may be linked to the known potential of nAChRs to improve synaptic plasticity in regions of the brain reward system such as the VTA and the NAc. Blood level of nicotine in smokers, ranging between 250 and 500 nM for about 10 min just after smoking a cigarette (Henningfield et al., 1993), may reach the firing threshold of mesolimbic dopaminergic neurons leading to dopamine release in the nucleus accumbens. The role of b2 nAChR is well established in the neural mechanisms of reward (Picciotto et al., 1998). Recovery of the endogenous cholinergic transmission may require 30 min following the elimination of nicotine.

6.8 Special Role of Nicotinic Receptors in Neural Plasticity The simplest case of cellular nicotinic function related to plasticity is the well‐described enhancement in the frequency of synaptic activity that outlasts for several minutes after the end of the stimulation (Radcliffe and Dani, 1998; Sharma and Vijayaraghavan, 2003). Nicotine can also enhance the short‐term depression observed at thalamocortical connections (Gil et al., 1997). Presynaptic a7 nAChRs are able to induce long‐ term potentiation in the mesolimbic system by pairing nicotine application to postsynaptic depolarization of dopaminergic neurons (Mansvelder and McGehee, 2000). Nicotinic stimulation can also induce transition of short‐term potentiation of synaptic potentials into long term in the hippocampus (Ji et al., 2001;

119

120

5

Cholinergic transmission

Mann and Greenfield, 2003). nAChRs use another possible mechanism to induce plastic changes in the hippocampus; nAChRs can transform the so‐called ‘‘silent’’ glutamatergic synapses into active synapses in CA1 pyramidal neurons (Maggi et al., 2003).

6.9 Nicotinic Receptors in Synaptic and Nonsynaptic Transmission There are experimental evidences for functional nAChRs, which mediate synaptic transmission. Cell types exhibiting nicotinic synaptic current include stratum radiatum and oriens interneurons (Frazier et al., 1998a), hippocampal CA1 pyramidal neurons (Hefft et al., 1999), pyramidal neurons and interneurons of the visual cortex (Roerig et al., 1997), and cells in the supraoptic nucleus (Hatton and Yang, 2002) and in the ciliary ganglion (Shoop et al., 2001). Both a7 and non‐a7 nAChRs can occur in synaptic transmission; in chick ciliary ganglion neurons, which innervate the iris and the choroid body, perisynaptic a7 nAChRs can also contribute to the fast synaptic current mediated by non‐a7 nAChRs (Chang and Berg, 1999). The dominant mode of the nicotinic influence on synaptic transmission, especially in the presynaptic actions, which, by nature, cannot be synaptic, seems to be the nonsynaptic information exchange. Activation of a7 nAChRs facilitates glutamatergic synaptic currents via a presynaptic action mechanism in cultured hippocampal neurons (Radcliffe and Dani, 1998), in CA1 and CA3 pyramidal neurons of the hippocampus (Ji et al., 2001; Maggi et al., 2003; Sharma and Vijayaraghavan, 2003), in pyramidal neurons of the rat auditory cortex (Aramakis and Metherate, 1998), and in mesolimbic dopaminergic neurons (Mansvelder and McGehee, 2000). Ca2þ influx seems to be a key player in the presynaptic mechanism of nAChR‐evoked responses. Low‐dose nicotine can increase frequency of miniature synaptic events, which are attributable to presynaptic activity, parallel with an increase in the presynaptic Ca2þ influx (McGehee et al., 1995; Gray et al., 1996; Maggi et al., 2003). There could be postsynaptic amplification by nAChRs, as well; low‐dose nicotine may enhance synaptic transmission by increasing the amplitude of evoked glutamatergic EPSCs via postsynaptic nAChRs in the interpeduncular nucleus (McGehee et al., 1995).

7

Muscarinic ACh Receptors

7.1 Subunits and Subtypes mAChRs received their name by their ability to bind muscarine, a product of the mushroom Amanita muscarina. mAChRs associate to G proteins and consist of five different receptors (M1–M5). Similar to nAChRs, a single neuron can express more than one mAChR subtype; for example, hippocampal pyramidal neurons have all five types of mAChRs (Levey et al., 1995). M1 receptors were first inactivated by genetic manipulation (Hamilton et al., 1997). Experiments on the M1 knockout mice revealed that seizure activity by muscarinic stimulation mostly connected to M1 receptors. In the striatum, M1 receptors might play a role in the early stages of Parkinson’s disease by increasing the dopamine release (Wess, 2003). Long‐ term potentiation in hippocampus is also reduced in M1 knockouts that exhibit a mild cognitive deficit in behavioral tests, suggesting the role of M1 receptors in learning and memory‐related processes (Anagnostaras et al., 2003). M2 receptor function has been associated with muscarinic stimulation‐induced tremor and akinesia. The block of VSCCs by muscarinic stimulation seems to be connected to activation of M2 receptors. M2 receptors participate in the presynaptic inhibition of transmitter release including ACh release. These autoreceptors of cholinergic transmission mediate feedback inhibition in the nervous system and in the neuromuscular junction. M3 receptors are widely expressed in the brain; the expression level is lower compared with other mAChR subtypes though. Adult M3 receptor‐deficient mice exhibit a weight loss of
25% (Yamada et al., 2001a). M3 receptors seem to play an important role in the regulation of appetite and daily food intake.

Cholinergic transmission

5

M3 receptors also mediate the contractile response of smooth muscle by cholinergic stimulation in various areas including urinary bladder, ileum, and trachea, and are involved in parasympathetic control of pupillary sphincter muscle contractility in the eye (Wess, 2003). M4 receptors, similarly to M2 receptors, participate in the autoinhibition of cholinergic axons. These receptors also occur as presynaptic heteroreceptors, for example, in the regulation of dopamine release. M4 receptors influence locomotion through an interaction with D1 dopamine receptors. M5 receptors were the last muscarinic receptors to be cloned; it is not surprising that the physiological role of this receptor is less understood. In addition, their low expression level in the brain and the lack of specific ligand make them difficult to study. M5 receptors mediate ACh‐induced dilation of cerebral blood vessels (Yamada et al., 2001b) that might be important in the development of Alzheimer’s disease. M5 receptor is the sole mAChR subtype expressed by the substantia nigra dopaminergic neurons. This observation is particularly important as the loss of the nigrostriatal dopaminergic input is the cellular defect underlying Parkinson’s disease and ligands targeting mAChRs are used in the therapy of this disease. M5 receptors are involved in the oxotremorine‐induced facilitation of dopamine release in the striatum (Yamada et al., 2001b). Nevertheless, indirect effects through M4 receptors on GABAergic cells also contribute to the oxotremorine‐induced facilitation.

7.2 Subcellular Action Mechanisms M1, M3, and M5 receptors couple to Gq proteins and their activation mobilizes intracellular Ca2þ through a store‐mediated release. Through Gq/11, these mAChRs activate phospholipase C, which initiates the phosphatidylinositol turnover and produces inositol trisphosphate (IP3)‐mediated Ca2þ release from the intracellular Ca2þ stores, such as the endoplasmic reticulum. The breakdown of phosphatidylinositol also leads to diacylglycerol production, which activates protein kinase C and initiates a number of downstream cellular effectors. Overall, M1 or M3 receptors can induce ‘‘slow’’ increase in neuronal excitability by the higher Ca2þ level. M2 and M4 mAChRs can activate Kþ channels of the plasma membrane through Gi proteins resulting in hyperpolarization that can be seen as inhibitory effects on neural activity. As a consequence, M2 and M4 receptors inhibit action potential firing. One or both of these subtypes are found presynaptically on cholinergic (autoreceptors) and other transmitter‐containing (heteroreceptors) axon terminals where they inhibit neurotransmitter release. In the subcellular level, activation of these mAChRs causes activation of inward rectifying Kþ channels, inhibition of Ca2þ channels, and inhibition of adenylate cyclase and the subsequent intracellular processes. Not only direct potentiation of Kþ channels can be used to hyperpolarize target cells but the increases in intracellular free Ca2þ by M1 receptors can also activate Kþ current. M3 mAChRs are able to form functional dimers that make the mAChR‐mediated function more complex in vivo (Wess, 2003). M1 and M2 mAChRs in neurons and M2 and M3 mAChRs in smooth muscle cells may also heterodimerize. The functional role of dimerization of mAChRs are not known. Although ACh is rapidly hydrolyzed after release, desensitization of mAChRs occurs under physiological conditions. As with a large number of G‐protein coupled receptors, agonist‐induced desensitization of mAChRs usually involves receptor phosphorylation (Haga and Haga, 1990). The M1 and M3 mAChRs have been shown to be phosphorylated by protein kinase C (PKC). This receptor modification occurs on serine and threonine residues in the third cytoplasmic loop and the C‐terminus of the mAChRs. An array of protein kinases is able to phosphorylate mAChRs, including various G‐protein coupled receptor kinases (GRKs), casein kinase 1a (CK1a), and diacylglycerol‐regulated PKC. The mitogen‐activated protein kinases ERK1/2 phosphorylate a serine residue in GRK2, which decreases the activity of the kinase toward G‐protein coupled receptor substrates. Cytosolic b‐arrestin interacts with the phosphorylated receptor, leading to uncoupling of the mAChRs from the G proteins and clathrin‐coated vesicle formation. Following release of dynamin, clathrin, and b‐arrestin, the vesicle recycles back to the plasma membrane. Receptor internalization may represent a molecular mechanism appropriate for selective attenuation of particular mAChR signaling pathways.

121

122

5

Cholinergic transmission

7.3 Pharmacology of Muscarinic Receptors General muscarinic agonists include several alkaloids, such as muscarine, arecoline, and synthetic compounds, such as carbachol. Bethanechol, metoclopramide, pilocarpine, and oxotremorine are other frequently used nonselective agonists of mAChRs (M1–M5). Absolute selectivity for any muscarinic agent has so far not been achieved. Subtype‐selective agonists are relatively rare; McN‐A‐343 is selective for the M1 receptor. There is more possibility to identify muscarinic subtypes with the use of selective antagonists. Subtypes of mAChRs can be identified by pharmacological tools (> Table 5-1): M1 receptors exhibit selectivity to the antagonist pirenzepine, M2 receptors can be selectively blocked by AF‐DX 116, M3 receptors can be selectively blocked by 4‐DAMP, whereas himbacine shows high affinity for the M4 receptor. M5 receptor has no selective ligand currently, and participation of this subtype is assumed when mAChR‐ mediated action is likely but all of the listed subtype‐selective antagonists are ineffective. The most used nonselective antagonists for mAChRs are atropine and scopolamine. Atropine, scopolamine, pirenzepine, and pilocarpine bind on the same binding site as the agonist but produce different conformational changes in the receptor structure that ultimately leads to the inhibition of the receptor function (van Koppen and Kaiser, 2003). All mAChR subtypes are susceptible to allosteric modulation; the binding of the allosteric modulator to the allosteric binding site results in a change in the conformation of the classical binding site leading to a change in the affinity of the receptor for classical muscarinic agonists and antagonists. There are also allosteric agents with positive cooperation on the binding of mAChR antagonists, such as strychnine (Lazareno and Birdsall, 1995). Other compounds, such as brucine, vincamine, and alcuronium, are able to allosterically modify the binding of mAChR agonists (Jakubik et al., 1997).

7.4 Muscarinic Receptor Functions mAChRs are responsible for postganglionic parasympathetic neurotransmission. Some sympathetic responses, such as sweating and piloerection, are also mediated by mAChRs. Although the ganglionic transmission is mediated by nAChRs by producing fast excitatory synaptic potential (EPSP) on the postsynaptic neuron, there is also a slow EPSP mediated by mAChRs. The slow EPSP decays in 1 sec and has a duration of 30–60 sec. Muscarinic agonists, acting on these ganglionic mAChRs, can enhance fast EPSPs under conditions of repetitive stimulation. It is important to note that not all of the peripherial organs are equipped with receptors to both sympathetic and parasympathetic transmitters. Therefore, nicotine‐sensitive ganglionic stimulation may end up with a non‐cholinergic effect, for example, in the blood vessels. M3 and M2 mAChRs are enriched in airway smooth muscle. In the CNS, the presynaptic mAChR (M2) inhibits cholinergic boutons as a muscarinic autoreceptor. Antagonism of this receptor by scopolamine results in release of choline into the extracellular space (Sarter and Parikh, 2005). It is well known that mAChRs play an important role in spike frequency adaptation in central neurons (Nicoll et al., 1990). Galantamine, a third generation cholinesterase inhibitor used in the therapy of Alzheimer’s disease, could dose dependently reduce the after hyperpolarization after a burst of action potentials and the spike‐ frequency accommodation of hippocampal CA1 neurons (Oh et al., 2005). Larger trains of backpropagating action potentials, exhibiting adaptation, were shown to be subject to modulation by mAChRs suggesting that dendritic integration can be modified by mAChRs (Tsubokawa and Ross, 1997). The cholinesterase blocker physostigmine and cholinomimetics evokes theta wave activity in the hippocampus through a muscarinic mechanism (Olpe et al., 1987; Konopacki et al., 1988; Golebiewski et al., 2002; Yoder and Pang, 2005). The rhythm is believed to be critical for the temporal coding or decoding of active neuronal ensembles and the modification of synaptic weights. Muscarinic antagonists, such as scopolamine or atropine, impair cognitive abilities in humans (Drachman, 1977). Recently it has been shown that transient activation of M1 mAChRs induces Ca2þ release from intracellular stores via IP3 and subsequent activation of an SK‐type Ca2þ‐activated Kþ conductance showing that ACh can directly inhibit neocortical pyramidal neurons through Ca2þ mobilization (Gulledge and Stuart, 2005). In the sensory system, presence of mAChRs has been shown on vestibular hair cells to evoke transmitter release from these cells (Derbenev et al., 2005).

Gene G protein Subcellular effect Selective agonist Selective antagonists

McN‐A‐343 Pilocarpine L‐689,660 Xanomeline Pirenzepine Telenzepine

M1 M1 Gq/11 Ic. Ca2þ (þ)

AF‐DX 116 Methoctramine AF‐DX 384 Gallamine Himbacine Tripitramine

M2 m2 Gi Kþ channels (þ) Ca2þ channels () adnylate cyclase () Bethanechol

. Table 5-1 mAChRs: basic pharmacology and subcellular effects of different subtypes

4‐DAMP Hexahydro‐sila‐ difenidol

L‐689,660

M3 m3 Gq/11 Ic. Ca2þ (þ)

M5 m5 Gq/11 Ic. Ca2þ (þ) – –

M4 m4 Gi Kþ channels (þ) Ca2þ channels () adnylate cyclase () McN‐A‐343 Himbacine Tropicamide AF‐DX 384

Cholinergic transmission

5 123

124

5 8

Cholinergic transmission

Conclusions

There are several examples that the effectors of the cholinergic, transmission, namely the muscarinic, and the nicotinic systems cooperate in the CNS. Although mAChRs are metabotropic receptors and mediate slow responses with considerable delay, nicotinic facilitation, following fast activation of nAChRs, can be sustained for up to 2 h. Interestingly, muscarinic inhibition seems to be more transient in certain cells (Girod and Role, 2001). Nicotinic agonists depolarize striatal interneurons and induce firing through non‐a 7 nAChRs, which, together with presynaptic inhibition through muscarinic receptors, form a dual cholinergic control on spiny interneurons of the striatum (Koos and Tepper, 2002). Main threads of current theories of nicotinic functions in the CNS include the following: (1) nAChRs may modulate, rather than mediate, fast synaptic transmission (McGehee and Role, 1995), (2) desensitization of the nAChRs, i.e., loss of function, is a key factor in the effect of nicotine during smoking and also shape the nAChR‐mediated activity in normal cholinergic transmission extending the computational power of the neuron, and (3) nAChRs directly release transmitters from presynaptic boutons skipping postsynaptic secondary modulations (Vizi and Lendvai, 1999). Taking these data together, we can conclude that the basis of the well‐known nicotinic enhancement of memory and learning function (Levin and Rezvani, 2000) is constructed on the level of cellular synaptic plasticity. Nicotine selectively improves the cognitive performance (especially those involving attentional processes) in deprived smokers and in cases with impaired cognition (Freedman et al., 1995), such as Alzheimer’s disease (Rezvani and Levin, 2001; Newhouse et al., 2004). Cholinergic therapy is the mainstay treatment for Alzheimer’s disease. In conclusion, cholinergic system plays an effective role in modulating cognitive functions in the brain acting on a rich assembly of metabotropic and ionotropic receptors. The current view that nonsynaptic communication is dominant in cholinergic transmission also supports the future perspective of drug therapy targeting high‐affinity nonsynaptic receptors.

References Adams CE, Stitzel JA, Collins AC, Freedman R. 2001. Alpha7‐ nicotinic receptor expression and the anatomical organization of hippocampal interneurons. Brain Res 922: 180-190. Alkondon M, Pereira EF, Albuquerque EX. 2003. NMDA and AMPA receptors contribute to the nicotinic cholinergic excitation of CA1 interneurons in the rat hippocampus. J Neurophysiol 90: 1613-1625. Alkondon M, Pereira EF, Cortes WS, Maelicke A, Albuquerque EX. 1997. Choline is a selective agonist of a7 nicotinic acetylcholine receptors in the rat brain neurons. Eur J Neurosci 9: 2734-2742. Anagnostaras SG, Murphy GG, Hamilton SE, Mitchell SL, Rahnama NP, et al. 2003. Selective cognitive dysfunction in acetylcholine M1 muscarinic receptor mutant mice. Nat Neurosci 6: 51-58. Aramakis VB, Metherate R. 1998. Nicotine selectively enhances NMDA receptor‐mediated synaptic transmission during postnatal development in sensory neocortex. J Neurosci 18: 8485-8495. Bennett BD, Wilson CJ. 1999. Spontaneous activity of neostriatal cholinergic interneurons in vitro. J Neurosci 19: 5586-5596. Chang KT, Berg DK. 1999. Nicotinic acetylcholine receptors containing a7 subunits are required for reliable synaptic transmission in situ. J Neurosci 19: 3701-3710.

Chiodini FC, Tassonyi E, Hulo S, Bertrand D, Muller D. 1999. Modulation of synaptic transmission by nicotine and nicotinic antagonists in hippocampus. Brain Res Bull 48: 623-628. Dani JA, Heinemann S. 1996. Molecular and cellular aspects of nicotine abuse. Neuron 16: 905-908. Dani JA, Ji D, Zhou FM. 2001. Synaptic plasticity and nicotine addiction. Neuron 31: 349-352. Derbenev AV, Linn CL, Guth PS. 2005. Muscarinic ACh receptor activation causes transmitter release from isolated frog vestibular hair cells. J Neurophysiol 94: 3134-3142. Descarries L, Mechawar N. 2000. Ultrastructural evidence for diffuse transmission by monoamine and acetylcholine neurons of the central nervous system. Prog Brain Res 125: 27-47. Didier M, Berman SA, Lindstrom J, Bursztajn S. 1995. Characterization of nicotinic acetylcholine receptors expressed in primary cultures of cerebellar granule cells. Mol Brain Res 30: 17-28. Dong H, Xiang YY, Farchi N, Ju W, Wu Y, et al. 2004. Excessive expression of acetylcholinesterase impairs glutamatergic synaptogenesis in hippocampal neurons. J Neurosci 24: 8950-8960. Drachman DA. 1977. Memory and cognitive function in man: Does the cholinergic system have a specific role? Neurology 27: 783-790.

Cholinergic transmission Eckenstein F, Baughman RW. 1984. Two types of cholinergic innervation in cortex, one co‐localized with vasoactive intestinal polypeptide. Nature 309: 153-155. Fabian‐Fine R, Skehel P, Errington ML, Davies HA, Sher E, et al. 2001. Ultrastructural distribution of the a7 nicotinic acetylcholine receptor subunit in rat hippocampus. J Neurosci 21: 7993-8003. Frazier CJ, Buhler AV, Weiner JL, Dunwiddie TV. 1998a. Synaptic potentials mediated via a‐bungarotoxin‐sensitive nicotinic acetylcholine receptors in rat hippocampal interneurons. J Neurosci 18: 8228-8235. Frazier CJ, Rollins YD, Breese CR, Leonard S, Freedman R, et al. 1998b. Acetylcholine activates an a‐bungarotoxin‐ sensitive nicotinic current in rat hippocampal interneurons, but not pyramidal cells. J Neurosci 18: 1187-1195. Freedman R, Hall M, Adler LE, Leonard S. 1995. Evidence in postmortem brain tissue for decreased numbers of hippocampal nicotinic receptors in schizophrenia. Biol Psychiatry 38: 22-33. Ge S, Dani JA. 2005. Nicotinic acetylcholine receptors at glutamate synapses facilitate long‐term depression or potentiation. J Neurosci 25: 6084-6091. Gil Z, Connors BW, Amitai Y. 1997. Differential regulation of neocortical synapses by neuromodulators and activity. Neuron 19: 679-686. Girod R, Role LW. 2001. Long‐lasting enhancement of glutamatergic synaptic transmission by acetylcholine contrasts with response adaptation after exposure to low‐level nicotine. J Neurosci 21: 5182-5190. Golebiewski H, Eckersdorf B, Konopacki J. 2002. Septal cholinergic mediation of hippocampal theta in the cat. Brain Res Bull 58: 323-335. Graham AJ, Ray MA, Perry EK, Jaros E, Perry RH, et al. 2003. Differential nicotinic acetylcholine receptor subunit expression in the human hippocampus. J Chem Neuroanat 25: 97-113. Gray R, Rajan AS, Radcliffe KA, Yakehiro M, Dani JA. 1996. Hippocampal synaptic transmission enhanced by low concentrations of nicotine. Nature 383: 713-716. Gulledge AT, Stuart GJ. 2005. Cholinergic inhibition of neocortical pyramidal neurons. J Neurosci 25: 10308-10320. Haga K, Haga T. 1990. Dual regulation by G proteins of agonistdependent phosphorylation of muscarinic acetylcholine receptors. FEBS Lett 268: 43-47. Hamilton SE, Loose MD, Qi M, Levey AI, Hille B, et al. 1997. Disruption of the m1 receptor gene ablates muscarinic receptor‐dependent M current regulation and seizure activity in mice. Proc Natl Acad Sci USA 94: 13311-13316. Hatton GI, Yang QZ. 2002. Synaptic potentials mediated by a7 nicotinic acetylcholine receptors in supraoptic nucleus. J Neurosci 22: 29-37.

5

Hefft S, Hulo S, Bertrand D, Muller D. 1999. Synaptic transmission at nicotinic acetylcholine receptors in rat hippocampal organotypic cultures and slices. J Physiol 515: 769-776. Henningfield JE, Stapleton JM, Benowitz NL, Grayson RF, London ED. 1993. Higher levels of nicotine in arterial than in venous blood after cigarette smoking. Drug Alcohol Depend 33: 23-29. Hill JA, Zoli M, Bourgeois J‐P, Changeux J‐P. 1993. Immunocytochemical localization of neural nicotinic receptor: The b2‐subunit. J Neurosci 13: 1551-1568. Horch HL, Sargent PB. 1995. Perisynaptic surface distribution of multiple classes of nicotinic acetylcholine receptors on neurons in the chicken ciliary ganglion. J Neurosci 15: 7778-7795. Jakubik J, Bacakova L, El‐Fakahany EE, Tucek S. 1997. Positive cooperativity of acetylcholine and other agonists with allosteric ligands on muscarinic acetylcholine receptors. Mol Pharmacol 52: 172-179. Ji D, Lape R, Dani JA. 2001. Timing and location of nicotinic activity enhances or depresses hippocampal synaptic plasticity. Neuron 31: 131-141. Jones S, Yakel JL. 1997. Functional nicotinic ACh receptors on interneurones in the rat hippocampus. J Physiol 504: 603-610. Kawai H, Zago W, Berg DK. 2002. Nicotinic a7 receptor clusters on hippocampal GABAergic neurons: Regulation by synaptic activity and neurotrophins. J Neurosci 22: 7903-7912. Khiroug L, Giniatullin R, Klein RC, Fayuk D, Yakel JL. 2003. Functional mapping and Ca2þ regulation of nicotinic acetylcholine receptor channels in rat hippocampal CA1 neurons. J Neurosci 23: 9024-9031. Khiroug L, Giniatullin R, Sokolova E, Talantova M, Nistri A. 1997. Imaging of intracellular calcium during desensitization of nicotinic acetylcholine receptors of rat chromaffin cells. Br J Pharmacol 122: 1323-1332. Kiss JP, Vizi ES, Westerink BHC. 1999. Effect of neostigmine on the hippocampal noradrenaline release: Role of cholinergic receptors. Neuroreport 10: 81-86. Kofalvi A, Sperlagh B, Zelles T, Vizi ES. 2000. Long‐lasting facilitation of 4‐amino‐n‐[2,3‐3H]butyric acid ([3H] GABA) release from rat hippocampal slices by nicotinic receptor activation. J Pharmacol Exp Ther 295: 453-462. Konopacki J, Bland BH, Roth SH. 1988. Evidence that activation of in vitro hippocampal theta rhythm only involves muscarinic receptors. Brain Res 455: 110-114. Koos T, Tepper JM. 2002. Dual cholinergic control of fast‐ spiking interneurons in the neostriatum. J Neurosci 22: 529-535. Kulak JM, Nguyen TA, Olivera BM, McIntosh JM. 1997. a‐ Conotoxin MII blocks nicotine‐ stimulated dopamine release in rat striatal synaptosomes. J Neurosci 17: 5263-5270.

125

126

5

Cholinergic transmission

Lazareno S, Birdsall NJ. 1995. Detection, quantitation, and verification of allosteric interactions of agents with labeled and unlabeled ligands at G protein‐coupled receptors: Interactions of strychnine and acetylcholine at muscarinic receptors. Mol Pharmacol 48: 362-78. Lena C, Changeux J‐P, Mulle C. 1993. Evidence for ‘preterminal’ nicotine receptors on GABAergic axons in the rat interpeduncular nucleus. J Neurosci 13: 2680-2688. Lena C, Changeux J-P. 1997. Role of Ca2+ ions in nicotinic facilitation of GABA release in mouse thalamus. J Neurosci 17: 576-585. Lendvai B, Sershen H, Lajtha A, Santha E, Baranyi M, et al. 1996. Differential mechanisms involved in the effect of nicotinic agonists DMPP and lobeline to release [3H]5‐ HT from rat hippocampal slices. Neuropharmacology 35: 1769-1777. Lester RA, Dani JA. 1994. Time‐dependent changes in central nicotinic acetylcholine channel kinetics in excised patches. Neuropharmacology 33: 27-34. Levey AI, Edmunds SM, Koliatsos V, Wiley RG, Heilman CJ. 1995. Expression of m1‐m4 muscarinic acetylcholine receptor proteins in rat hippocampus and regulation by cholinergic innervation. J Neurosci 15: 4077-4092. Levin ED, Rezvani AH. 2000. Development of nicotinic drug therapy for cognitive disorders. Eur J Pharmacol 393: 141-146. Liang SD, Vizi ES. 1997. Positive feedback modulation of acetylcholine release from isolated superior cervical ganglion. J Pharmacol Exp Ther 280: 650-655. Lukas RJ. 1991. Effects of chronic nicotinic ligand exposure on functional activity of nicotinic acetylcholine receptors expressed by cells of the PC12 rat pheochromocytoma or the TE671/RD human clonal line. J Neurochem 56: 1134-1145. Maggi L, Le Magueresse C, Changeux JP, Cherubini E. 2003. Nicotine activates immature ‘‘silent’’ connections in the developing hippocampus. Proc Natl Acad Sci USA 100: 2059-2064. Mann EO, Greenfield SA. 2003. Novel modulatory mechanisms revealed by the sustained application of nicotine in the guinea‐pig hippocampus in vitro. J Physiol 551.2: 539-550. Mansvelder HD, McGehee DS. 2000. Long‐term potentiation of excitatory inputs to brain reward areas by nicotine. Neuron 27: 349-357. McGehee DS, Role LW. 1995. Physiological diversity of nicotinic acetylcholine receptors expressed by vertebrate neurons. Annu Rev Physiol 57: 521-546. McGehee DS, Heath MJ, Gelber S, Devay P, Role LW. 1995. Nicotine enhancement of fast excitatory synaptic transmission in CNS by presynaptic receptors. Science 269: 1692-1696.

McQuiston AR, Madison DV. 1999. Nicotinic receptor activation excites distinct subtypes of interneurons in the rat hippocampus. J Neurosci 19: 2887-2896. Mulle C, Choquet D, Korn H, Changeux JP. 1992. Calcium influx through nicotinic receptor in rat central neurons: Its relevance to cellular regulation. Neuron 8: 135-143. Newhouse PA, Potter A, Singh A. 2004. Effects of nicotinic stimulation on cognitive performance. Curr Opin Pharmacol 4: 36-46. Nicoll RA, Malenka RC, Kauer JA. 1990. Functional comparison of neurotransmitter receptor subtypes in mammalian central nervous system. Physiol Rev 70: 513-565. Oh MM, Wu WW, Power JM, Disterhoft JF. 2005. Galantamine increases excitability of CA1 hippocampal pyramidal neurons. Neuroscience Oct 19; [Epub ahead of print]. Olpe HR, Klebs K, Kung E, Campiche P, Glatt A, et al. 1987. Cholinomimetics induce theta rhythm and reduce hippocampal pyramidal cell excitability. Eur J Pharmacol 142: 275-283. Picciotto MR, Zoli M, Rimondini R, Lena C, Marubio LM, et al. 1998. Acetylcholine receptors containing the b2 subunit are involved in the reinforcing properties of nicotine. Nature 391: 173-177. Pidoplichko VI, De Biasi M, Williams JT, Dani JA. 1997. Nicotine activates and desensitizes midbrain dopamine neurons. Nature 390: 401-404. Radcliffe KA, Dani JA. 1998. Nicotinic stimulation produces multiple forms of increased glutamatergic synaptic transmission. J Neurosci 18: 7075-7083. Rezvani AH, Levin ED. 2001. Cognitive effects of nicotine. Biol Psychiatry 49: 258-267. Roerig B, Nelson DA, Katz LC. 1997. Fast synaptic signaling by nicotinic acetylcholine and serotonin 5‐HT3 receptors in developing visual cortex. J Neurosci 17: 8353-8362. Role LW, Berg DK. 1996. Nicotinic receptors in the development and modulation of CNS synapses. Neuron 16: 1077-1085. Rowell PP, Duggan DS. 1998. Long‐lasting inactivation of nicotinic receptor function in vitro by treatment with high concentrations of nicotine. Neuropharmacology 37: 103-111. Sarter M, Parikh V. 2005. Choline transporters, cholinergic transmission and cognition. Nat Rev Neurosci 6: 48-56. Seguela P, Wadiche J, Dineley‐Miller K, Patrick JW. 1993. Molecular cloning, functional properties, and distribution of rat brain a7: A nicotinic cation channel highly permeable to calcium. J Neurosci 13: 596-604. Sershen H, Balla A, Lajtha A, Vizi ES. 1997. Characterization of nicotinic receptors involved in the release of norepinephrine from hippocampus. Neuroscience 77: 121-130.

Cholinergic transmission Sharma G, Vijayaraghavan S. 2003. Modulation of presynaptic store calcium induces release of glutamate and postsynaptic firing. Neuron 38: 929-939. Shoop RD, Chang KT, Ellisman MH, Berg DK. 2001. Synaptically driven calcium transients via nicotinic receptors on somatic spines. J Neurosci 21: 771-781. Soliakov L, Wonnacott S. 1996. Voltage sensitive Ca2þ channels involved in nicotinic receptor‐mediated 3Hdopamine release from rat striatal synaptosomes. J Neurochem 67: 163-170. Szasz BK, Mayer A, Zsilla G, Lendvai B, Vizi ES, et al. 2005. Carrier‐mediated release of monoamines induced by the nicotinic acetylcholine receptor agonist DMPP. Neuropharmacology 49: 400-409. Towart LA, Alves SE, Znamensky V, Hayashi S, McEwen BS, et al. 2003. Subcellular relationships between cholinergic terminals and estrogen receptor‐alpha in the dorsal hippocampus. J Comp Neurol 463: 390-401. Tsubokawa H, Ross WN. 1997. Muscarinic modulation of spike backpropagating in the apical dendrites of hippocampal CA1 pyramidal neurons. J Neurosci 17: 5782-5791. Ullian EM, Sargent PB. 1995. Pronounced cellular diversity and extrasynaptic location of nicotinic acetylcholine receptor subunit immunreactivities in the chicken pretectum. J Neurosci 15: 7012-7023. Umbriaco D, Garcia S, Beaulieu C, Descarries L. 1995. Relational features of acetylcholine, noradrenaline, serotonin and GABA axon terminals in the stratum radiatum of adult rat hippocampus (CA1). Hippocampus 5: 605-620. Umbriaco D, Watkins KC, Descarries L, Cozzari C, Hartman BK. 1994. Ultrastructural and morphometric features of the acetylcholine innervation in adult rat parietal cortex: An electron microscopic study in serial sections. J Comp Neurol 348: 351-373. van Koppen CJ, Kaiser B. 2003. Regulation of muscarinic acetylcholine receptor signaling. Pharmacol Ther 98: 197-220. Vernino S, Amador M, Luetje CW, Patrick J, Dani JA. 1992. Calcium modulation and high calcium permeability of neuronal nicotinic acetylcholine receptors. Neuron 8: 127-134.

5

Vijayaraghavan S, Pugh PC, Zhang ZW, Rathouz MM, Berg DK. 1992. Nicotinic receptors that bind agr‐bungarotoxin on neurons raise intracellular free Ca2þ . Neuron 8: 353-362. Vizi ES. 2000. Role of high‐affinity receptors and membrane transporters in nonsynaptic communication and drug action in the central nervous system. Pharmacol Rev 52: 63-89. Vizi ES, Lendvai B. 1999. Modulatory role of presynaptic nicotinic receptors in synaptic and non‐synaptic chemical communication in the central nervous system. Brain Res Rev 30: 219-235. Vizi ES, Sershen H, Balla A, Mike A, Windish K, et al. 1995. Neurochemical evidence of heterogeneity of presynaptic and somatodendritic nicotinic acetylcholine receptors. Ann NY Acad Sci 757: 84-99. Wess J. 2003. Novel insights into muscarinic acetylcholine receptor function using gene targeting technology. Trends Pharmacol Sci 24: 414-420. Williams BM, Temburni MK, Levey MS, Bertrand S, Bertrand D, et al. 1998. The long internal loop of the a3 subunit targets nAChRs to subdomains within individual synapses on neurons in vivo. Nat Neurosci 1: 557-562. Wonnacott S. 1997. Presynaptic nicotinic ACh receptors. Trends Neurosci 20: 92-98. Xiang Z, Huguenard JR, Prince DA. 1998. Cholinergic switching within neocortical inhibitory networks. Science 281: 985-988. Yamada M, Lamping KG, Duttaroy A, Zhang W, Cui Y, et al. 2001b. Cholinergic dilation of cerebral blood vessels is abolished in M(5) muscarinic acetylcholine receptor knockout mice. Proc Natl Acad Sci USA 98: 14096-14101. Yamada M, Miyakawa T, Duttaroy A, Yamanaka A, Moriguchi T, et al. 2001a. Mice lacking the M3 muscarinic acetylcholine receptor are hypophagic and lean. Nature 410: 207-212. Yoder RM, Pang KC. 2005. Involvement of GABAergic and cholinergic medial septal neurons in hippocampal theta rhythm. Hippocampus 15: 381-392. Zhou FM, Liang Y, Dani JA. 2001. Endogenous nicotinic cholinergic activity regulates dopamine release in the striatum. Nat Neurosci 4: 1224-1229.

127

6

Molecular Genetics of Brain Noradrenergic Neurotransmission

R. Meloni

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 130

2 2.1 2.2 2.3 2.4 2.5 2.6 2.6.1 2.6.2

Brain Noradrenergic System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 130 Ontogeny . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 130 Nuclei and Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 NE Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 NE Storage and Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 NE Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 132 Termination of NE Synaptic Transmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 Reuptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134

3 3.1 3.1.1 3.2

Functional Noradrenergic System Neurotransmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134 Brain Noradrenergic Neurotransmission and Arousal . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134 Tonic Versus Phasic Excitation of NE Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134 Brain Noradrenergic Neurotransmission, Stress, and Depression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135

4 4.1 4.2 4.3 4.4

Genetics of Noradrenergic Neurotransmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Tyrosine Hydroxylase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136 Dopamine b Hydroxylase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137 Mono‐Amine‐Oxydase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 138 Catechol‐O‐Methyltransferase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139

5

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 140

#

2008 Springer ScienceþBusiness Media, LLC.

130

6

Molecular genetics of brain noradrenergic neurotransmission

Abstract: The brain noradrenergic system plays a pivotal role in mediating the responses of the organism to the external and internal milieu and particularly to stress. The different components of this system have been implicated in a wide range of normal and pathological behavior. Recent advances in molecular psychiatric genetics may pave the way for a better understanding of the etiology of different symptoms of mental diseases and their relationship to environmental factors. List of Abbreviations: AAD, Aromatic Amino acid Decarboxylase; AC, adenylyl cyclase; ADs, antidepressant drugs; BMP, Bone Morphogenetic Proteins; COMT, catechol‐O‐methyltransferase; DAG, diacylglycerol; DOPA, 3,4 dihydroxyphenylalanine; DHPG, 3,4‐dihydroxyphenyl‐glycol; MHPG, 3‐methoxy‐4‐ hydroxyphenylglycol; fmri, functional magnetic resonance imaging; Gata3, GATA‐binding protein 3; GDP, guanosine diphosphate; GPCRs, G protein‐coupled receptors; GTP, guanosine triphosphate; HPA, hypothalamic‐pituitary‐adrenal; LC, Locus Coeruleus; MAOs, monoamineoxidases; NET, NE transporter; PKC, protein kinase C; PKA, protein kinases A; PNMT, Phenylethanolamine‐N‐Methyl‐Transferase; PLC, phospholipase C; PTSD, posttraumatic stress disorder; SSRIs, selective serotonin reuptake inhibitors; TH, Tyrosine Hydroxylase; VMA, vanyl‐mandelic acid; VCF, velocardiofacial syndrome; VMAT, vesicular monoamine transporter

1

Introduction

Adrenaline (L(
)‐epinephrine) and noradrenaline (L(
)‐norepinephrine) (NE), along with dopamine, are catecholamines, a class of molecules characterized by a catechol and an amine group, which belong with the indolamines (such as serotonin) to the monoamines group of compounds. They act as neurotransmitters as well as hormones, NE being the predominant neurotransmitter whereas epinephrine is the major hormone. Their names derive from the ancient Greek ‘‘epi’’ (upon) and ‘‘nephron’’ (kidney), since epinephrine is produced mainly by the adrenal glands located in the apex of the kidneys. Epinephrine released from the adrenal glands is carried in the bloodstream and acts upon the autonomous nervous system regulating a wide range of functions such as the heart rate, dilation of the pupils, glucose storage, intestinal motility, secretion of sweat, and salivary glands etc. In the brain the cell bodies that contain NE are found in the brainstem, and their projections extend from the forebrain to the spinal cord. These neurons are associated with the stress response and with the control of drive and motivation, alertness and sleep patterns, along with stress‐related manifestations such as anxiety and fear. Taken together the physiological reactions in the peripheral and central nervous system mediated by the adrenergic and NE‐ergic systems make up the substrate of the response to stress that is best illustrated by the ‘‘Fight or Flight’’ paradigm. The central role of the stress response in the homeostasis of the living organism has placed it also at the crossroads of several peripheral and central pathologies that are totally or partially related to inadequate, excessive, or prolonged activation of the mechanism underlying stress. In the brain, stress has been associated with such pathologies as depression and anxiety, but also bipolar disorder and schizophrenia (Berridge and Waterhouse, 2003).

2

Brain Noradrenergic System

2.1 Ontogeny Mash‐1, a bHLH domain protein, as well as Phox2a and Phox2b, two paralogous homeodomain proteins belonging to the Q50 paired‐like class, are transcription factors essential for the development of all central and peripheral neurons with a NE‐ergic phenotype. These factors have been implicated in synchronizing pan‐neuronal and catecholaminergic specific neurogenesis. Their expression domain is restricted, in the peripheral nervous system, to all autonomic ganglia (sympathetic, parasympathetic, and enteric) and the distal ganglia of the facial, hypoglossal, and vagus cranial nerves. In the central nervous system, it comprises all NE‐ergic centers, all cranial motor nuclei with the exception of the abducens and hypoglossal,

Molecular genetics of brain noradrenergic neurotransmission

6

the nucleus of the solitary tract and associated area postrema, as well as scattered interneurons in the hindbrain and spinal cord (Brunet and Pattyn, 2002). The cascade of evenements leading to the NE‐ergic phenotype encompasses upstream Mash‐1 and Phox2a/2b, the bone morphogenetic proteins (BMP) class of factors and, downstream, the bHLH transcription factor dHand (heart and neural‐crest derivatives expressed 2), and the zinc finger protein GATA‐binding protein 3 (Gata3). In the periphery, Mash‐1 appears to regulate, with Phox2b, the expression of dHand and Phox2a. These two factors may interact and regulate with Gata3 the expression of the tyrosine hydroxylase (TH) and dopamine‐b‐hydroxylase genes, which constitute the hallmarks of the NE‐ergic phenotype. In the brain, this pathway appears to be more straightforward forming a direct chain with the factors acting in the sequence BMP, Mash‐1, Phox2a, and Phox2b. This latter then regulates (with Phox2a?) the expression of the NE‐ergic synthesizing enzyme tyrosine hydroxylase and dopamine‐b‐hydroxylase (Goridis and Rohrer, 2002). Interestingly, point mutations in the HASH‐1 gene and a polyalanine expansion in the PHOX2B gene product have been associated with congenital central hypoventilation syndrome in humans, a rare breathing disorder accompanied by dysautomia manifestations, which is also known as Ondine’s curse (de Pontual et al., 2003).

2.2 Nuclei and Pathways All NE‐ergic neurons in the brain are located in the brainstem. In the pons they form the locus coeruleus (LC), and in the medulla they are found in the reticular formation, the solitary nucleus, and the dorsal motor nucleus of the vagus. The NE‐ergic system originates in a relatively small number of cells, but it innervates, by generating an extensive network of NE‐ergic projections, essentially the whole neuraxis from the olfactory bulb to the spinal cord. In this way, this system may potentially influence the activity of a widespread array of brain centers under conditions of elevated NE‐ergic activity. The projections originating from the LC form the dorsal noradrenergic bundle, a pathway that merges with the median forebrain bundle, the medial forebrain bundle, and the dorsal longitudinal fasciculum. The LC is at the origin of most of the innervation to the forebrain, with over 40% of its neurons projecting to the neocortex (frontal lobes, hippocampus, and olfactory bulbs), representing the most important NE‐ergic input related to psychological functions (Aston‐Jones and Cohen, 2005). Other projections are directed to the basal forebrain, the thalamus, and the cerebellum, whereas another important output is toward the brainstem sensory and association nuclei and all the levels of the spinal cord, mediating autonomic regulation of, for example, the cardiac activity or the axis hypothalamic–pituitary–adrenal (HPA) stress axis. Afferents to the LC are inhibitory GABA‐ergic inputs from the rostral medulla and the nucleus prepositus hypoglossi, and excitatory inputs, probably glutamatergic, from the ventromedial zone of the nucleus paragigantocellularis.

2.3 NE Synthesis Tyrosine is the common precursor in the synthesis of NE and adrenaline, as well as dopamine. Tyrosine is converted inside the nerve terminal to 3,4‐dihydroxyphenylalanine (DOPA) by tyrosine hydroxylase (TH), the rate‐limiting enzyme in the synthesis of catecholamines. DOPA is then decarboxylated by the aromatic amino acid decarboxylase (AAD or DOPA decarboxylase) to dopamine, which is then converted to NE by dopamine‐b‐hydroxylase. Thereafter, the phenylethanolamine‐N‐methyl‐transferase (PNMT) converts the NE to adrenaline in the adrenal medulla.

2.4 NE Storage and Release NE, as all monoamines, is concentrated in vesicles at the nerve terminal by a specific vesicular monoamine transporter (VMAT‐1 and VMAT‐2) (Njus et al., 1986). VMAT‐1 is primarily present in endocrine and paracrine cells of peripheral organs, whereas VMAT‐2 is the predominant monoamine vesicular transporter in the central nervous system (Masson et al., 1999).

131

132

6

Molecular genetics of brain noradrenergic neurotransmission

The calcium inflow triggered by the opening of voltage‐activated calcium channels upon the arrival of an action potential to the nerve terminal induces the fusion of the vesicle with the presynaptic membrane and the release of the neurotransmitter into the synaptic cleft. There it will act on specific receptors located both on the postsynaptic and presynaptic membranes. Stimulation of the postsynaptic receptors results in changes in the properties of the postsynaptic membrane with either a shift in membrane potential when the receptors are coupled with ion channels (ionotropic receptors) or in biochemical changes when the receptors are coupled with G proteins (metabotropic receptors) (Starke, 2001). Stimulation of the presynaptic receptors located on the nerve terminal will regulate the transmitter release triggered by the action potential, providing, therefore, a feedback mechanism for the control of the neurotransmitter’s concentration in the synaptic cleft (Boehm and Kubista, 2002).

2.5 NE Receptors NE and adrenaline exert their cellular action via binding to specific membrane proteins, the adrenergic receptors or adrenoceptors. The adrenergic receptors are classified into two main categories, alpha and beta, and can be divided into three main classes based on sequence similarity, receptor pharmacology, and signaling mechanisms: alphal (a1), alpha2 (a2), and beta (b). All three classes are present in the brain. Further subdivisions exist within each class (Bylund, 1992; Bylund, 2005). All adrenergic receptors belong to the family of G protein‐coupled receptors (GPCRs), the largest single family of integral membrane receptors. It has been calculated that 3–4% of the genes encodes a member of this family. GPCRs are characterized by seven transmembrane domains and are coupled to heterotrimeric—since they are composed of three subunits (a, b, and g)—guanine nucleotide‐binding proteins (G proteins). The binding of the agonist to the receptor induces a conformational change characterized by a high affinity agonist state that turn G proteins on by promoting the binding of the activating nucleotide guanosine triphosphate (GTP) in exchange for guanosine diphosphate (GDP) on the a subunit of the G protein. This activation of the specific G protein initiates a cascade of molecular events resulting in the positive or negative regulation of the effectors systems (Neer, 1995). The G proteins are named after their a subunit and are divided into four subgroups: Gs and Gi/o, which stimulate and inhibit, respectively, adenylate cyclase, Gq/11, which stimulates phospholipase C (PLC), and the less characterized G12/13 subgroup, which activates the Naþ/Hþ exchanger pathway (Gether, 2000). The distinct classes of adrenergic receptors are differently coupled to these subgroups. The b‐adrenergic receptor, which presents three subtypes (b1, b2, and b3) and is exclusively postsynaptic, is coupled to the Gs proteins. The stimulation of the b‐adrenergic receptor leads to the prototypic cellular effect of Gs proteins, which is activation via stimulation of adenylyl cyclase (AC) resulting in the accumulation of the second messenger cyclic adenosine monophosphate (c‐AMP) and subsequent activation of the c‐AMP‐dependent protein kinases A (PKA). PKA causes the phosphorylation of various cellular proteins, which produce the specific responses of b‐adrenergic receptor stimulation (Taussig and Gilman, 1995). Molecular cloning has identified four different subtypes of a2 receptors designated as a2A, a2B, a2C, and a2D (Bylund et al., 1994). The a2 receptors are both presynaptic and postsynaptic and the a2A receptor appears to be far more represented in the brain than in the sympathetic nervous system (Boehm and Kubista, 2002). The a2 adrenergic receptors are coupled to the Gi/o protein family whose activation results in the inhibition of c‐AMP accumulation (Neer, 1995). The a2 adrenergic receptors mediate a hyperpolarization of the neuronal membrane, which renders the neuron less excitable by increasing Kþ conductance via activation of G protein‐gated Kþ channels. By their localization and action the presynaptic a2 receptors regulate the synthesis and release of NE. The al adrenergic receptors (comprising the 1A, 1B, 1C, and 1D subtypes) are generally excitatory in nature and are usually coupled to the Gi/Gq family of G proteins. The Gi/Gq proteins are linked to a signaling cascade different from the AC pathway. They activate the phosphatidylinositol‐specific PLC, which subsequently generates the second set of messengers, inositol triphosphate (IP3) and diacylglycerol (DAG). IP3 stimulates the release of Ca2þ from intracellular stores via a specific receptor‐mediated process,

Molecular genetics of brain noradrenergic neurotransmission

6

thus increasing available intracellular calcium that will be involved in the regulation of several protein kinases (Berridge, 1993). DAG is a potent activator of protein kinase C (PKC), which is involved in the activation of many substrates including membrane proteins such as channels, pumps, and ion exchange proteins (Fields and Casey, 1997). Stimulation of the membrane PLC leading to the formation of the second messengers IP3 and DAG through activation of the al receptors on central adrenergic target neurons produces depolarizing responses due to a decrease in Kþ conductance, thus making the neuron more excitable. The activation of a GPCR not only results in the G protein‐dependent activation of effector systems but also allows for feedback regulation of G protein coupling, receptor endocytosis, and signaling through G protein independent signal transduction pathways. Therefore, GPCR activity represents a coordinated balance between molecular mechanisms governing receptor signaling, desensitization, and resensitization as well as downregulation. Desensitization is the consequence of the uncoupling of the receptor from heterotrimeric G proteins in response to receptor phosphorylation by specific kinases (GRKs). This phosphorylation favorizes the interactions with G proteins to mediate the effects of receptor stimulation and, at the same time, target the receptor for the binding of the arrestin class of proteins. The arrestins bind to the phosphorylated receptor on the cytoplasmatic side and block further G protein activation, operating in this way to desensitize the receptor. Arrestins act subsequently as a molecular adaptor in binding with clathrin and the clathrin adaptor complex AP‐2. This produces the internalization by endocytosis of cell surface receptors and their sequestration in intracellular membranous compartments. Then the sequestered receptors can be recycled, producing resensitization, or directed to the lysosomial apparatus for degradation, resulting in a downregulation, which is a reduction of the total number of receptors. Conversely, after reduction in the chronic level of receptor stimulation by chronic antagonist administration or denervation, hyperreactivity and upregulation of the GPCRs are stimulated (Ferguson, 2001). The pivotal role played by arrestins in a NE‐ergic transmission, and in general in the activity of all GPCRs, is not limited just to the extinguishing of the signal. They have been shown to act also as a multifunctional adaptor and as scaffolding proteins implicated in recruiting signaling molecules. In mammals there are two types of arrestins: the visual arrestins that are mainly restricted to photoreceptor cells, and the b‐arrestins (1 and 2) that are ubiquitously expressed. Upon recruitment following agonist‐ induced phosphorylation of GPCRs by GRKs, the b‐arrestins, in addition to their role in the repression of signaling, may participate in the signal transduction acting as scaffolds for recruiting phosphatases and kinases, such as, for example, AKT and ERK (Hubbard and Hepler, 2006). In this perspective, b‐arrestins are essential for the AKT signaling involved in the Dopamine D2 receptor‐mediated behavior in mice (Beaulieu et al., 2004). Moreover, the action of b‐arrestins may not be limited to the role of a cytoplasmic adaptor in a signaling cascade, but may also intervene directly in gene regulation. b‐Arrestin1 has been found, after GPCR stimulation, in the nucleus as part of the nuclear complex formed, due to the promoter of target genes, by the transcription factor CREB and the histone acetyl transferase protein P300 (Kang et al., 2005).

2.6 Termination of NE Synaptic Transmission 2.6.1 Reuptake The synaptic activity of NE, as with the actions of all monoamines, is terminated by its active reuptake into the presynaptic neuron (known as uptake1) and/or glial cells. The uptake one mechanisms utilize Naþ/ Cl
‐dependent transporters (Lester et al., 1996). These transporters are members of a large family of Naþ/Cl
containing putative transmembrane domains, which control the concentration of the transmitter released into the intraplasmatic and extrasynaptic spaces via rapid reuptake into the nerve terminals, thus maintaining low concentration of the neurotransmitter at these sites (Masson et al., 1999). Since transporter velocity is increased by hyperpolarization of the membrane and is decreased by depolarization, autoreceptors or heteroreceptors affect the activity of the reuptake system by changing the membrane potential.

133

134

6

Molecular genetics of brain noradrenergic neurotransmission

The NE transporter (NET), like the serotonin or dopamine transporters, is a target for antidepressant drugs (ADs) and psychostimulant drugs. The NET is regulated by a wide range of intracellular signaling molecules such as PKC, c‐AMP/PKA, and CaM Kinase (Mandela and Ordway, 2006).

2.6.2 Metabolism The two enzymes that are important in the initial steps of metabolic transformation of NE and the other catecholamines are the monoamineoxidases (MAOs) and the catechol‐O‐methyltransferase (COMT). Two isoforms of MAO (MAO‐A and MAO‐B) are differentiated on the basis of substrate and inhibitor specificities. The preferential metabolizer of NE (and serotonin) is the MAO‐A whose specific inhibitor is clorgyline. Neurons contain both isoforms of MAOs, localized primarily in the outer membrane of mitochondria (Shih et al., 1999). Inhibitors of MAO cause an increase in the amount of monoamines stored and released from the nerve terminals, thus increasing their availability at the synapsis. COMT is bound to membranes and appears to be located principally in postsynaptic neurons. Both MAO and COMT enzymes, starting with either one of them, sequentially coordinate the degradation of NE. They generate aldehyde intermediates, which are reduced to 3,4‐dihydroxyphenyl‐glycol (DHPG) and 3‐methoxy‐4‐hydroxyphenylglycol (MHPG) by cytosolic aldehyde reductase or oxidized to vanyl‐mandelic acid (VMA) by mitochondrial aldehyde dehydrogenase (Eisenhofer et al., 2004).

3

Functional Noradrenergic System Neurotransmission

The postsynaptic effects of NE, at the cellular or neural circuit level, are essentially modulatory in nature. NE, rather than inducing simple inhibition or excitation, facilitates responses evoked in target cells by altering the ‘‘signal to noise ratio’’ (Servan‐Schreiber et al., 1990) of responses evoked by other afferents, both excitatory and inhibitory, thus enhancing synaptic transmission in target circuits (Woodward et al., 1991).

3.1 Brain Noradrenergic Neurotransmission and Arousal The involvement of the LC in the brain arousal system is essential for mediating attention in processes related to successful behavior. The LC is implicated in either selective (i.e. focused for rapid adaptative responses) or labile attention (Aston‐Jones et al., 1999). The LC innervates many more cerebral areas than any other brain nucleus, but it is also extremely specific in this innervation, which is most dense in areas associated with attention such as the parietal cortex, the pulvinar nucleus, and the superior colliculus. An overly focused or labile attention is a pathological behavior, which is a component of several neuropsychiatric diseases such as schizophrenia, ADHD, OCD, and depression.

3.1.1 Tonic Versus Phasic Excitation of NE Neurons NE is able to increase the evoked either excitatory or inhibitory activity while decreasing the spontaneous activity of its target neurons. NE plays a more specific role in regulating motivation and arousal, nonspecific aspects of behavior that are related in task‐related cognitive processes. In this aspect LC neurons display a stimulus‐specific activity similar to that of brainstem dopaminergic neurons responding selectively to stimuli that predict reward (Schultz et al., 1997). As observed in monkeys performing a visual discrimination task, LC neurons show short‐latency stimulus‐evoked (phasic) responses to target stimuli correlated with the corresponding behavioral responses but not to distractor stimuli or other task events. Thus, optimal behavioral performance is correlated with phasic activation of LC neurons. In contrast, periods of poor performance are associated with significantly higher tonic LC activity (Aston‐Jones et al., 1994). However, the LC tonic activity is necessary for an adaptative phasic activation. The LC tonic activity is high in waking, slow in sleeping, and almost absent during paradoxical sleep. The relationship between tonic and phasic activity related to behavioral performances corresponds graphically to an inverted‐U. The increase in

Molecular genetics of brain noradrenergic neurotransmission

6

phasic activation of the LC coupled with an intermediate tonic activity results in an excellent behavioral performance, whereas an increase or a decrease of the LC tonic activity diminishes the phasic activity with a concomitant reduction of the attentional performances. Thus, phasic LC discharge is related to behavioral responses and the level of LC tonic activity to behavioral performance (Aston‐Jones and Cohen, 2005).

3.2 Brain Noradrenergic Neurotransmission, Stress, and Depression Chronic stress has been implicated as a predisposing and/or triggering element for neuropsychiatric diseases such as major depression, bipolar disorder, and schizophrenia (Schildkraut, 1965) and has an important impact on the catecholaminergic system (Sabban and Kvetnansky, 2001) and in particular the NE‐ergic system (Morilak et al., 2005). Stress can be defined as any menace either real or perceived to the homeostasis and wellbeing of an organism. In the first case, stress is systemic or physiologic and the physical characteristics of the stressor require an immediate restorative response from the organism. Awareness or perception of the stimulus is not required to initiate the stress response. The second form is psychogenic stress, which depends upon perception, cognitive processing, and interpretation of the stimulus to confer upon it a stressful quality. In this case, the relative severity of the stressor and its physiologic impact is variable between individuals. The severity of stress presented by a stimulus, whether physiologic or psychogenic, has typically been defined in terms of the magnitude of the physiological response it elicits, for instance by measuring activation of the hormonal HPA stress axis, or of the peripheral sympathoadrenal autonomic response system. Stress in the experimental animal (restrain or foot shock) increases the firing in the LC and, in parallel, also the MHPG, which is a reliable index of the NE turnover. The inescapable stress not only induces a condition of ‘‘learned helplessness,’’ which is a preclinical correlate of depression, but also anxiety. Disregulation of NE‐ergic neurotransmission has been implicated in stress‐related psychiatric diseases such as depression, posttraumatic stress disorder (PTSD), and other anxiety disorders (Southwick et al., 1993; Sullivan et al., 1999; Leonard et al., 2001). Stress is able to deplete NE, and depletion of NE or repeated stress in rats upregulates TH in the LC (Melia and Duman, 1991; Melia et al., 1992). Elevated levels of TH have been found in the LC of victims of suicide (Ordway, 1997) and in patients with major depression as compared with normal controls (Zhu et al., 1999), whereas the NE transporter is downregulated in major depression (Klimek et al., 1997). Furthermore, the impairment of the NE‐ergic input from locus coeruleus to the forebrain limbic system may contribute to the symptoms of schizophrenia (Yamamoto and Hornykiewicz, 2004). In fact, these NE‐ergic projections are normally essential for screening and filtering the incoming sensory stimuli to discard irrelevant information (Archer, 1982). Alterations in NE‐ergic neurotransmission are important in the actions of many classes of antidepressant drugs (ADs), such as MAO inhibitors and tricyclic antidepressants, these latter blocking the reuptake of both NE and serotonin (Nelson, 1999). Serotonin in particular has been heralded as the main player in depression, since inhibition of its synthesis counteracts the effects of antidepressants, and selective serotonin reuptake inhibitors (SSRIs) are highly effective in the regulation of mood (Fuller et al., 1995). However, the observation that the extremely selective NE reuptake inhibitor reboxetine is an effective and powerful antidepressant has reactualized the role of NE in depression (Schatzberg, 2000). Activation of NE release in the limbic forebrain by acute stressors may facilitate anxiety‐like behavioral responses, making anxiety a prominent component of depression. The new generation of dual uptake inhibitors, as well as selective NE reuptake inhibitors, alleviate depressed mood, social withdrawal, and other symptoms of depression including anxiety (Nelson, 1999; Versiani et al., 2002; Ferguson et al., 2003; Morilak et al., 2005).

4

Genetics of Noradrenergic Neurotransmission

The role played by central NE‐ergic neurotransmission in behavior and its implication in related disturbances have prompted, with the advent of positional cloning, the investigation of linkage or association to neuropsychiatric diseases of the genes encoding the components of this system. These studies have been hampered by the genetic complexity inherent to psychiatric diseases, which are characterized by

135

136

6

Molecular genetics of brain noradrenergic neurotransmission

heterogeneity, polygenicity, and the interplay between genetic and environmental factors in such a way that no clear‐cut mutations but only predisposing polymorphism are implicated in their etiology. Taking into account these caveats, the most relevant results in this domain have been obtained for the TH, DBH, MAO, and COMT genes.

4.1 Tyrosine Hydroxylase The TH gene, located in the chromosome 11p15, encodes the rate‐limiting enzyme in the synthesis of catecholamines, and is thus a strong candidate gene for neuropsychiatric diseases (Mallet, 1996). A first report showing a genetic linkage in the Amish population between bipolar disorder and markers at the chromosomal region that contains the TH locus also put forward the TH as a ‘‘positional’’ candidate gene (Egeland et al., 1987). This result was questioned because the lod score method utilized did not take into account genetic heterogeneity that characterizes bipolar disorder and other complex diseases (Hodgkinson et al., 1987), and neither further genetic analysis of the Amish nor studies of other populations confirmed this initial result (Detera‐Wadleigh et al., 1987; Kelsoe et al., 1989; Ginns et al., 1992; Gerhard et al., 1994; Gershon et al., 1996; Ginns et al., 1996). However, other studies finding significant linkage between markers at the TH locus and bipolar disorder maintained the implication of the TH gene in this disease and strengthened the case for genetic heterogeneity (Pakstis et al., 1991; Byerley et al., 1992; Lim et al., 1993; Gurling et al., 1995; Smyth et al., 1996; Malafosse et al., 1997). In the first association study of the TH gene, a significant genetic association was found between restriction fragment polymorphism markers at the TH locus and bipolar disorder in a French population sample (Leboyer et al., 1990), a result that was not always replicated in other studies (Korner et al., 1990, 1994; Gill et al., 1991; Inayama et al., 1993; Kawada et al., 1995). Another association analysis was conducted using the more informative microsatellite HUMTH01 marker in order to further investigating the implication of the TH gene in the genetic predisposition to bipolar disorder. This microsatellite is a polymorphic polypyrimidine sequence localized in the first intron of the TH gene and is characterized by a core (TCAT)n tetranucleotide repeat iterated usually between 5 and 10 times (Polymeropoulos et al., 1991; Puers et al., 1993; Brinkmann et al., 1996). In a new sample of French case–controls, a significant genotypic association was found between the HUMTH01 and bipolar disorder as well as familial history of bipolar disorder and/or delusive symptoms during manic or depressive episodes (Meloni et al., 1995a). Moreover, a rare allele of the HUMTH01 microsatellite was significantly associated with schizophrenia in two different ethnic samples from Normandy in northwestern France and the Sousse region of eastern Tunisia (Meloni et al., 1995b). Several further studies inspired by these results have been inconclusive for association (Cavazzoni et al., 1996; Souery et al., 1996, 1999; Turecki et al., 1997; Burgert et al., 1998; Jonsson et al., 1998) or have replicated the positive association between this microsatellite and both bipolar disorder (Perez de Castro et al., 1995; Lobos and Todd, 1997; Serretti et al., 1998a, b; Furlong et al., 1999; Chiba et al., 2000) and schizophrenia (Wei et al., 1995, 1997; Kurumaji et al., 2001). It is noteworthy that the HUMTH01 microsatellite is associated with other physiological or behavioral traits such as personality (Persson et al., 2000), longevity (De Benedictis et al., 1998), symptoms of alcohol withdrawal (Sander et al., 1998), hypertension (Jindra et al., 2000), and stress response parameters in twins (Zhang et al., 2004), but also directly with catecholamine neurotransmission. This is shown by measuring catecholamine metabolite levels in cerebrospinal fluid (Jonsson et al., 1996) or in plasma (Wei et al., 1995, 1997) as an indirect index of their turnover in the brain. Moreover, in a clinical study in the original Normandy sample the schizophrenic patients bearing the rare allele associated with schizophrenia presented significantly lower plasma concentrations of the catecholaminergic metabolites MHPG and HVA, which are indices of central NE‐ergic and DA‐ergic function, respectively, as compared with patients bearing other alleles (Thibaut et al., 1997). These results suggest a functional link between allelic variations at the HUMTH01 marker and TH activity. Indeed, the (TCAT)n motif of this microsatellite differs by only one nucleotide from the consensus AP1 sequence (TGATTCA) present in the rat and the human TH gene (Icard Liepkalns et al., 1992), a sequence that is specifically recognized by transcription factors of the Fos and Jun proto‐onco‐gene families

Molecular genetics of brain noradrenergic neurotransmission

6

(Sassone‐Corsi et al., 1988). Moreover, a less polymorphic HUMTH01 repeated sequence is conserved at its orthologous position in the first intron of the TH gene in several non‐human primate species (Meyer et al., 1995), suggesting that this motif may be an evolutionary conserved regulatory element that has expanded in the human lineage. Therefore, the functional role of the HUMTH01 microsatellite was assessed to investigate the biological significance of the genetic association findings. Indeed, the alleles of this microsatellite enhanced transcription when placed upstream from a minimal promoter driving the expression of a luciferase‐reporter gene. Moreover, these repeated sequences interacted specifically with factors of the fos/jun type and with an even higher affinity with other nuclear proteins (Meloni et al., 1998). Subsequently, ZNF191, a zinc finger protein, and HBP1, a HMG box transcription factor, were identified as the proteins specifically binding the TCAT motif. Interestingly, the specific binding of ZNF191 to the HUMTH01 sequence was correlated in a quantitative fashion to the number of TCAT repeats. Moreover, in vitro experiments with a TH‐reporter gene construct established that the HUMTH01 microsatellite regulates the TH gene expression by a quantitative silencing effect that correlates with the number of repetitions of the (TCAT) motif (Albanese et al., 2001). Thus, the HUMTH01 sequence may participate in the transcriptional regulation of the TH gene by modulating its expression in a quantitative fashion. It is noteworthy that HBP1 was characterized as a chromatin remodeling factor, which binds specifically a TCAT short repeat in the locus control region of the CD2 gene (Zhuma et al., 1999). Moreover, a polypyrimidine trait similar to the TCAT repeated motif has been shown to regulate the expression of the CD30 gene (Croager et al., 2000). Finally, a TCAT stretch is the only difference that characterizes the regulatory region of the vasopressin receptor gene that accounts for a different distribution of the gene product in a brain region of two different species of voles. This single difference is responsible for completely different mating and parental behavior between species (Young et al., 1999). Since the (TCAT)n polymorphic sequence is widespread in the genome and present in several genes, it may provide a molecular basis for the modulation of gene expression relevant to the genetics of quantitative traits.

4.2 Dopamine b Hydroxylase The DBH gene is located on chromosome 9q34 (Craig et al., 1988) and encodes the enzyme that catalyzes the conversion of DA to NE. Mutations in the DBH gene result in a lack of sympathetic NE‐ergic function and orthostatic hypotension (Garland et al., 2002; Deinum et al., 2004). The DBH enzyme is localized within the soluble and membrane fractions of secretory catecholamine‐containing vesicles of NE‐ergic and adrenergic cells. These two forms originate from optional cleavage of the signal peptide. The form retaining the signal peptide is completely associated with the membrane, whereas the cleaved form is mostly soluble (Houhou et al., 1995). The soluble form of the enzyme is secreted into the circulation from nerve terminals allowing for assaying its activity in plasma or serum. DBH presents stable differences in enzymatic activity that appear to be genetically determined (Stolk et al., 1982). Several polymorphisms in the human gene have been implicated in these variations. A G/T polymorphism at the nucleotide 910 of the coding sequence results in a change of amino acid residue 304 between Ala (A) and Ser (S) (DBH/A and DBH/S). The resulting proteins have similar kinetic constants, but DBH/S has a homospecific activity that is about 1/13 lower than that of DBH/A (Ishii et al., 1991). In European, African, and several other populations the DBH/ A allele is the most common with allele frequencies greater than 0.80 in each sample and significant heterogeneity in allele frequency across population groups (Cubells et al., 1997). These allelic differences cannot alone account for the differences in the activity of DBH in blood since circulating DBH concentrations also vary considerably in the general population. Recently, a novel polymorphism (
1021 C/T) in the 50 promoter region of the DBH gene was shown to strongly influence plasma DBH activity, accounting for 35–52% of its variation in different populations (Zabetian et al., 2001; Kohnke et al., 2002). A further study showed that ten biallelic markers in a 10 Kb surrounding the
1021C/T polymorphisms were all associated with plasma DBH activity and that this association was strongly correlated with the degree of Linkage Disequilibrium between each marker and the
1021C/T polymorphism (Zabetian et al., 2003). Another association was found between a DBH TaqI polymorphism and plasma metabolites of catecholamines

137

138

6

Molecular genetics of brain noradrenergic neurotransmission

(Wei et al., 1998). Other polymorphic variants in the DBH gene are represented by a GT dinucleotide microsatellite, a single‐base, 444 g/a, substitution at the 30 end of DBH exon 2 and a diallelic variant, DBH50 ‐ins/del, located approximately 3 kb 50 to the DBH transcriptional start site. All these markers, which are in linkage disequilibrium, were also associated with plasma DBH activity (Wei et al., 1997; Cubells et al., 1998, 2000; Jo¨nsson et al., 2004). Moreover, the 444 g/a marker was also associated with differences in DBH concentration in the CSF (Cubells et al., 1998; Zabetian et al., 2003). Psychiatric genetic studies using the polymorphisms at the DBH gene have shown a significant association between the DBH TaqI polymorphism and ADHD (Daly et al., 1999). Also, albeit it did not reach statistical significance, the DBH GT repeat four allele, which is associated with high serum levels of DBH, occurred more frequently in the ADHD group than controls (Mu¨ller Smith et al., 2003). However, other groups failed in replicating these results either with the same (Wigg et al., 2002) or other markers (Hawi et al., 2003). A positive association was shown between the DBH 50 ‐444a haplotype and cocaine‐ induced paranoia (Cubells et al., 2000) as well as a lack of response to antipsychotic drug treatment in schizophrenic patients (Yamamoto et al., 2003). These results may indicate that the DBH gene is indirectly involved in schizophrenia as a modulatory factor of psychotic symptoms, severity of the disorder, and therapeutic response to neuroleptic drugs.

4.3 Mono‐Amine‐Oxydase The MAO‐A and MAO‐B genes are situated on the X chromosome at Xp11.23–11.4 and result from the duplication of a common ancestral gene. In humans both genes are deleted in patients with Norrie’s disease, a rare X‐linked recessive neurological disorder characterized by blindness, hearing loss, and mental retardation (Lan et al., 1989). A point deletion resulting in a complete MAO‐A inactivation was linked to abnormally aggressive behavior in the males of a Dutch family (Brunner et al., 1993). Conversely, MAO‐A knock‐out mice show an increased aggressivity in males, which is related to increased levels of NE and serotonin during development and result in brain structural changes (Cases et al., 1995). A functional polymorphism located in the MAO‐A gene promoter 1.2kb upstream of the encoding sequence consists of a 30bp repeated sequence present in 3, 3.5, 4, or 5 copies. This polymorphism displays significant variations in allele frequencies across ethnic groups and is able to affect the transcriptional activity of the MAO‐A gene promoter (Sabol et al., 1998). Genetic studies with this polymorphism have found that the high‐activity MAO‐A gene promoter alleles were associated with panic disorder (Deckert et al., 1999) and major depressive disorder (Schulze et al., 2000) in females, whereas the low‐activity alleles were associated with schizophrenia in males (Jonsson et al., 2003). Other studies have yielded negative results for panic disorders (Hamilton et al., 2000), schizophrenia (Syagailo et al., 2001; Fan et al., 2004), and mood disorders (Kirov et al., 1999; Kunugi et al., 1999; Jorm et al., 2000; Syagailo et al., 2001; Huang et al., 2004). However, more probant results have been found when genetic studies have taken into account an environmental component. A leading study has shown that this functional polymorphism can modulate the association between childhood maltreatment and subsequent antisocial behavior. In males, who have only one copy of the X chromosome, the MAO‐A functional polymorphism confers either a high or a low‐activity genotype. The low‐activity MAO‐A genotype is associated with antisocial behavior in up to 85% of a cohort of males who had been severely maltreated in their childhood but not in boys who had suffered little or no abuse. In contrast, the high‐activity MAO‐A genotype has a protective effect from developing antisocial behavior in maltreated children. In women a similar trend for association between MAO‐A genotype, antisocial behavior and child maltreatment, was present, but the interpretation of this result was hindered by the presence of two X chromosomes with sometimes heterozygous low/high MAO‐A activity alleles, one of which is randomly inactivated. However, taken together, these results show a clear influence of the MAO‐A genotype in the behavioral effects of an environmental factor and may help in understanding the marked differences in the frequency of antisocial behavior between sexes (Caspi et al., 2002). The association between the lower expression MAO‐A genotype and antisocial behavior consequent to childhood maltreatment has been replicated

Molecular genetics of brain noradrenergic neurotransmission

6

(Foley et al., 2004), and this risk genotype has also been associated with impulsive traits in males who have experienced early abuse (Huang et al., 2004) and with pathological gambling always in males (Ibanez et al., 2000).

4.4 Catechol‐O‐Methyltransferase The COMT gene, localized to chromosome 22q11.1–q11.2, encodes a soluble (S‐COMT) and a membrane‐ bound (MB‐COMT) form of the enzyme, the latter characterized by an additional 50 amino acids at the N‐terminal (Bertocci et al., 1991; Grossman et al., 1992). The two length variants of the COMT are expressed from two mRNA transcripts: a long mRNA, which is able to translate both S‐COMT and MB‐COMT from two different initiation sites, and a short mRNA producing S‐COMT only. The long mRNA and the larger MB‐COMT are predominant in the brain whereas the short mRNA and the S‐COMT prevail in the other tissues (Tenhunen et al., 1994; Lundstrom et al., 1995). The COMT enzymatic activity shows high, intermediate, and low rates consistent with inheritance of two codominant alleles (Weinshilboum, 1978). This difference in enzyme activity is independent of protein length variations but is caused by an amino acid substitution. A G/A polymorphism in exon 4 at position 472 in the long mRNA, and 322 in the short mRNA, results in a Val to Met amino acid change at codon 158 of MB‐COMT and codon 108 of S‐COMT. The G (Val) allele encodes the thermostable, high‐activity form of the enzyme, whereas the A (Met) allele encodes the thermolabile, low‐activity variant that exhibits a 3‐ to 4‐fold decrease in the enzymatic activity level (Lotta et al., 1995; Lachman et al., 1996a, b). The G (Val) and A (Met) alleles correspond also to the absence or presence, respectively, of a NlaIII polymorphic restriction site that allows for easily genotyping the functional variations (Karayiorgou et al., 1998). COMT is an obvious a priori candidate gene for neuropsychiatric disorders that involve dopaminergic or NE‐ergic systems (Palmatier et al., 1999) but also a strong positional candidate gene for schizophrenia because of its chromosomal location in the locus of the velocardiofacial (VCF) syndrome. Microdeletions of 22q11 are associated with VCF syndrome which is characterized by congenital abnormalities, learning difficulties, and psychosis in up to one third of patients. Conversely, the deletion is also 80‐fold more common in patients with psychosis than the normal population (Sugama et al., 1999). Both linkage and association studies have implied that chromosome 22q11 is a locus for schizophrenia (Pulver et al., 1994a, b; Karayiorgou et al., 1995; Karayiorgou and Gogos, 1997). The case–control association approach has consequently been used to study the role of COMT in schizophrenia and other psychiatric diseases, mostly using the Val108/158Met polymorphism. Positive associations have been found between COMT and schizophrenia (Ohmori et al., 1998; de Chaldee et al., 1999), violence in schizophrenia (Lachman et al., 1998), bipolar disorder (Li et al., 1997; Mynett‐Johnson et al., 1998), unipolar disorder (Ohara et al., 1998), bipolar disorder or ADHD in VCF syndrome patients (Lachman et al., 1996a), OCD (Karayiorgou et al., 1997), drug abuse (Vandenbergh et al., 1997), and Parkinson’s disease (Kunugi et al., 1997a). However, other studies have excluded a major contribution of the COMT gene to schizophrenia (Daniels et al., 1996; Chen et al., 1997; Strous et al., 1997; Karayiorgou et al., 1998; Wei and Hemmings, 1999), bipolar disorder (Craddock et al., 1997; Gutierrez et al., 1997; Kunugi et al., 1997b; Lachman et al., 1997; Geller and Cook, 2000), ADHD or bipolar disorder in VCF syndrome patients (Lachman et al., 1996b), substance abuse and violence (Vandenbergh et al., 1997; Lachman et al., 1998), as well as Parkinson’s disease (Hoda et al., 1996; Syvanen et al., 1997; Xie et al., 1997). These conflicting results have prompted a meta‐analysis indicating that the COMT Met allele that characterizes the instable form of the enzyme with low‐activity phenotype is not associated with schizophrenia (Lohmueller et al., 2003). However, a new association study conducted in a genetically homogeneous population yielded a highly significant association between a COMT haplotype and schizophrenia (Shifman et al., 2003). This study is the largest case/control analysis in schizophrenia that has been reported with more than 700 patients and 4,000 control subjects. Genotyping was conducted using 12 SNPs, comprising the Val108/158Met polymorphism, across the COMT gene, and haplotypes with seven of these SNPs were established in the large sample of an Israeli Ashkenazi Jewish population. This population has the advantage of presenting a founder effect that allows for reducing genetic heterogeneity thus increasing gene effect and avoiding false‐positive results due to population stratification.

139

140

6

Molecular genetics of brain noradrenergic neurotransmission

The association between schizophrenia and the Val108/158Met polymorphism was moderate, but extremely high levels of statistical significance were attained when this marker was analyzed as part of a haplotype including two other noncoding SNPs that were more significantly associated with schizophrenia. Moreover, one of these polymorphisms represented a higher risk factor essentially for women than men, hinting at a possible sex‐specific genetic component in schizophrenia. These results confirmed a complex association of the COMT locus to schizophrenia and suggested that other functional variants besides the Val108/158Met polymorphism are likely to be involved in susceptibility to the disease (Shifman et al., 2003). A significant association was also found between bipolar disorder and both the allele and haplotype of the COMT gene and they were previously found to be associated with schizophrenia (Shifman et al., 2004). In addition to these association studies, the role of COMT in schizophrenia and other neuropsychiatric diseases is further supported by functional genetic studies that have essentially focused on the Val108/ 158Met polymorphism. A large amount of experimental data suggests that heritable abnormalities of prefrontal cortex function are a prominent feature of schizophrenia (Grace, 1991, 1993; Moore et al., 1999). COMT may constitute a major contributor to these abnormalities by virtue of its unique role in regulating catecholamine‐mediated prefrontal information processing, since COMT inhibitors can improve working memory in both rodents and humans (Weinberger et al., 2001). In this perspective, a study combining a genetic and a functional approach has shown that the Val allele of the Val108/158Met polymorphism that characterizes the high‐activity form of the COMT occurs at higher rates in both schizophrenics and their unaffected siblings. Moreover, patients and siblings bearing this allele performed poorly on the Wisconsin card sorting test (a neuropsychological test of frontal lobe function for working memory) and manifested inefficient brain activation as assessed by functional magnetic resonance imaging (fMRI) (Egan et al., 2001). Interestingly, amphetamine, a drug that increases catecholaminergic neurotransmission, enhances the efficiency of prefrontal cortex function as assayed with fMRI during a working memory task in subjects with the high‐activity val/val genotype but not in subjects with the low‐activity met/met genotype (Mattay et al., 2003). Moreover, this polymorphism is also associated with personality traits, as assessed by the tridimensional personality questionnaire (Benjamin et al., 2000). In addition, homozygosity for the Met allele is associated, particularly in schizophrenic patients, with lower frontal P300 amplitudes, which is an index of catecholaminergic efficacy in reducing noise during information processing (Gallinat et al., 2003). In agreement with these findings and with the results of the association studies in Ashkenazi Jews (Shifman et al., 2003, 2004), the analysis of the allele‐specific expression using mRNA from human brains indicated that the haplotype implicated in schizophrenia and bipolar disorder is associated with lower expression of COMT mRNA (Bray et al., 2003). These findings suggest that the COMT Val allele impairs prefrontal cognition and physiology and, by virtue of this effect, may condition some pathological features of schizophrenia thus contributing, with other sequence variations at the COMT locus, to the increase of the risk for schizophrenia or other psychiatric disorder characterized by impaired frontal cortex functioning.

5

Conclusions

The brain NE‐ergic system plays a pivotal role in integrating and fine‐tuning the adaptative responses to basic arousal and stress‐generated stimuli. Anatomical, functional, and genetic studies may further contribute to the understanding of how the mechanism underlying these responses may intervene in normal and pathological behavior and yield new entries for therapeutic interventions in related psychiatric diseases.

References Albanese V, Biguet NF, Kiefer H, Bayard E, Mallet, J, et al. 2001. Quantitative effects on gene silencing by allelic variation at a tetranucleotide microsatellite. Hum Mol Genet 10: 1785-1792. Archer T. 1982. The role of noradrenaline in learned behaviors: Studies using DSP4. Scand J Psychol Suppl 1: 61-71.

Aston‐Jones G, Cohen, JD. 2005. An integrative theory of locus coeruleus‐norepinephrine function: Adaptive gain and optimal performance. Annu Rev Neurosci 28: 403-450. Aston‐Jones G, Rajkowski J, Cohen J. 1999. Role of locus coeruleus in attention and behavioral flexibility. Biol Psychiatry 46: 1309-1320.

Molecular genetics of brain noradrenergic neurotransmission Aston‐Jones G, Rajkowski J, Kubiak P, Alexinsky T. 1994. Locus coeruleus neurons in monkey are selectively activated by attended cues in a vigilance task. J Neurosci 14: 4467-4480. Beaulieu J‐M, Sotnikova TD, Yao W.‐D, Kockeritz L, Woodgett JR, et al. 2004. Lithium antagonizes dopamine‐ dependent behaviors mediated by an AKT/glycogen synthase kinase 3 signaling cascade. Proc Natl Acad Sci USA 101: 5099-5104. Benjamin J, Osher Y, Kotler M, Gritsenko I, Nemanov L, et al. 2000. Association between tridimensional personality questionnaire (TPQ) traits and three functional polymorphisms: Dopamine receptor D4 (DRD4), serotonin transporter promoter region (5‐HTTLPR) and catechol‐ O‐methyltransferase (COMT). Mol Psychiatry 5: 96-100. Berridge M. 1993. Cell signalling. A tale of two messengers. Nature 365: 388-389. Berridge CW, Waterhouse BD. 2003. The locus coeruleus‐ noradrenergic system: Modulation of behavioral state and state‐dependent cognitive processes. Brain Res Rev 42: 33-84. Bertocci B, Miggiano V, Da Prada M, Dembic Z, Lahm, HW, et al. 1991. Human catechol‐O‐methyltransferase: Cloning and expression of the membrane‐associated form. Proc Natl Acad Sci USA 88: 1416-1420. Boehm S, Kubista, H. 2002. Fine tuning of sympathetic transmitter release via ionotropic and metabotropic presynaptic receptors. Pharmacol Rev 54: 43-99. Bray N, Buckland P, Williams N, Williams H, Norton N, et al. 2003. A haplotype implicated in schizophrenia susceptibility is associated with reduced COMT expression in human brain. Am J Hum Genet 73: 152-161. Brinkmann B, Sajantila A, Goedde H, Matsumoto H, Nishi K, et al. 1996. Population genetic comparisons among eight populations using allele frequency and sequence data from three microsatellite loci. Eur J Hum Genet 4: 175-182. Brunet J‐F, Pattyn A. 2002. Phox2 genes—from patterning to connectivity. Curr Opin Genet Dev 12: 435-440. Brunner HG, Nelen M, Breakfield XO, Ropers HH, Van Oost B. 1993. Abnormal behavior associated with a point mutation in the structural gene for monoamine oxidase A. Science 262: 578-580. Burgert E, Crocq MA, Bausch E, Macher JP, Morris‐ Rosendahl DJ. 1998. No association between the tyrosine hydroxylase microsatellite marker HUMTH01 and schizophrenia or bipolar I disorder. Psychiatr Genet 8: 45-48. Byerley W, Plaetke R, Hoff M, Jensen S, Holik J, et al. 1992. Tyrosine hydroxylase gene not linked to manic‐depression in seven of eight pedigrees. Hum Hered 42: 259-263. Bylund D. 1992. Subtypes of alpha 1‐ and alpha 2‐adrenergic receptors. FASEB J 6: 832-839. Bylund, DB. 2005. Alpha‐2 adrenoceptor subtypes: Are more better? 144: 159‐160.

6

Bylund D, Eikenberg D, Hieble J, Langer S, Lefkowitz R, et al. 1994. International union of pharmacology nomenclature of adrenoceptors. Pharmacol Rev 46: 121-136. Cases O, Self I, Grimsby J, Gaspar P, Chen K, et al. 1995. Aggressive behavior and altered amounts of brain serotonin and norepinephrine in mice lacking MAOA. Science 268: 1763-1766. Caspi A, McClay J, Moffitt TE, Mill J, Martin J, et al. 2002. Role of genotype in the cycle of violence in maltreated children. Science 297: 851-854. Cavazzoni P, Alda M, Turecki G, Rouleau G, Grof E, et al. 1996. Lithium‐responsive affective disorders: No association with the tyrosine hydroxylase gene. Psychiatry Res 64: 91-96. de Chaldee M, Laurent C, Thibaut F, Martinez M, Samolyk D, et al. 1999. Linkage disequilibrium on the COMT gene in French schizophrenics and controls. Am J Med Genet 88: 452-457. Chen CH, Lee YR, Wei FC, Koong FJ, Hwu HG, et al. 1997. Association study of NlaIII and MspI genetic polymorphisms of catechol‐O‐methyltransferase gene and susceptibility to schizophrenia. Biol Psychiatry 41: 985-987. Chiba M, Suzuki S, Hinokio Y, Hirai M, Satoh Y, et al. 2000. Tyrosine hydroxylase gene microsatellite polymorphism associated with insulin resistance in depressive disorder. Metabolism 49: 1145-1149. Craddock N, Spurlock G, McGuffin P, Owen MJ, Nosten‐ Bertrand M, et al. 1997. No association between bipolar disorder and alleles at a functional polymorphism in the COMT gene. Br J Psychiatry 170: 526-528. Craig SP, Buckle VJ, Lamouroux A, Mallet J, Craig IW. 1988. Localization of the human dopamine beta hydroxylase (DBH) gene to chromosome 9q34. Cytogenet Cell Genet 48: 48-50. Croager EJ, Gout AM, Abraham LJ. 2000. Involvement of Sp1 and microsatellite repressor sequences in the transcriptional control of the human CD30 gene. Am J Pathol 156: 1723-1731. Cubells JF, van Kammen DP, Kelley ME, Anderson GM, O’Connor DT, et al. 1998. Dopamine beta‐hydroxylase: Two polymorphisms in linkage disequilibrium at the structural gene DBH associate with biochemical phenotypic variation. Hum Genet 102: 533-540. Cubells JF, Kobayashi K, Nagatsu T, Kidd KK, Kidd JR, et al. 1997. Population genetics of a functional variant of the dopamine beta‐hydroxylase gene (DBH). Am J Med Genet 74: 374-379. Cubells JF, Kranzler HR, McCance‐Katz E, Anderson GM, Malison RT, et al. 2000. A haplotype at the DBH locus, associated with low plasma dopamine beta‐hydroxylase activity, also associates with cocaine‐induced paranoia. Mol Psychiatry 5: 56-63. Daly G, Hawi Z, Fitzgerald M, Gill M. 1999. Mapping susceptibility loci in attention deficit hyperactivity disorder:

141

142

6

Molecular genetics of brain noradrenergic neurotransmission

Preferential transmission of parental alleles at DAT1, DBH and DRD5 to affected children. Mol Psychiatry 4: 192-196. Daniels JK, Williams NM, Williams J, Jones LA, Cardno AG, et al. 1996. No evidence for allelic association between schizophrenia and a polymorphism determining high or low catechol‐O‐methyltransferase activity. Am J Psychiatry 153: 268-270. De Benedictis G, Carotenuto L, Carrieri G, De Luca M, Falcone E, et al. 1998. Gene/longevity association studies at four autosomal loci (REN, THO, PARP, SOD2). Eur J Hum Genet 6: 534-541. Deckert J, Catalano M, Syagailo Y, Bosi M, Okladnova O, et al. 1999. Excess of high activity monoamine oxidase A gene promoter alleles in female patients with panic disorder. Hum Mol Genet 8: 621-624. Deinum J, Steenbergen‐Spanjers GCH, Jansen M, Boomsma F, Lenders JWM, et al. 2004. DBH gene variants that cause low plasma dopamine {beta} hydroxylase with or without a severe orthostatic syndrome. J Med Genet 41: e38. Detera‐Wadleigh S, Berrettini W, Goldin L, Boorman D, Anderson S, et al. 1987. Close linkage of c‐Harvey‐ras‐1 and the insulin gene to affective disorder is ruled out in three North American pedigrees. Nature 325: 806-808. Egan MF, Goldberg TE, Kolachana BS, Callicott JH, Mazzanti CM, et al. 2001. Effect of COMT val108/158 met genotype on frontal lobe function and risk for schizophrenia. Proc Natl Acad Sci USA 98: 6917-6922. Egeland JA, Gerhard DS, Pauls DL, Sussex JN, Kidd KK, et al. 1987. Bipolar affective disorders linked to DNA markers on chromosome 11. Nature 325: 783-787. Eisenhofer G, Kopin IJ, Goldstein DS. 2004. Catecholamine metabolism: A contemporary view with implications for physiology and medicine. Pharmacol Rev 56: 331-349. Fan J‐B, Yang M‐S, Tang J‐X, He L, Xing YL, et al. 2004. Family‐based association study of the functional monoamine oxidase a gene promoter polymorphism and schizophrenia. Schizophr Res 67: 107-109. Ferguson SSG. 2001. Evolving concepts in G protein‐coupled receptor endocytosis: The role in receptor desensitization and signaling. Pharmacol Rev 53: 1-24. Ferguson J, Wesnes K, Schwartz G. 2003. Reboxetine versus paroxetine versus placebo: Effects on cognitive functioning in depressed patients. Int Clin Psychopharmacol 18: 9-14. Fields TA, Casey PJ. 1997. Signalling functions and biochemical properties of pertussis toxin‐resistant G‐proteins. Biochem J 321: 561-571. Foley DL, Eaves LJ, Wormley B, Silberg JL, Maes HH, et al. 2004. Childhood adversity, monoamine oxidase a genotype, and risk for conduct disorder. Arch Gen Psychiatry 61: 738-744. Fuller R, Hemrick‐Luecke S, Littlefield E, Audia J. 1995. Comparison of desmethylsertraline with sertraline as a

monoamine uptake inhibitor in vivo. Prog Neuropsychopharmacol Biol Psychiatry 19: 135-149. Furlong RA, Rubinsztein JS, Ho L, Walsh C, Coleman TA, et al. 1999. Analysis and metaanalysis of two polymorphisms within the tyrosine hydroxylase gene in bipolar and unipolar affective disorders. Am J Med Genet 88: 88-94. Gallinat J, Bajbouj M, Sander T, Schlattmann P, Xu K, et al. 2003. Association of the G1947A COMT (Val108/158Met) gene polymorphism with prefrontal P300 during information processing. Biol Psychiatry 54: 40-48. Garland EM, Hahn MK, Ketch TP, Keller NR, Kim CH, et al. 2002. Genetic Basis of clinical catecholamine disorders. Ann NY Acad Sci 971: 506-514. Geller B, Cook Jr., EH. 2000. Ultradian rapid cycling in prepubertal and early adolescent bipolarity is not in transmission disequilibrium with val/met COMT alleles. Biol Psychiatry 47: 605-609. Gerhard D, LaBuda M, Bland S, Allen C, Egeland J, et al. 1994. Initial report of a genome search for the affective disorder predisposition gene in the old order amish pedigrees: Chromosomes 1 and 11. Am J Med Genet 54: 398-404. Gershon E, Goldin L, Badner J, Berrettini W. 1996. Detection of linkage to affective disorders in the catalogued amish pedigrees: A reply to Pauls et al. Am J Hum Genet 58: 1381-1385. Gether U. 2000. Uncovering molecular mechanisms involved in activation of G protein‐coupled receptors. Endocr Rev 21: 90-113. Gill M, Castle D, Hunt N, Clements A, Sham P, et al. 1991. Tyrosine hydroxylase polymorphisms and bipolar affective disorder. J Psychiatr Res 25: 179-184. Ginns E, Egeland J, Allen C, Pauls D, Falls K, et al. 1992. Update on the search for DNA markers linked to manic‐ depressive illness in the old order amish. J Psychiatr Res 26: 305-308. Ginns E, Ott J, Egeland J, Allen C, Fann C, et al. 1996. A genome‐wide search for chromosomal loci linked to bipolar affective disorder in the old order amish. Nat Genet 12: 431-435. Goridis C, Rohrer H. 2002. Specification of catecholaminergic and serotonergic neurons. Nat Rev Neurosci 3: 531-541. Grace AA. 1991. Phasic versus tonic dopamine release and the modulation of dopamine system responsivity: A hypothesis for the etiology of schizophrenia. Neuroscience 41: 1-24. Grace AA. 1993. Cortical regulation of subcortical dopamine systems and its possible relevance to schizophrenia. J Neural Transm Gen Sect 91: 111-134. Grossman MH, Emanuel BS, Budarf ML. 1992. Chromosomal mapping of the human catechol‐O‐methyltransferase gene to 22q11.1–q11.2. Genomics 12: 822-825.

Molecular genetics of brain noradrenergic neurotransmission Gurling H, Smyth C, Kalsi G, Moloney E, Rifkin L, et al. 1995. Linkage findings in bipolar disorder. Nat Genet 10: 8-9. Gutierrez B, Bertranpetit J, Guillamat R, Valles V, Arranz MJ, et al. 1997. Association analysis of the catechol‐ O‐methyltransferase gene and bipolar affective disorder. Am J Psychiatry 154: 113-115. HamiltonS, Slager S, Heiman G, Haghighi F, Klein D, et al. 2000. No genetic linkage or association between a functional promoter polymorphism in the monoamine oxidase‐ A gene and panic disorder. Mol Psychiatry 5: 465-466. Hawi Z, Lowe N, Kirley A, Gruenhage F, Nothen M, et al. 2003. Linkage disequilibrium mapping at DAT1, DRD5 and DBH narrows the search for ADHD susceptibility alleles at these loci. Mol Psychiatry 8: 299-308. Hoda F, Nicholl D, Bennett P, Arranz M, Aitchison KJ, et al. 1996. No association between Parkinson’s disease and low‐activity alleles of catechol‐O‐methyltransferase. Biochem Biophys Res Commun 228: 780-784. Hodgkinson S, Sherrington R, Gurling H, Marchbanks R, Reeders S, et al. 1987. Molecular genetic evidence for heterogeneity in manic depression. Nature 325: 805-806. Houhou L, Lamouroux A, Faucon Biguet N, Mallet J. 1995. Expression of human dopamine b‐hydroxylase in mammalian cells infected by recombinant vaccinia virus. J Biol Chem 270: 12601-12606. Huang Y, Cate S, Battistuzzi C, Oquendo M, Brent D, et al. 2004. An association between a functional polymorphism in the monoamine oxidase a gene promoter, impulsive traits and early abuse experiences. Neuropsychopharmacology 29: 1498-1505. Hubbard KB, Hepler JR. 2006. Cell signalling diversity of the Gq[alpha] family of heterotrimeric G proteins. Cellular Signalling 18: 135-150. Ibanez A, Perez de Castro I, Fernandez‐Piqueras J, Blanco C, Saiz‐Ruiz J. 2000. Pathological gambling and DNA polymorphic markers at MAO‐A and MAO‐B genes. Mol Psychiatry 5: 105-109. Icard Liepkalns C, Faucon Biguet N, Vyas S, Robert JJ, Sassone Corsi P, et al. 1992. AP‐1 complex and c‐fos transcription are involved in TPA provoked and trans‐synaptic inductions of the tyrosine hydroxylase gene: Insights into long‐term regulatory mechanisms. J Neurosci Res 32: 290-298. Inayama Y, Yoneda H, Sakai T, Ishida T, Kobayashi S, et al. 1993. Lack of association between bipolar affective disorder and tyrosine hydroxylase DNA marker. Am J Med Genet 48: 87-89. Ishii A, Kobayashi K, Kiuchi K, Nagatsu T. 1991. Expression of two forms of human dopamine‐beta‐hydroxylase in COS cells. Neurosci Lett 125: 25-28. Jindra A, Jachymova M, Horky K, Peleska J, Umnerova V, et al. 2000. Association analysis of two tyrosine hydroxylase

6

gene polymorphisms in normotensive offspring from hypertensive families. Blood Press 9: 250-254. Jo¨nsson EG, Bah J, Melke J, Jamra RA, Schumacher J, et al. 2004. Monoamine related functional gene variants and relationships to monoamine metabolite concentrations in CSF of healthy volunteers. BMC Psychiatry 4: 4. Jonsson EG, Geijer T, Gyllander A, Terenius L, Sedvall GC. 1998. Failure to replicate an association between a rare allele of a tyrosine hydroxylase gene microsatellite and schizophrenia. Eur Arch Psychiatry Clin Neurosci 248: 61-63. Jonsson EG, Norton N, Forslund K, Mattila‐Evenden M, Rylander G, et al. 2003. Association between a promoter variant in the monoamine oxidase A gene and schizophrenia. Schizophr Res 61: 31-37. Jonsson E, Sedvall G, Brene S, Gustavsson JP, Geijer T, et al. 1996. Dopamine‐related genes and their relationships to monoamine metabolites in CSF. Biol Psychiatry 40: 1032-1043. Jorm A, Henderson A, Jacomb P, Christensen H, Korten A, et al. 2000. Association of a functional polymorphism of the monoamine oxidase A gene promoter with personality and psychiatric symptoms. Psychiatr Genet 10: 87-90. Kang J, Shi Y, Xiang B, Qu B, Su W, et al. 2005. A nuclear function of [beta]‐arrestin1 in GPCR signaling: Regulation of histone acetylation and gene transcription. Cell 123: 833-847. Karayiorgou M, Altemus M, Galke BL, Goldman D, Murphy DL, et al. 1997. Genotype determining low catechol‐O‐methyltransferase activity as a risk factor for obsessive‐compulsive disorder. Proc Natl Acad Sci USA 94: 4572-4575. Karayiorgou M, Gogos J. 1997. A turning point in schizophrenia genetics. Neuron 19: 967-979. Karayiorgou M, Gogos JA, Galke BL, Wolyniec PS, Nestadt G, et al. 1998. Identification of sequence variants and analysis of the role of the catechol‐O‐methyl‐transferase gene in schizophrenia susceptibility. Biolo Psychiatry 43: 425-431. Karayiorgou M, Morris MA, Morrow B, Shprintzen RJ, Goldberg R, et al. 1995. Schizophrenia susceptibility associated with interstitial deletions of chromosome 22q11. Proc Natl Acad Sci USA 92: 7612-7616. Kawada Y, Hattori M, Fukuda R, Arai H, Inoue R, et al. 1995. No evidence of linkage or association between tyrosine hydroxylase gene and affective disorder. J Affect Disord 34: 89-94. Kelsoe J, Ginns E, Egeland J, Gerhard D, Goldstein A, et al. 1989. Re‐evaluation of the linkage relationship between chromosome 11p loci and the gene for bipolar affective disorder in the old order amish. Nature 342: 238-243. Kirov G, Norton N, Jones I, McCandless F, Craddock N, et al. 1999. A functional polymorphism in the promoter of

143

144

6

Molecular genetics of brain noradrenergic neurotransmission

monoamine oxidase A gene and bipolar affective disorder. Int J Neuropsychopharmcol 2: 293-298. Klimek V, Stockmeier C, Overholser J, Meltzer HY, Kalka S, et al. 1997. Reduced levels of norepinephrine transporters in the locus coeruleus in major depression. J Neurosci 17: 8451-8458. Kohnke MD, Zabetian CP, Anderson GM, Kolb W, Gaertner I, et al. 2002. A genotype‐controlled analysis of plasma dopamine [beta]‐hydroxylase in healthy and alcoholic subjects: Evidence for alcohol‐related differences in noradrenergic function. Biol Psychiatry 52: 1151-1158. Korner J, Fritze J, Propping P. 1990. RFLP alleles at the tyrosine hydroxylase locus: No association found to affective disorders. Psychiatry Res 32: 275-280. Korner J, Rietschel M, Hunt N, Castle D, Gill M, et al. 1994. Association and haplotype analysis at the tyrosine hydroxylase locus in a combined German–British sample of manic depressive patients and controls. Psychiatr Genet 4: 167-175. Kunugi H, Ishida S, Kato T, Tatsumi M, Sakai T, et al. 1999. A functional polymorphism in the promoter region of monoamine oxidase‐A gene and mood disorders. Mol Psychiatry 4: 393-395. Kunugi H, Nanko S, Ueki A, Otsuka E, Hattori M, et al. 1997a. High and low activity alleles of catechol‐O‐methyltransferase gene: Ethnic difference and possible association with Parkinson’s disease. Neurosci Lett 221: 202-204. Kunugi H, Vallada HP, Hoda F, Kirov G, Gill M, et al. 1997b. No evidence for an association of affective disorders with high‐ or low‐activity allele of catechol‐O‐methyltransferase gene. Biol Psychiatry 42: 282-285. Kurumaji A, Kuroda T, Yamada K, Yoshikawa T, Toru M. 2001. An association of the polymorphic repeat of tetranucleotide (TCAT) in the first intron of the human tyrosine hydroxylase gene with schizophrenia in a Japanese sample. J Neural Transm 108: 489-495. Lachman HM, Kelsoe J, Moreno L, Katz S, Papolos DF. 1997. Lack of association of catechol‐O‐methyltransferase (COMT) functional polymorphism in bipolar affective disorder. Psychiatric Genet 7: 13-17. Lachman HM, Morrow B, Shprintzen R, Veit S, Parsia SS, et al. 1996b. Association of codon 108/158 catechol‐O‐ methyltransferase gene polymorphism with the psychiatric manifestations of velo‐cardio‐facial syndrome. Am J Med Genet 67: 468-472. Lachman HM, Nolan KA, Mohr P, Saito T, Volavka J. 1998. Association between catechol‐O‐methyltransferase genotype and violence in schizophrenia and schizoaffective disorder. Am J Psychiatry 155: 835-837. Lachman HM, Papolos DF, Saito T, Yu Y‐M, Szumlanski CL, et al. 1996a. Human catechol‐O‐methyltransferase pharmacogenetics: Description of a functional polymorphism and

its potential application to neuropsychiatric disorders. Pharmacogenetics 6: 243-250. Lan NC, Heinzmann C, Gal A, Klisak I, Orth U, et al. 1989. Human monoamine oxidase A and B genes map to Xp 11.23 and are deleted in a patient with norrie disease. Genomics 4: 552-559. Leboyer M, Malafosse A, Boularand S, Campion D, Gheysen F, et al. 1990. Tyrosine hydroxylase polymorphisms associated with manic‐depressive illness. Lancet 335: 1219. Leonard B. 2001. Stress, norepinephrine and depression. J Psychiatry Neurosci 26: Suppl: S11-S16. Lester HA, Cao Y, Mager S. 1996. Listening to neurotransmitter transporters. Neuron 17: 807-810. Li T, Vallada H, Curtis D, Arranz M, Xu K, et al. 1997. Catechol‐O‐methyltransferase Val158Met polymorphism: Frequency analysis in Han Chinese subjects and allelic association of the low activity allele with bipolar affective disorder. Pharmacogenetics 7: 349-353. Lim LC, Gurling H, Curtis D, Brynjolfsson J, Petursson H, et al. 1993. Linkage between tyrosine hydroxylase gene and affective disorder cannot be excluded in two of six pedigrees. Am J Med Genet 48: 223-228. Lobos EA, Todd RD. 1997. Cladistic analysis of disease association with tyrosine hydroxylase: Application to manic‐ depressive disease and alcoholism. Am J Med Genet 74: 289-295. Lohmueller KE, Pearce CL, Pike M, Lander ES, Hirschhorn JN. 2003. Meta‐analysis of genetic association studies supports a contribution of common variants to susceptibility to common disease. Nat Genet 33: 177-182. Lotta T, Vidgren J, Tilgmann C, Ulmanen I, Melen K, et al. 1995. Kinetics of human soluble and membrane‐bound catechol‐O‐methyltransferase: A revised mechanism and description of the thermolabile variant of the enzyme. Biochemistry 34: 4202-4210. Lundstrom K, Tenhunen J, Tilgmann C, Karhunen T, Panula P, et al. 1995. Cloning, expression and structure of catechol‐ O‐methyltransferase. Biochim Biophys Acta Protein Structure and Molecular Enzymology 1251: 1-10. Malafosse A, Leboyer M, d’Amato T, Amadeo S, Abbar M, et al. 1997. Manic depressive illness and tyrosine hydroxylase gene: Linkage heterogeneity and association. Neurobiol Dis 4: 337-349. Mallet J. 1996. The TiPS/TINS lecture. Catecholamines: From gene regulation to neuropsychiatric disorders. Trends Pharmacol Sci 17: 129-135. Mandela P, Ordway GA. 2006. The norepinephrine transporter and its regulation. J Neurochem 97: 310-333. Masson J, Sagne C, Hamon M, Mestikawy SE. 1999. Neurotransmitter transporters in the central nervous system. Pharmacol Rev 51: 439-464.

Molecular genetics of brain noradrenergic neurotransmission Mattay VS, Goldberg TE., Fera F, Hariri AR., Tessitore A, et al. 2003. Catechol‐O‐methyltransferase val158‐met genotype and individual variation in the brain response to amphetamine. Proc Natl Acad Sci USA 100: 6186-6191. Melia K, Duman R. 1991. Involvement of corticotropin‐ releasing factor in chronic stress regulation of the brain noradrenergic system. Proc Natl Acad Sci USA 88: 8382-8386. Melia K, Rasmussen K, Terwilliger R, Haycock J, Nestler E, et al. 1992. Coordinate regulation of the cyclic AMP system with firing rate and expression of tyrosine hydroxylase in the rat locus coeruleus: Effects of chronic stress and drug treatments. J Neurochem 58: 494-502. Meloni R, Albanese V, Ravassard P, Treilhou F, Mallet J. 1998. A tetranucleotide polymorphic microsatellite, located in the first intron of the tyrosine hydroxylase gene, acts as a transcription regulatory element in vitro. Hum Mol Genet 7: 423-428. Meloni R, Laurent C, Campion D, Ben Hadjali B, Thibaut F, et al. 1995b. A rare allele of a microsatellite located in the tyrosine hydroxylase gene found in schizophrenic patients. C R Acad Sci III 318: 803-809. Meloni R, Leboyer M, Bellivier F, Barbe B, Samolyk D, et al. 1995a. Association of manic‐depressive illness with tyrosine hydroxylase microsatellite marker. Lancet 345: 932. Meyer E, Wiegand P, Rand SP, Kuhlmann D, Brack M, et al. 1995. Microsatellite polymorphisms reveal phylogenetic relationships in primates. J Mol Evol 41: 10-14. Moore H, West AR, Grace AA. 1999. The regulation of forebrain dopamine transmission: Relevance to the pathophysiology and psychopathology of schizophrenia. Biol Psychiatry 46: 40-55. Morilak DA, Barrera G, Echevarria DJ, Garcia AS, Hernandez A, et al. 2005. Role of brain norepinephrine in the behavioral response to stress. Prog Neuropsychopharmacol Biol Psychiatry—Experimental Stress: From Basic to Clinical Aspects 29: 1214-1224. Mu¨ller Smith K, Daly M, Fischer M, Yiannoutsos CT, Bauer L, et al. 2003. Association of the dopamine beta hydroxylase gene with attention deficit hyperactivity disorder: Genetic analysis of the Milwaukee longitudinal study. Am J Med Genet 119B: 77-85. Mynett‐Johnson LA, Murphy VE, Claffey E, Shields DC, McKeon P. 1998. Preliminary evidence of an association between bipolar disorder in females and the catechol‐O‐ methyltransferase gene. Psychiatr Genet 8: 221-225. Neer EJ. 1995. Heterotrimeric C proteins: Organizers of transmembrane signals. Cell 80: 249-257. Nelson JC. 1999. A review of the efficacy of serotonergic and noradrenergic reuptake inhibitors for treatment of major depression. Biol Psychiatry 46: 1301-1308. Njus D, Kelley P, Harnadek G. 1986. Bioenergetics of secretory vesicles. Biochim Biophys Acta 853: 237-265.

6

Ohmori O, Shinkai T, Kojima H, Terao T, Suzuki T, et al. 1998. Association study of a functional catechol‐O‐methyltransferase gene polymorphism in Japanese schizophrenics. Neurosci Lett 243: 109-112. Ordway G. 1997. Pathophysiology of the locus coeruleus in suicide. Ann N Y Acad Sci 836: 233-252. Pakstis A, Kidd J, Castiglione C, Kidd K. 1991. Status of the search for a major genetic locus for affective disorder in the old order amish. Hum Genet 87: 475-483. Palmatier MA, Kang AM, Kidd KK. 1999. Global variation in the frequencies of functionally different catechol‐O‐ methyltransferase alleles. Biol Psychiatry 46: 557-567. Perez de Castro I, Santos J, Torres P, Visedo G, Saiz‐Ruiz J, et al. 1995. A weak association between TH and DRD2 genes and bipolar affective disorder in a Spanish sample. J Med Genet 32: 131-134. Persson ML, Wasserman D, Jonsson EG, Bergman H, Terenius L, et al. 2000. Search for the influence of the tyrosine hydroxylase (TCAT)(n) repeat polymorphism on personality traits. Psychiatry Res 95: 1-8. Polymeropoulos M, Xiao H, Rath D, Merril C. 1991. Tetranucleotide repeat polymorphism at the human tyrosine hydroxylase gene (TH). Nucleic Acids Res 19: 3753. de Pontual L, Nepote V, Attie‐Bitach T, Al Halabiah H, Trang H, et al. 2003. Noradrenergic neuronal development is impaired by mutation of the proneural HASH‐1 gene in congenital central hypoventilation syndrome (ondine’s curse). Hum Mol Genet 12: 3173-3180. Puers C, Hammond HA, Jin L, Caskey T, Schumm JW. 1993. Identification of repeat sequence heterogeneity at the polymorphic short tandem repeat locus HUMTH01 (AATG)n and reassignment of alleles in population analysis by using a locus‐specific allelic ladder. Am J Hum Genet 53: 953-958. Pulver AE, Karayiorgou M, Lasseter VK, Wolyniec P, Kasch L, et al. 1994b. Follow‐up of a report of a potential linkage for schizophrenia on chromosome 22q12–q13.1: Part 2. Am J Med Genet 54: 44-50. Pulver AE, Karayiorgou M, Wolyniec PS, Lasseter VK, Kasch L, et al. 1994a. Sequential strategy to identify a susceptibility gene for schizophrenia: Report of potential linkage on chromosome 22q12–q13.1: Part 1. Am J Med Genet 54: 36-43. Sabban EL, Kvetnansky R. 2001. Stress‐triggered activation of gene expression in catecholaminergic systems: Dynamics of transcriptional events. Trends Neurosci 24: 91-98. Sabol S, Hu S, Hamer D. 1998. A functional polymorphism in the monoamine oxidase A gene promoter. Hum Genet 103: 273-279. Sander T, Harms H, Rommelspacher H, Hoehe M, Schmidt LG. 1998. Possible allelic association of a tyrosine hydroxylase polymorphism with vulnerability to alcohol‐ withdrawal delirium. Psychiatr Genet 8: 13-17.

145

146

6

Molecular genetics of brain noradrenergic neurotransmission

Sassone‐Corsi P, Lamph WW, Kamps M, Verma IM. 1988. Fos‐associated cellular p39 is related to nuclear transcription factor AP‐1. Cell 54: 553-560. Schatzberg A. 2000. Clinical efficacy of reboxetine in major depression. J Clin Psychiatry 61: 31-38. Schildkraut JJ. 1965. The catecholamine hypothesis of affective disorders: A review of supporting evidence. Am J Psychiatry 122: 509-522. Schultz, W, Dayan P, Montague P. 1997. A neural substrate of prediction and reward. Science 275: 1593-1599. Schulze T, Muller D, Krauss H, Scherk H, Ohlraun S, et al. 2000. Association between a functional polymorphism in the monoamine oxidase A gene promoter and major depressive disorder. Am J Med Genet 96: 801-803. SerrettiA, Macciardi F, Cusin C, Verga M, Pedrini S, 1998a. Tyrosine hydroxylase gene in linkage disequilibrium with mood disorders. Mol Psychiatry 3: 169-174. Serretti A, Macciardi F, Verga M, Cusin C, Pedrini S, et al. 1998b. Tyrosine hydroxylase gene associated with depressive symptomatology in mood disorder. Am J Med Genet 81: 127-130. Servan‐Schreiber D, Printz H, Cohen J. 1990. A network model of catecholamine effects: Gain, signal‐to‐noise ratio, and behavior. Science 249: 892-895. Shifman S, Bronstein M, Sternfeld M, Pisante´ A, Weizman A, et al. 2004. COMT: A common susceptibility gene in bipolar disorder and schizophrenia. Am J Med Genet 128B: 61-64. Shifman S, Bronstein M, Sternfeld M, Pisante‐Shalom A, Lev‐Lehman E, et al. 2003. A highly significant association between a COMT haplotype and schizophrenia. Am J Hum Genet 71: 1296-1302. Shih JC, Chen K, Ridd, J. 1999. Monoamine oxidase: From genes to behavior. Annu Rev Neurosci 22: 197-217. Smyth C, Kalsi G, Brynjolfsson J, O’Neill J, Curtis D, et al. 1996. Further tests for linkage of bipolar affective disorder to the tyrosine hydroxylase gene locus on chromosome 11p15 in a new series of multiplex British affective disorder pedigrees. Am J Psychiatry 153: 271-274. Souery D, Lipp O, Mahieu B, Mendelbaum K, De Martelaer V, et al. 1996. Association study of bipolar disorder with candidate genes involved in catecholamine neurotransmission: DRD2, DRD3, DAT1, and TH genes. Am J Med Genet 67: 551-555. Souery D, Lipp O, Rivelli SK, Massat I, Serretti A, et al. 1999. Tyrosine hydroxylase polymorphism and phenotypic heterogeneity in bipolar affective disorder: A multicenter association study. Am J Med Genet 88: 527-532. Southwick S, Yehuda R, Giller EJ. 1993. Personality disorders in treatment‐seeking combat veterans with posttraumatic stress disorder. Am J Psychiatry 150: 1020-1023. Starke K. 2001. Presynaptic autoreceptors in the third decade: Focus on alpha2‐adrenoceptors. J Neurochem 78: 685-693.

Stolk JM, Hurst JH, Nisula BC. 1982. Regulation and inheritance of dopamine‐beta‐hydroxylase. Behav Genet 12: 37-52. Strous RD, Bark N, Woerner M, Lachman HM. 1997. Lack of association of a functional catechol‐O‐methyltransferase gene polymorphism in schizophrenia. Biol Psychiatry 41: 493-495. Sugama S, Namihira T, Matsuoka R, Taira N, Eto Y, et al. 1999. Psychiatric inpatients and chromosome deletions within 22q11.2. J Neurol Neurosurg Psychiatry 67: 803-806. Sullivan G, Coplan J, Kent J, Gorman J. 1999. The noradrenergic system in pathological anxiety: A focus on panic with relevance to generalized anxiety and phobias. Biol Psychiatry 46: 1205-1218. Syagailo Y, Stober G, Grassle M, Reimer E, Knapp M, et al. 2001. Association analysis of the functional monoamine oxidase a gene promoter polymorphism in psychiatric disorders. Am J Med Genet 105: 168-171. Syvanen A‐C, Tilgmann C, Rinne J, Ulmanen I. 1997. Genetic polymorphism of catechol‐O‐methyltransferase (COMT): Correlation of genotype with individual variation of S‐COMT activity and comparison of the allele frequencies in the normal population and Parkinsonian patients in Finland. Pharmacogenetics 7: 65-71. Taussig R, Gilman AG. 1995. Mammalian membrane‐bound adenylyl cyclases. J Biol Chem 270: 1-4. Tenhunen J, Salminen M, Lundstrom K, Savolainen R, Ulmanen I. 1994. Genomic organization of the human catechol‐O‐methyltransferase gene and its expression from two distinct promoters. Eur J Biochem 223: 1049-1059. Thibaut F, Ribeyre JM, Dourmap N, Meloni R, Laurent C, et al. 1997. Association of DNA polymorphism in the first intron of the tyrosine hydroxylase gene with disturbances of the catecholaminergic system in schizophrenia. Schizophr Res 23: 259-264. Turecki G, Rouleau GA, Mari J, Joober R, Morgan K. 1997. Lack of association between bipolar disorder and tyrosine hydroxylase: A meta‐analysis. Am J Med Genet 74: 348-352. Vandenbergh DJ, Rodriguez LA, Miller IT, Uhl GR, Lachman HM. 1997. High‐activity catechol‐O‐methyltransferase allele is more prevalent in polysubstance abusers. Am J Med Genet—Neuropsychiatric Genetics 74: 439-442. Versiani M, Cassano G, Perugi G, Benedetti A, Mastalli L, et al. 2002. Reboxetine, a selective norepinephrine reuptake inhibitor, is an effective and well‐tolerated treatment for panic disorder. J Clin Psychiatry 63: 31-37. Wei J, Hemmings GP. 1999. Lack of evidence for association between the COMT locus and schizophrenia. Psychiatr Genet 9: 183-186. Wei J, Ramchand CN, Hemmings GP. 1995. Association of polymorphic VNTR region in the first intron of the human

Molecular genetics of brain noradrenergic neurotransmission TH gene with disturbances of the catecholamine pathway in schizophrenia. Psychiatr Genet 5: 83-88. Wei J, Ramchand CN, Hemmings GP. 1997. Possible association of catecholamine turnover with the polymorphic (TCAT)n repeat in the first intron of the human tyrosine hydroxylase gene. Life Sci 61: 1341-1347. Wei J, Ramchand CN, Hemmings G. 1998. TaqI polymorphic sites at the human dopamine beta‐hydroxylase gene possibly associated with biochemical alterations of the catecholamine pathway in schizophrenia. Psychiatr Genet 8: 19-24. Weinberger DR, Egan MF, Bertolino A, Callicott JH, Mattay VS, et al. 2001. Prefrontal neurons and the genetics of schizophrenia. Biol Psychiatry 50: 825-844. Weinshilboum R. 1978. Human biochemical genetics of plasma dopamine‐[beta]‐hydroxylase and erythrocyte catechol‐ O‐methyltransferase. Hum Genet 45: 101-112. Wigg K, Zai G, Schachar R, Tannock R, Roberts W, et al. 2002. Attention deficit hyperactivity disorder and the gene for dopamine beta‐hydroxylase. Am J Psychiatry 159: 1046-1048. Woodward D, Moises H, Waterhouse B, Yeh H, Cheun J. 1991. Modulatory actions of norepinephrine on neural circuits. Adv Exp Med Biol 287: 193-208. Xie T, Ho SL, Li LSW, Ma OCK. 1997. G/A1947 polymorphism in catechol‐O‐methyltransferase (COMT) gene in Parkinson’s disease. Mov Disord 12: 426-427. Yamamoto K, Cubells J, Gelernter J, Benkelfat C, Lalonde P, et al. 2003. Dopamine beta‐hydroxylase (DBH) gene and schizophrenia phenotypic variability: A genetic association study. Am J Med Genet 117B: 33-38.

6

Yamamoto K, Hornykiewicz O. 2004. Proposal for a noradrenaline hypothesis of schizophrenia. Prog Neuropsychopharmacol Biol Psychiatry 28: 913-922. Young LJ, Nilsen R, Waymire KG, Mac Gregor GR, Insel TR. 1999. Increased affiliative response to vasopressin in mice expressing the V1a receptor from a monogamous vole. Nature 400: 766-768. Zabetian CP, Anderson GM, Buxbaum SG, Elston RC, Ichinose H, et al. 2001. A quantitative‐trait analysis of human plasma‐dopamine beta‐hydroxylase activity: Evidence for a major functional polymorphism at the DBH locus. Am J Hum Genet 68: 515-522. Zabetian CP, Buxbaum SG, Elston RC, Ko¨hnke MD, Anderson GM, et al. 2003. The structure of linkage disequilibrium at the DBH locus strongly influences the magnitude of association between diallelic markers and plasma dopamine—hydroxylase activity. Am J Hum Genet 72: 1389-1400. Zhang L, Rao F, Wessel J, Kennedy BP, Rana BK, et al. 2004. Functional allelic heterogeneity and pleiotropy of a repeat polymorphism in tyrosine hydroxylase: Prediction of catecholamines and response to stress in twins. Physiol Genomics 19: 277-291. Zhu MY, Klimek V, Dilley GE, Haycock JW, Stockmeier C, et al. 1999. Elevated levels of tyrosine hydroxylase in the locus coeruleus in major depression. Biol Psychiatry 46: 1275-1286. Zhuma T, Tyrrell R, Sekkali B, Skavdis G, Saveliev A, et al. 1999. Human HMG box transcription factor HBP1: A role in hCD2 LCR function. EMBO J 18: 6396-6406.

147

7

Dopamine and the Dopaminergic Systems of the Brain

L. G. Harsing Jr.

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151

2 2.1 2.2 2.3 2.4

The Synthetic Pathway of Dopamine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152 Tyrosine Hydroxylase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152 Estimation of Dopamine Turnover Rate from Dopamine Synthesis Inhibition . . . . . . . . . . . . . . . . . . 154 L‐Aromatic Amino Acid Carboxylase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 154 Estimation of Dopamine Synthesis Rate from Dopa Decarboxylase Inhibition . . . . . . . . . . . . . . . . . . 154

3 3.1 3.2 3.3

The Degradative Pathway of Dopamine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 Monoamine Oxidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 Catechol‐O‐Methyl‐Transferase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 Estimation of Dopamine Turnover Rate by Calculation of Metabolites/Dopamine Ratio . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155

4 Storage of Dopamine in Neuronal Pools . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 156 4.1 Vesicular Monoamine Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158 5 5.1 5.2 5.3

Dopamine Plasma Membrane Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158 Structure of Dopamine Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158 Operation of Dopamine Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159 Drugs Acting on Dopamine Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159

6 6.1 6.2 6.3 6.4 6.5

Dopamine Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159 Classification of Dopamine Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 160 Structure of Dopamine Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 160 Postsynaptic Dopamine Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 Presynaptic Dopamine Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 Regional Distribution of Dopamine Autoreceptors Assessed by the g‐Butyrolactone Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 6.6 Changes in Dopamine Receptor Sensitivity and Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 7 7.1 7.2 7.3 7.4

Dopamine Release from Neuronal Stores . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163 Action Potential Propagation‐Induced Dopamine Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163 Dopamine Release Evoked by Reverse‐Mode Operation of Dopamine Transporters . . . . . . . . . . . . . 164 Dopamine Release Evoked by Ion Channel‐Coupled Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165 Regulation of Dopamine Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165

8 Dopaminergic Innervations in the Central Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165 8.1 Dopamine in the Striatum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166

#

2008 Springer ScienceþBusiness Media, LLC.

150

7

Dopamine and the dopaminergic systems of the brain

8.2 Dopamine in the Cerebral Cortex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166 9 Altered Dopaminergic Neurotransmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166 9.1 Neurotoxins Used for Destruction of Dopaminergic Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166 9.2 Dopaminergic Neurotransmission in Knockout Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167 10

Conclusions and Future Avenues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167

Dopamine and the dopaminergic systems of the brain

7

Abstract: The great majority of dopaminergic neurons in the brain (in human 300,000–400,000 cells) is organized in three nuclei, the substantia nigra pars compacta, the ventral tegmental area and the arcuate nucleus. Long pathways originating from these areas project to the neostriatum, the cerebral cortex and limbic structures, and the hypothalamus. Dopaminergic neurotransmission is believed to have a central role in the integration of the reward system; it organizes complete motor programs and voluntary movements, and regulates some endocrine hormone secretion. In pathological conditions, this neurotransmitter system mediates extrapyramidal side effects, drug abuse and dependence, and it is responsible to develop psychiatric disorders such as schizophrenia and attention‐deficit hyperactivity disorder. There are two enzymes, tyrosine hydroxylase and dopa decarboxylase, involved in the synthesis and two enzymes, monoamine oxidase and catechol‐O‐ methyl transferase, take part in the degradation of dopamine. Dopamine in the axon terminals is stored in the cytoplasm and in at least two different vesicular pools. Vesicular monoamine transporter concentrates dopamine into vesicles; this storage protects dopamine from enzymatic degradation, retards dopamine from diffusion out from nerve endings, and serves as pool from where dopamine can be released in response to physiological stimuli. Dopamine release from axon terminals may occur with different mechanisms: action potential propagation‐induced membrane depolarization evokes release from the vesicular stores, whereas the amphetamine class of drugs may induce dopamine efflux by forcing dopamine transporter to operate in reverse mode. The action of dopamine released is primarily terminated by its reuptake into the presynaptic terminals. Dopamine transporter is a transmembrane protein that removes dopamine from the synaptic cleft before it can escape into the biophase. Dopamine transporter knockout mice exhibit a wide range of deficit in dopaminergic functions and altered dopamine‐mediated behavior. There is growing evidence for the importance of extracellular dopamine in the regulation of nonsynaptic neurotransmission, drug actions, and mediation of some dopamine‐related psychiatric disorders. Dopamine, if once released into the synaptic cleft, acts on dopamine receptors. Five dopamine receptors, the D1‐like D1 and D5 and the D2‐like D2, D3, and D4 receptors were identified. Dopamine receptors are coupled to G proteins and they positively or negatively regulate intracellular messenger cascade in which the phosphoprotein DARP‐32 possesses central role. Based upon their expression, presynaptic and postsynaptic dopamine receptors can be distinguished. Response to postsynaptic D1 and D2 receptors alters cellular messenger cascades and that to presynaptic D2 receptors stimulation influences dopamine release and synthesis and neuronal firing rate. Newly synthesized compounds with affinity to dopamine D2/D3 receptors or inhibitors of dopamine degradation are now clinically tested for treatment of schizophrenia and Parkinson’s disease. List of Abbreviations: AADC, amino acid carboxylase; AC, adenylate cyclase; cAMP, cyclic adenosine 50 ‐monophosphate; COMT, catechol‐O‐methyl‐transferase; DA, dopamine; DARPP‐32, dopamine‐ and cAMP‐regulated phosphoprotein‐32; DAT, dopamine plasma membrane transporter; DOPAC, 3,4‐dihydroxyphenylacetic acid; DOPA, dihydroxyphenylalanine; GABA, g‐amino‐butyric acid; GBL, g‐butyrolactone; G‐protein, guanine nucleotide binding protein; HVA, homovanillic acid; MAO, monoamine oxidase; MPPþ, 1‐methyl‐4‐phenylpyridinium; MPTP, 1‐methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine; NO, nitric oxide; NOS, nitric oxide synthase; pDARPP‐32, phosphorylated dopamine‐ and cAMP‐regulated phosphoprotein‐32; PKA, protein kinase A; PP‐1, protein phosphatase‐1; PP‐2B, protein phosphatase‐2B; SAM, S‐adenosyl‐methionine; SLC, solute carrier; TH, tyrosine hydroxylase; 3‐MT, 3‐methoxytyramine; TR, turnover rate; VMATs, vesicular monoamine transporter proteins

1

Introduction

Dopamine (3,4‐dihydroxyphenylalanine) is an endogenous compound containing a benzene ring with two hydroxyl substituents (catechol nucleus) and an aminoethyl group attached to the substituted ring. Dopamine and other compounds with similar molecular structures (norepinephrine and epinephrine) belong to catecholamines and they are often referred in neurosciences as biogenic amines or monoamine neurotransmitters. Dopamine is an important neurotransmitter in the central nervous system and mediates a number of physiological regulations and also involved in the development of neurological and psychiatric disorders. Thus, dopamine has a major role in the pathology of control of movement (Parkinson’s disease)

151

152

7

Dopamine and the dopaminergic systems of the brain

or in psychiatric disorders such as schizophrenia and attention deficit hyperactivity disorder. In addition, dopaminergic neurotransmission has a central role in the mechanism of drug abuse associated with dependence and also in the integration of the reward system of the brain. Long dopaminergic projections in the central nervous system that relate to these functions were visualized first by a fluorescent method described by Dahlstrom and Fuxe (1964). Dopamine and other biogenic amines (norepinephrine, epinephrine, serotonin, and histamine) are members of the classic neurotransmitters. There are several differences between the morphological appearance and neurochemistry of the classic and nonclassic (peptide) neurotransmitters. Storage vesicles, when they are present, are smaller for classic neurotransmitters and they are larger for nonclassic neurotransmitters. Another different feature is that classic neurotransmitters or, in some cases, their metabolic products are subject to reuptake but there is no energy dependent, high‐affinity transport system for peptide neurotransmitters. Moreover, classic neurotransmitters are synthesized in the presynaptic axon terminals, the synthesis of nonclassic transmitters occurs in the cell body and the precursor proteins reach nerve endings by axonal transport.

2

The Synthetic Pathway of Dopamine

There are two enzymatic steps involved in dopamine synthesis (Von Bohlen und Halbach and Dermietzel, 2002). The biosynthetic pathway of dopamine begins with the amino acid precursor tyrosine (> Table 7-1). Phenylalanine is converted to tyrosine by the enzyme phenylalanine hydroxylase although dopamine biosynthesis is usually considered to begin with tyrosine (> Figure 7-1). Tyrosine is hydroxylated at

. Table 7-1 Enzymes involved in the biosynthesis and degradation of dopamine Enzymes Tyrosine hydroxylase L‐aromatic amino acid carboxylase Dopamine‐b‐hydroxylase

Substrate Tyrosine Dopa

Product Dihydroxy‐phenylalanine Dopamine

Dopamine

Norepinephrine

Monoamine oxidase A

Norepinephrine

Monoamine oxidase B

Dopaminea

Catechol‐O‐methyl‐ transferase

Dopamine Norepinephrine

3,4‐Dihydroxy‐ phenylglycolaldehyde 3,4‐Dihydroxyphenyl‐ acetaldehyde 3‐Methoxytyramine Normetanephrine

Inhibitors a‐Methyl p‐tyrosine Carbidopa Benserazide Copper chalators FLA‐63 Clorgyline L‐deprenyl Tolcapone

a

Dopamine is a substrate for type A monoamine oxidase in rat striatum (Demarest et al., 1980), but it is monoamine oxidase B substrate in human (Glover et al., 1977)

position 3 by the enzyme tyrosine hydroxylase (TH) and 3,4‐dihydroxy‐L‐phenylalanine (L‐dopa) is formed. Dopamine synthesis is followed by the next step, when L‐aromatic amino acid decarboxylase converts dopa to dopamine (Deutch and Roth, 2004).

2.1 Tyrosine Hydroxylase Tyrosine hydroxylase catalyzes the addition of hydroxyl group to the meta position of tyrosine, thus forming L‐dopa. Physiological tyrosine concentrations saturate tyrosine hydroxylase and increase of tyrosine concentrations usually does not elevate the rate of dopamine synthesis. Tyrosine hydroxylase is the

Dopamine and the dopaminergic systems of the brain

7

. Figure 7-1 The life cycle of dopamine

rate‐limiting step in dopamine synthesis and the enzyme controls the neuronal concentrations of dopamine. Tyrosine hydroxylase is a mixed‐function oxidase that has moderate substrate specificity; it hydroxylates not only tyrosine but phenylalanine also, particularly in conditions when phenylalanine hydroxylase in suppressed (phenylketonuria). The actions of tyrosine hydroxylase require tetrahydrobiopterin cofactor as hydrogen donor, Fe2þ ion, and molecular oxygen (Thony et al., 2000). Activation of dopaminergic neurons leads to increase of tyrosine hydroxylase activity and dopaminergic neurotransmission is elevated. The short‐term regulation of tyrosine hydroxylase occurs through the posttranslational levels whereas long‐term regulation of tyrosine hydroxylase activity can occur through transcriptional regulation of the gene. Rapid and short term activation of tyrosine hydroxylase occur through phosphorylation and dephosphorylation of at least four serine residues (Ser‐8, Ser‐19, Ser‐31, Ser‐40) in the N terminal part of the enzyme by a series of distinct protein kinases including protein kinase A, protein kinase C, and Ca2þ/calmodulin‐dependent protein kinase II. The conformational changes of the enzyme during phosphorylation result in higher affinity to tetrahydrobiopterin cofactor and lower affinity to dopamine and thus, the endproduct inhibition of the enzyme is decreasing. Tyrosine hydroxylase expression can be either upregulated or downregulated by different drugs such as nicotine, caffeine, morphine, or antidepressants by activating or repressing transcriptional regulatory elements of tyrosine hydroxylase gene promoter. These regulatory elements may include cAMP response element (CRE), glycocorticoid response element (GRE), activator proteins‐1 (AP‐1) or NF‐kB sites (Nestler et al., 2001).

153

154

7

Dopamine and the dopaminergic systems of the brain

2.2 Estimation of Dopamine Turnover Rate from Dopamine Synthesis Inhibition Analogs of tyrosine (a‐methyl‐p‐tyrosine, 3‐iodotyrosine) are competitive inhibitor of tyrosine hydroxylase (Walter et al., 1984; Snyder et al., 1990). a‐Methyl‐p‐tyrosine is widely used to study catecholamine neurotransmission functions, including dopamine turnover rate (Walter et al., 1984; Vizi et al., 1986). Dopamine biosynthesis can be assessed by estimating in vivo activity of tyrosine hydroxylase after treatment with a‐methyl‐p‐tyrosine. Apparent rate of dopamine turnover in brain regions can be determined by following the rate of decline of dopamine concentrations at certain time intervals after i.p. injection of a‐methyl‐p‐tyrosine. The levels of dopamine (as well as its precursors and metabolites) can be determined by HPLC‐electrochemistry. Drugs that are tested to alter dopamine turnover, are usually given before a‐methyl‐p‐tyrosine administration. After blockade of synthesis by a‐methyl‐p‐tyrosine, dopamine concentrations decline at a rate that is proportional to concentration, i.e., log½DA
t ¼ log½DA
0  0:434 kt where [DA]0 is the initial level, [DA]t is the level at time t, and k is the rate constant of dopamine efflux. The turnover rate is product of steady‐state level and of k (the rate constant of dopamine decline): TRDA ¼ k[DA]0. The turnover time of dopamine (the time required to replace the amine pool) is then calculated as Tt ¼1/k. The validity of this method for determining turnover rates of dopamine depends on a‐methyl‐p‐ tyrosine being maintained at an inhibitory level. This was evidenced in that the complete blockade of the enzyme resulted in a decline of dopamine level to an almost zero value at an exponential rate and by the fact that higher doses of a‐methyl‐p‐tyrosine does not increase further the decline.

2.3 L‐Aromatic Amino Acid Carboxylase The hydroxylation of tyrosine by tyrosine hydroxylase generates L‐dopa, which is then decarboxylated by the enzyme L‐aromatic amino acid carboxylase (AADC), also referred as dopa decarboxylase. This enzyme has low Km and high Vmax values and levels of L‐dopa are virtually unmeasurable in the brain under basal conditions. This is because the activity of dopa decarboxylase is so high that L‐dopa is converted into dopamine almost instantaneously. L‐aromatic AADC has low substrate specificity and decarboxylases tyrosine and tryptophane as well as other aromatic amino acids. L‐aromatic AADC is a cytoplasmic enzyme that is present in catecholamine‐ and serotonin‐containing neurons. The enzyme requires pyridoxal 5‐phosphate as a cofactor for its activity. L‐aromatic AADC is exploited in the treatment of Parkinson’s disease by giving L‐dopa to patients to enhance dopamine production in the remaining dopaminergic axon terminals. As dopamine does not cross the blood–brain barrier and L‐dopa readily enters the brain, the latter is used for substitution therapy. A series of L‐dopa analogs such as a‐methyldopa and carbidopa inhibit dopa decarboxylase. Benserazide and RO4‐4602 are also known inhibitors of the enzyme. NSD‐1015 (3‐hydroxybenzylhydrazine) is a centrally active decarboxylase inhibitor and it is widely used to study drugs modifying dopamine synthesis.

2.4 Estimation of Dopamine Synthesis Rate from Dopa Decarboxylase Inhibition The synthesis rate of dopamine can be determined from measuring L‐dopa accumulation in brain tissue samples after inhibition of L‐aromatic amino acid decarboxylase with NSD‐1015. The accumulation of L‐dopa in the rat brain after administration of dopa decarboxylase inhibitors can be used as a measure to estimate tyrosine hydroxylase activity in vivo. L‐dopa accumulation is linear for a certain period after the

Dopamine and the dopaminergic systems of the brain

7

administration of NSD‐1015. The increase in L‐dopa accumulation can be reduced by the dopamine D2 receptor agonist apomorphine and reversed by dopamine antagonists. NSD‐1015, when blocks L‐aromatic amino acid decarboxylase, also inhibits the conversion of the amino acid b‐phenylalanine into b‐phenylethylamine. Administration of NSD‐1015 also leads to elevation of 5‐hydroxytryptophan levels; a technique allows the measurement of serotonin synthesis rate in brain tissue samples.

3

The Degradative Pathway of Dopamine

3.1 Monoamine Oxidase Two major enzymes take part in dopamine catabolism: monoamine oxidase (MAO) and catechol‐O‐ methyl‐transferase (COMT) (Von Bohlen und Halbach and Dermietzel, 2002). MAO occurs in the tissues in the form of two isoenzymes, MAOA and MAOB. MAOA displays high affinity to norepinephrine and serotonin, MAOB exhibits the highest affinity to phenylethylamine (> Table 7-1). MAOA and MAOB have similar affinity to dopamine. MAOs oxidatively deaminate dopamine and its O‐methylated derivative, 3‐methoxytyramine (3‐MT), to form inactive and unstable aldehyde derivatives, 3,4‐dihydroxyphenylacetaldehyde and 3‐methoxy‐4‐hydroxyphenylacetaldehyde (> Figure 7-1). These aldehydes can then be further catabolized by aldehyde dehydrogenases to form corresponding acid metabolites, 3,4‐dihydroxyphenylacetic acid (DOPAC) and homovanillic acid (HVA). MAO requires flavin adenine dinucleotide as cofactor for its activity. The two different forms of MAO are derived from distinct genes and differ not only in their substrate specificity but also in their cellular locations and regulation by pharmacological agents. MAOA is selectively inhibited by clorgyline, and L‐deprenyl selectively inhibits MAOB. Both A and B forms of MAO are associated with the outer membrane of mitochondria. MAOB is also present in glial cells (Nestler et al., 2001).

3.2 Catechol‐O‐Methyl‐Transferase COMT acts to methylate catecholamines and requires S‐adenosyl‐methionine (SAM) as methyl donor for its activity. COMT is relatively nonspecific enzyme that transfers methyl groups from SAM to the meta‐ hydroxy group of catechols. This enzyme was identified in the synaptic cleft. COMT inhibitors (tolcapone, entacapone) increase dopamine levels within the synapses and prolong dopamine receptor activation.

3.3 Estimation of Dopamine Turnover Rate by Calculation of Metabolites/Dopamine Ratio The ratio of DOPAC to dopamine indicates the rate of dopamine metabolism, whereas changes in the levels of dopamine metabolites, DOPAC and HVA, reflect changes in MAO activity. Dopaminergic neuronal activity can be further estimated by calculation of DOPACþHVA/dopamine ratio. This ratio, which indicates alterations in the rate of dopamine turnover, was found to be changed in a number of pathological conditions (experimental parkinsonism or following stroke) as well as during drug treatments (Ogawa et al., 2000; Megyeri et al., 2007). Changes in the levels of 3‐methoxytyramine, a minor metabolite of dopamine, also reflect the turnover and utilization of dopamine. a‐Methyl‐p‐tyrosine produces a parallel decrease in dopamine and 3‐methoxytyramine levels in the striatum and nucleus accumbens. An enhanced 3‐methoxytyramine accumulation can be observed in rats pretreated with MAO inhibitors such as tranylcypromine or pargyline. In addition, accumulation of 3‐methoxytyramine in rat brain provides a sensitive assay to distinguish between dopamine‐releasing agents and uptake inhibitors (Heal et al., 1990).

155

156

7 4

Dopamine and the dopaminergic systems of the brain

Storage of Dopamine in Neuronal Pools

Dopamine is synthesized in the nerve terminal cytoplasm to where enzymes involved in this process are transported from the cell body area (> Figure 7-2). Dopamine is then packed in storage vehicles by means of vesicular monoamine transporter (VMAT) proteins. Dopamine storage vesicles are present at high density within nerve terminals. Vesicles protect dopamine from inactivation by intraneuronal enzymes,

. Figure 7-2 The dopaminergic synapse: schematic diagram of the presynaptic nerve terminal, the synaptic cleft and the postsynaptic cell. At the presynaptic nerve terminals, dopamine is released by exocytosis or by reverse‐mode operation of the dopamine transporter. The released dopamine activates presynaptic D2 autoreceptors inhibiting further release of the neurotransmitter. This inhibition may be the consequence of inhibition of voltage‐ sensitive Ca2þ‐channels and/or inhibition of Kþpermeability. In addition, activation of D2 receptor‐coupled Gi/o protein leads to decreased cAMP production and inhibition of protein kinase A, which regulates proteins involved in the release process. For reversal of dopamine transporter, dopamine releaser drugs enter the nerve endings by the carrier causing intraterminal transfer of dopamine. The elevated external Naþ concentrations force the transporter into reverse‐mode operation and dopamine is extruded out from the nerve terminals. Dopamine released into the synaptic cleft will activate postsynaptic D1‐like (D1 and D5) and D2‐like (D2, D3, and D4) receptors. Activation of the corresponding Gs proteins by D1‐like receptors or Gi/o proteins by D2‐like receptors results in either stimulation or inhibition of adenylate cyclase and increase or decrease of cAMP production. D1 receptor stimulation acts to phosphorylate the phosphoprotein DARPP‐32 via cAMP and protein kinase A, pDARPP‐32 will then inhibit protein phosphatase‐1 (PP‐1) increasing ion channel and receptor phosphorylation. In cells expressing D2‐like receptors, Ca2þ entry activates calcineurin, which in turn, leads to dephosphorylation of pDARPP‐32

whereas dopamine in cytoplasmic store is less protected. Vesicles also retard dopamine from diffusion out of the neuron. Moreover, storage vesicles are ready for fusion with the cellular membrane and they undergo subsequent exocytosis (Hammond, 1996). Vesicular storage of dopamine serves as a depot from where dopamine can be released by appropriate physiological stimuli. The granules that store dopamine also contain ATP but in case of vesicular dopamine release, no enzyme is coreleased.

Dopamine and the dopaminergic systems of the brain

7

Dopamine storage in and efflux from neuronal tissue pools can be characterized by the de Langer–Mulder compartmental analysis (de Langen and Mulder, 1979) by using [3H]dopamine to label (> Figure 7-3). Dopamine pools vary in their turnover rate and turnover time and also in the rate constant of release occurs . Figure 7-3 Compartmental analysis of dopamine pools in rat striatum where [3H]dopamine is taken up to and released from. The release of [3H]dopamine (kBg/g/fraction) was plotted in function of tissue [3H]dopamine content (kBq/g). Desaturation curves of [3H]dopamine efflux with different slopes yield more than on compartment in the release process. Distributional analysis revealed that the efflux compartment consists of various pools with different rate constant, turnover rate and turnover time; these stores participate in the fast, slow and resting release. Striatal slices were prepared from rat brain, loaded with [3H]dopamine and superfused. The slices were stimulated (S) electrically (20 V, 2 Hz, 2‐msec for 2 min) in fraction 4 (S). The radioactivity released from the tissue and that remained in the tissue was determined by liquid scintillation spectrometry

from them. In brain slices, accumulated [3H]dopamine distributes mainly in two compartments: an efflux compartment where release of [3H]dopamine originates from and a bound fraction, which contributes to the efflux with a limited rate (slowly exchanging compartment). Further analysis of radioactivity stored in brain tissue after incubation indicates that the efflux compartment consists of at least three different pools of [3H]dopamine from which fast and slow efflux occur in response to stimulation and a third one, which represents the source of resting dopamine outflow (Harsing, 2006) (> Figure 7-3). Other kinetic analysis has indicated that dopamine is compartmentalized into three separate pools within the presynaptic nerve terminal (Justice et al., 1988). These intraneuronal compartments are the cytosolic dopamine and two vesicular compartments. Dopamine is present in low concentrations in the cytosol where it is subject to metabolism by MAOs; their inhibitors enhance dopamine levels in this compartment. Cytosolic dopamine pool has a pivotal role in distributing newly synthesized dopamine into vesicular storage. From this store, dopamine can be released by reverse‐mode operation of dopamine transporter. There are two sources of cytosolic dopamine: one is uptake from the synaptic cleft and the other is diffusion of dopamine from the vesicular stores. Cytosolic dopamine is lost in three processes: uptake into vesicles, metabolism by MAOs to DOPAC, and efflux into the extracellular fluid. One vesicular compartment is designated as releasable bound dopamine from where action potential‐ induced depolarization evokes transmitter release. The other is a larger inactive compartment, which communicates with the active releasable compartment. The releasable compartment represents the vesicles located near the presynaptic membrane. The rapidly turning over pool contains 5–20% of the total dopamine content in the striatum. The curve of disappearance of dopamine following a‐methyl‐p‐tyrosine administration also reveals the existence of two distinct and separate phases of dopamine decline: an initial

157

158

7

Dopamine and the dopaminergic systems of the brain

and rapid one and a much slower, long‐lasting one starting after drug administration (McMillen et al., 1980). These multiexponential curves of dopamine disappearance observed after a‐methyl‐p‐tyrosine indicate that dopamine is not homogeneously stored in the dopaminergic axon terminals. The biphasic decline in dopamine tissue concentrations is separated by a brief interval when dopamine levels rose to refill the rapidly turning over pool. Dopamine store with fast turnover rate preferentially releases dopamine by depolarization and the other with slow turnover rate considered as an inactive form of dopamine stored. Additional dopamine stores are those present in synaptic cleft and in extrasynaptic space, the latter can be sampled by microdialysis. The main storage site for dopamine in the dendrites may be the smooth endoplasmatic reticulum where dendritic dopamine release may occur from (Bergquist and Nissbrandt, 2005). Dopamine is also present in glial cells.

4.1 Vesicular Monoamine Transporter Accumulation of dopamine in the vesicles depends on the operation of the VMAT (Weihe and Eiden, 2000). VMAT belongs to the intracellular transporters, solute carrier (SLC)18‐gene family (Gether et al., 2006). Two VMATs were identified, one is found in the adrenal medulla and the other is present in the central nervous system, in catecholamine and serotonin neurons. The vesicular uptake process has a low substrate specificity and a variety of biogenic amines including tryptamine, tyramine, and amphetamines can be transported. These amines may compete with endogenous catecholamines for vesicular storage sites. The intravesicularly stored dopamine exists in a complex with ATP and the acidic proteins, chromogranins. The mechanism that concentrates dopamine within the vesicles is an ATP‐dependent process and it is linked to a proton pump. The driving force for uptake into synaptic vesicles is a proton electrochemical gradient generated by a vacuolar Hþ‐ATPase in the synaptic vesicle membrane. The transporter proteins have 12 transmembrane domains and are homologous to a family of bacterial drug resistance transporters. VMAT‐2 has a high affinity to reserpine, which irreversibly blocks vesicular uptake in vivo. Reserpine and related compounds (tetrabenazine, benzquinamide) also inhibit dopamine uptake into the storage vesicles and deplete available stores of dopamine. Reserpine induces depression in human due to depletion of neuronally stored catecholamines and serotonin.

5

Dopamine Plasma Membrane Transporter

Dopamine plasma membrane transporter (DAT) is a transmembrane protein, which effectively removes dopamine from synaptic cleft and returns it into the presynaptic terminals. Dopamine transporter belongs to the Naþ–Cl‐coupled transporters, SLC6‐gene family (Gether et al., 2006). The reuptake of dopamine limits its duration of action on pre‐ and postsynaptic receptors and also its diffusion to other synapses within the biophase. Moreover, the uptake process also allows the recycling and reuse of nonmetabolized dopamine molecules in the neurotransmission process. Reuptake of released dopamine by neurons is the major mode of inactivation.

5.1 Structure of Dopamine Transporter Human dopamine transporter consists of 620 amino acids. The genes for transporters responsible for uptake of dopamine have been cloned revealing protein that belongs to a larger family of neurotransporters. The protein is thought to have 12 transmembrane domains with intracellularly oriented N and C termini and a large glycosylated extracellular loop between transmembrane domains 3 and 4. Dopamine transporter possesses two to four extracellular N‐linked glycosylation sites. Domains 1, 2, and 4–8 may be involved in moving the transmitter across the membrane. The transporter is substrate for protein kinase C‐dependent phosphorylation, which reduces its activity. The dopamine transporter is phosphorylated on the N terminal tail but there are other phosphorylation sites for protein kinase A, protein kinase C, and Ca2þ/calmodulin protein kinase as well (Vaughan, 2004).

Dopamine and the dopaminergic systems of the brain

7

Dopamine transporter gene expression occurs in brain areas in which dopamine is synthesized: the substantia nigra and the ventral tegmental area, it is less prevalent in the arcuate nucleus, olfactory bulb, and the retina. It is most commonly translated in cell bodies and transported to dendrites and axon fibers. Regional distribution of the carrier follows the expected localization of distinct dopamine neurons; however, dopamine transporter is not expressed in all dopamine neurons. The tuberoinfundibular dopamine cells in the hypothalamus that release dopamine into the pituitary portal blood stream, lack demonstrable dopamine transporter mRNA and protein. Dopamine transporter expression is also low in primate prefrontal cortex and a substantial amount of dopamine released is taken up by noradrenergic terminals (Gresch et al., 1995). There is dopamine reuptake into glial cells, and the functional significance of the glial reuptake remains, however, unknown.

5.2 Operation of Dopamine Transporter Dopamine transporter is saturable and its operation can be characterized by the Km and Vmax values determined from the Michaelis–Menten kinetics. The uptake process is energy‐dependent since it can be inhibited by incubation at a low temperature or by metabolic inhibitors. The neuronal reuptake is saturable and depends on Naþ cotransport as well as requiring extracellular Cl (Norregaard and Gether, 2001). Because reuptake depends on cocoupling to the Naþ gradient across the neuronal membrane, drugs such as ouabain, which inhibit Naþ–Kþ‐ATPase, inhibit the reuptake process. Veratridine, which opens Naþchannels, also inhibits the operation of the carrier. The linkage of uptake to the Naþ gradient may have physiological importance since transport is temporarily suspended at the time of depolarization‐induced release of dopamine. Coupling of transporter function to Naþ flow may lead to local changes in the Naþ gradient across the plasma membrane and thereby it can paradoxically extrude dopamine from the nerve endings. Thus, dopamine transporter may act in reverse‐mode operation, a process that conveys dopamine out of the neurons (Gainetdinov et al., 2002). The membrane transporter is not Mg2þ‐dependent, this characteristics distinguishes the neuronal membrane transporters from the vesicular transporters.

5.3 Drugs Acting on Dopamine Transporter Dopamine transporter has limited substrate specificity. Amphetamine and related drugs (methamphetamine, phenmetrazine) are taken up and force transporters actively pump dopamine out from the terminals (Norregaard and Gether, 2001). The amphetamine‐related compounds act as substrate for this transporter and thus, they compete with dopamine reuptake, and are direct releaser also. Other drugs (methylphenidate, nomifensine, amfonelic acid) block dopamine uptake but possess no dopamine releasing effects in vitro in brain slices or synaptosomal preparations. Cocaine binds to the carrier and blocks reuptake of synaptically released dopamine. The cocaine‐binding site in dopamine transporter is distinct from the substrate recognition site. Cocaine and amphetamine exert their effects on arousal by increasing extracellular dopamine concentrations. Some antidepressant and psychostimulant agents block dopamine transporter. Chronic administration of inhibitors alters the number of transporter sites. The membrane transporter is insensitive to reserpine.

6

Dopamine Receptors

Synaptically released dopamine that is not degraded enzymatically, or transported back into the presynaptic cell may activate dopamine receptors. Numerous dopamine receptors have been identified (> Table 7-2), whereas only one transporter has been cloned to dopamine. Dopamine receptors may be located on dendrites and cell bodies of neurons but also occur on axons or nerve terminals. Activation of dopamine receptors may cause decrease of dopamine release or lead to decrease or increase of various other neurotransmitters.

159

160

7

Dopamine and the dopaminergic systems of the brain

. Table 7-2 Types of dopamine receptors and their agonist and antagonist ligands Receptor assays Dopamine D1 Dopamine D2

Agonist SKF38393 SKF81297 SKF82958 Bromocriptine Lisuride

Dopamine D3

BP897 Quinpirole

Dopamine D4

PD168077

Dopamine D5

SKF38393

Antagonist SCH23390

G protein Gs

Haloperidol Domperidone SKF83566 Nafadotrine U99194A Raclopide NGD941 L745870 Clozapine SCH23390

Gi/Go

Gi/Go Gi/Go Gs

Transduction coupling Stimulation of cAMP production Inhibition of cAMP production Inhibition of cAMP production Inhibition of cAMP production Stimulation of cAMP production

6.1 Classification of Dopamine Receptors The first classification of dopamine receptors into two types was proposed based upon a combination of biochemical and pharmacological criteria (Kebabian and Calne, 1979). Two types of dopamine receptors were identified based on differences in their drug specificities and signaling mechanisms. Recent evidence indicated that members of the dopamine receptor family can be generally classified as either D1‐like or D2‐ like receptors. The dopamine D1 receptor was mainly defined as the receptor associated with adenylate cyclase activation in striatal and retinal membranes and displaying low affinity for some antipsychotic drugs, such as sulpiride. The dopamine D2 receptor was defined as being associated with inhibition of prolactin release and displaying high affinity for all antipsychotic agents using radioligand binding experiments. Molecular cloning identified multiple D1 and D2‐like receptors (The IUPHAR Compendium of Receptor Characterization and Classification. 2000). According to our current view, the effects of dopamine are mediated through interaction with five different receptors usually referred to as D1 and D5 receptors and D2, D3, and D4 receptors. D1‐like receptors comprise the D1 and D5 receptors, both exhibit similar pharmacology and activate adenylate cyclase via coupling to a Gs protein and activation of protein kinase A. Subsequently, D1‐like receptors have a high affinity for benzazepines like SCH‐23390 and exhibit low affinity for benzamides (sulpiride). The D2‐like receptors compose D2, D3, and D4 receptors. Molecular cloning has demonstrated the presence of two isoforms of D2 receptors, designated D2long and D2short (Giros et al., 1989). Like D2 receptors, the receptors of type D3 also exist in different isoforms. D2‐like receptors are with similar pharmacology; they inhibit adenylate cyclase via coupling to Gi/Go proteins. D3 receptors express relatively high affinity for atypical antipsychotics and for dopamine autoreceptor inhibitors [(þ)UH‐232, (þ)AJ‐76] while D4 receptors have high affinity for clozapine. D3 receptors may in part modulate the synthesis and release of dopamine in striatum and mesolimbic regions. Some antipsychotics exhibit high to moderate affinity to D4 dopamine receptors.

6.2 Structure of Dopamine Receptors All dopamine receptor subtypes are members of the large G protein‐coupled receptor superfamily, which is characterized by seven transmembrane hydrophobic domains, an extracellular N‐ and an intracellular C terminus. Dopamine receptors contain one aspartic acid and two serine residues in transmembrane

Dopamine and the dopaminergic systems of the brain

7

domains 3 and 5, which may bind the amino and hydroxyl groups of dopamine. These receptors are subject to posttranslational modifications including glycosylation, palmytoilation, and phosphorylation. The glycosylation sites at asparagines in the N terminus are essential for the transport of receptor protein through the cell and proper folding within the plasma membrane. Cysteines in the first and second extracellular loops form disulfide bond that has importance to maintain the three‐dimensional structure of the protein within the membrane. Sequences of phosphorylation are found in the second and third intracellular loops and the C terminal tail of the receptor. D1 receptors possess small i3 loops and long C‐tails and D2 receptors have large i3 loops and short C‐tails. Dopamine receptors form a packet‐like structure in which dopamine is recognized and bound. The binding of dopamine to a membrane‐bound receptor initiates a conformational change in the receptor such that it alters a G protein, which in turn is coupled to ion channels or second messenger systems. Two regions in the third intracellular loop are essential for binding of receptors to G protein a subunit.

6.3 Postsynaptic Dopamine Receptors Response to postsynaptic D1 and D2 receptor stimulation either activates or inhibits a messenger cascade involving the phosphorylation of a dopamine‐ and cAMP‐regulated phosphoprotein, DARPP‐32 (Greengard et al., 1999). The DARPP‐32 signaling pathway has a central role in mediating signal transduction within neurons like GABAergic medium‐sized spiny neurons in the striatum. Stimulation of D1 receptors by dopamine acts via cAMP and protein kinase A to phosphorylate phosphoprotein DARPP‐32, which in turn inhibits the activity of protein phosphatase‐1 (PP‐1). Besides changes in signal transduction system, electrophysiological changes after D1 receptor stimulation have also been reported. Thus, D1 receptor stimulation in the striatum reduces fast sodium conductance and N‐ and P‐type calcium currents and also enhances L‐type Ca2þ currents via a protein kinase A‐mediated process (Grace, 2002). In contrast, stimulation of D2 receptors leads to calcium stimulation of protein phosphatase‐2B. D2 receptor activation in enkephalin‐containing striatal GABAergic neurons (striatopallidal pathway) causes dephosphorylation of DARPP‐32 by Ca2þ influx‐activated calcineurin. Thus, D1 and D2 receptors exert opposite effects on centrally positioned DARPP‐32 in the signal transduction of striatal GABA neurons. Furthermore, dopamine agonists can exert excitation on GABA release within the striatum via D1 receptors and an inhibitory influence via D2 receptors as shown in a functional assay (Harsing and Zigmond, 1997).

6.4 Presynaptic Dopamine Receptors Whereas postsynaptic D2 receptors are associated with intracellular signaling, presynaptic dopamine D2 receptors regulate the release and synthesis of dopamine as well as the firing activity of dopaminergic neurons (> Figure 7-2). This regulation is primarily inhibitory in nature as activation of dopamine autoreceptors on the same presynaptic terminals can curtail the release and synthesis of dopamine, whereas those reside on cell bodies reduce neuronal firing activity. All three types of dopamine autoreceptors belong to the D2 family of dopamine receptors, which includes D2, D3, and D4 receptors. Presynaptic D2 dopamine receptors serve as autoreceptors because their activation can inhibit the cells by responding dopamine released from the same neurons; this kind of regulation is often designated as negative feedback regulation. Autoreceptors are distributed in all parts of the neurons, including the soma, the dendrites, and the nerve terminals. The release‐mediating dopamine autoreceptors at the nerve terminal respond to the transmitter released into the synaptic cleft, and those located on the cell body may respond to dendritic dopamine release. Release‐mediated autoreceptors are coupled to Gi protein and they dampen further release of dopamine by one or more of the following mechanisms (Bagdy and Harsing, 1995; Nestler et al., 2001): 1. Inhibitions of G protein‐coupled voltage‐sensitive Ca2þ channels that leads to restriction of available Ca2þ for depolarization.

161

162

7 2. 3.

Dopamine and the dopaminergic systems of the brain

Activation of the inwardly rectifying Kþ channels, which may increase potassium ion permeability through the cell membrane. Inhibition of presynaptically located receptor‐coupled adenylate cyclase, which is mediated by Gi/o protein. This may cause inhibition of protein kinase A that regulates proteins involved in neurotransmitter release.

D2 autoreceptors with different functions may be coupled to distinct transduction mechanisms. The released dopamine acts homeostatically at the synthesis‐modulating autoreceptors to control neurotransmitter synthesis. Dopamine receptors directly regulate dopamine synthesis: dopamine agonists decrease and antagonists increase synthesis of the neurotransmitter. Synthesis‐modulating autoreceptors are not present on all neurons; some midbrain dopamine neurons projecting to the prefrontal cortex appear to lack synthesis‐modulating autoreceptors. Synthesis‐regulating dopamine autoreceptors may belong to D3 rather than the D2 subtype in the striatum (Meller et al., 1993). Dopamine‐containing neurons in the midbrain exhibit spontaneous firing that is driven by an endogenous pacemaker conductance with their activity modulated by autoreceptors and afferent inputs. One of the prominent regulators of dopamine neuronal activity is the impulse‐mediating dopamine autoreceptor. These autoreceptors are located on the soma and dendrites of dopamine neurons. Impulse‐mediating autoreceptors are believed to exert a tonic downregulation of dopamine neuron activity, maintaining their firing rate within a stable range of activity. Somatodendritic autoreceptors are stimulated by an extracellular pool of dopamine released from the dendrites of the same or neighboring dopamine neurons. Dopamine agonists inhibit spike firing and corresponding antagonists reverse this effect.

6.5 Regional Distribution of Dopamine Autoreceptors Assessed by the g‐Butyrolactone Model g‐Butyrolactone (GBL) increases the concentrations of dopamine in several regions of the rat brain (Roth, 1984). This increase in dopamine levels is due to inhibition of impulse flow in dopamine neurons and the reduced release leads to disinhibition of synthesis‐mediated autoreceptors. The GBL‐evoked increase in dopamine synthesis can be further enhanced with inhibition of dopa decarboxylase elicited by concomitant administration of NSD‐1015 (Harsing and Vizi, 1991). The increase in dopamine levels after GBL injection can be antagonized by the D2 receptor agonist apomorphine in those brain areas where dopaminergic terminals possess dopamine‐sensitive autoreceptors. The use of this technique led to differentiate among dopamine nerve terminals: as those in the striatum, olfactorial bulb, amygdala, and piriform cortex express dopamine‐sensitive autoreceptors, whereas they are absent in dopaminergic nerve terminals of the prefrontal, cingulated, and entorthinal cortices. GBL pretreatment also decreases the formation of 3‐methoxytyramine, an indirect index for dopamine release in rat striatum (Westerink and Spaan, 1982).

6.6 Changes in Dopamine Receptor Sensitivity and Expression Receptor desensitization may occur when transmitter no longer causes a cellular response (Kuhar et al., 2006). Chronic agonist stimulation may induce this phenomenon. Desensitization may be due to receptor phosphorylation by protein kinase A, protein kinase C, or G protein‐coupled receptor kinases. A related phenomenon is termed downregulation, which usually occurs with a slower time course and involves cellular adaptations such as receptor degradation. MAO inhibitors and tricyclic antidepressants that increase concentrations of dopamine within the synaptic cleft leads to functional subsensitivity. Partial lesion of the dopamine system results in dopamine receptor supersensitivity in the remaining neurons. Destruction of neurons with neurotoxins such as 6‐hydroxydopamine evokes functional supersensitivity. Supersensitivity can result from a rapid loss of release sites following denervation. Chronic

Dopamine and the dopaminergic systems of the brain

7

administration of antagonists increases the density of D1 and D2 receptors, this increase in postsynaptic sites occurs in a longer time course. Supersensitivity of dopamine receptors that have been chronically blocked during treatment of patients with first generation antipsychotic drugs leads to development of excessive motor activity called tardive dyskinesia.

7

Dopamine Release from Neuronal Stores

Dopamine release may occur from axon terminals or dendrites of neurons by action potential propagation or reverse‐mode operation of the plasma dopamine transporter. Agonists of ion channel‐coupled receptors (glutamate or nicotinic receptors) may also increase neurotransmitter outflow from dopamine neurons. The mechanisms of these processes are different.

7.1 Action Potential Propagation‐Induced Dopamine Release Membrane depolarization induced by action potential propagation evokes dopamine release from synaptic vesicles located in nerve endings (> Figure 7-4). This release occurs from storage vesicles by the

. Figure 7-4 The time course of [3H]dopamine release measured from striatal slices. Slices were prepared from rat brain, loaded with [3H]dopamine and superfused with Krebs‐bicarbonate buffer. The slices were stimulated electrically (20 V, 2 Hz, 2‐msec for 2 min in fractions 4(S1) and 15(S2)) and the fractional release of [3H]dopamine (i.e., a percentage of the amount of [3H]dopamine in the tissue at the time of the release) was calculated. The calculated ratio of fractional release S2 over fractional release S1 (S2/S1) was 1.021. The radioactivity released from the tissue and that remained in the tissue was determined by liquid scintillation spectrometry

mechanism of exocytosis. Vesicular dopamine release is an external Ca2þ‐dependent process, as increase in free Ca2þ concentrations in the cytosol triggers fusion of secretory vesicles with plasma membrane and lack of free Ca2þ abolishes it. The rise of intracellular free Ca2þ is a consequence of Ca2þ entry through plasma membrane Ca2þ channels and other entry processes like opening of receptor‐coupled ion channels

163

164

7

Dopamine and the dopaminergic systems of the brain

permeable to bivalent cations. Release of Ca2þ from intracellular stores may have less importance in the exocytotic dopamine release. Membrane depolarization induced by elevated external potassium concentration leads to increase in dopamine release and this release also needs the presence of external Ca2þ. Dopamine release evoked by electrical stimulation, which mimics action potential propagation, can be abolished by tetrodotoxin, a drug which blocks voltage‐dependent sodium channels. On the contrary, tetrodotoxin does not abolish dopamine release evoked by high potassium depolarization, a stimulation procedure confines to the plasma membrane of the nerve terminals. Dopamine release that occurs typically by a Ca2þ‐dependent process is also designated as phasic release (Grace, 1991). The compartment, where phasic dopamine release originates from, can be characterized by carbon fiber voltammetry that allows to measure dopamine release in a real time base. Dopamine released from nerve ending with a phasic manner is removed from the synaptic cleft by the process of reuptake. The reuptake process does not capture all dopamine released as some may diffuse out from the synaptic cleft establishing dopamine pool in the biophase. Dopamine present in the extrasynaptic space represents the bases of nonsynaptic dopaminergic neurotransmission (Vizi, 2000) and it is often called tonic dopamine. Extrasynaptic compartments of tonic dopamine can be sampled by brain microdialysis, a technique that exhibits slower measures of dopamine dynamics. Phasic versus tonic dopamine release has been shown to have importance in normal and dysfunctional dopamine regulations related to certain psychiatric disorders such as schizophrenia, attention deficit hyperactivity disorder or drug abuse (Grace, 1991). From dendrites, dopamine can be released through a process that may not necessarily involve conventional exocytosis, i.e., the release, according to some observations, is not Ca2þ‐dependent.

7.2 Dopamine Release Evoked by Reverse‐Mode Operation of Dopamine Transporters Dopamine active transport could be bidirectional and able to evoke release by the same exchange‐diffusion process as that involved in the uptake function. Amphetamine and its analogs, methamphetamine and phenmetrazine, evoke direct release of dopamine and also inhibit its reuptake; these effects can be demonstrated both in vivo and in vitro. The releasing effect of amphetamine and its derivatives is mediated by outward transport of dopamine (Leviel, 2001). The releasing molecules that evoke dopamine release by reverse‐mode operation of the transporter induce the following steps: 1. Dopamine releaser drugs enter axon terminals by an operation of the transporter in the normal mode. 2. The releaser molecules evoke intraterminal transfer of dopamine from an electrically releasable pool to cytosolic compartment and thus dopamine concentrations increase within the terminals. 3. Dopamine is then transported out from the nerve terminals by reverse‐mode operation of the transporter. The release of dopamine by exchange‐diffusion is temperature‐ and Naþ‐dependent, saturable, and stereoselective. Exocytosis requires extracellular Ca2þ but dopamine transporter‐mediated outward transport does not. This effect is also not receptor‐mediated. Protein kinase C is involved in the external Ca2þ‐ independent dopamine release: activators of protein kinase C result in increase of dopamine efflux even in the absence of external Ca2þ (Gnegy, 2003). (þ)Amphetamine releases dopamine from a pool that is insensitive to reserpine but dependent on newly synthesized dopamine (i.e., a‐methyl‐p‐tyrosine sensitive). Other classes of drugs that inhibit dopamine reuptake (methylphenidate, cocaine, and amphenolic acid) do not induce dopamine release in vitro but they block the dopamine releasing effect of (þ)amphetamine. Methylphenidate evokes ‘‘neurogenic overflow’’ of dopamine in vivo and this effect may be explained by increased exchange of dopamine from the large, reserpine‐sensitive storage pool, too much smaller releasable sites (McMillen, 1983). This effect of methylphenidate has first been demonstrated as a potentiation of dopaminergic drugs in animal behavioral tests.

Dopamine and the dopaminergic systems of the brain

7

7.3 Dopamine Release Evoked by Ion Channel‐Coupled Receptors Ligands for ion channel‐coupled receptors (glutamate, nicotinic) can release dopamine by a mechanism that involves reverse‐mode operation of dopamine transporter and, in some cases, vesicular process as well (Lonart and Zigmond, 1991). Thus, glutamate is taken up into dopamine nerve endings by heterotransporter and Naþ is cotransported during this process. This leads a rise in intracellular Naþ concentrations, which promotes reduced membrane potential across nerve terminals. Enhanced intracellular Naþ concentrations shift dopamine transporter operation into reverse mode resulting in an increase in dopamine efflux from the nerve terminals. In addition, activation of ionotropic glutamate receptors permits influx of Naþ through receptor‐coupled ion channels and the rise of intracellular Naþ causes further reversal of Naþ‐dependent dopamine transport with a consequent efflux of dopamine from cytoplasmic stores. Moreover, excitatory amino acids also induce Ca2þ entry through receptor‐coupled ion channels followed by a further increase in intracellular Ca2þ concentrations due to opening of voltage‐dependent Ca2þ channels. The rise in intracellular Ca2þ concentrations may then be utilized for dopamine release process originating from nerve ending neurotransmitter stores. Activation of nicotinic receptors by agonists may also directly open ligand‐gated ion channels that are permeable to monovalent cations and also Ca2þ (Wonnacott et al., 1995). When nicotine binds to and activates nicotinic receptors, Naþ enters the cells through nicotinic receptor‐coupled ion channels inducing local membrane depolarization. Activation of nicotinic receptors may elevate free Ca2þ intracellularly in an indirect way by depolarizing the cell membrane enough for Ca2þ entry through voltage sensitive Ca2þ channels. As a consequence of the enhanced free intracellular Ca2þ, an exocytotic process may trigger dopamine release from vesicular pool. In many experimental conditions, dopamine release evoked by nicotine has been reported to be partly external Ca2þ‐dependent (Harsing et al., 1992).

7.4 Regulation of Dopamine Release The activity of dopaminergic neurons is determined by at least three different events. These are (1) the spontaneous discharge activity of dopamine neurons, (2) the autoinhibitory properties that include regulation of release, synthesis, and neural firing rate, and (3) excitatory and inhibitory afferent inputs mediated by heteroreceptor population of dopamine neurons. The glutamate‐ and GABA heteroreceptor‐ mediated regulation of dopamine release is particularly well characterized in the striatum. In addition, dopamine release can also be controlled by number local factors such as nitric oxide (NO) that is located released from striatal interneurons. When added, substrates for nitric oxide synthase (NOS) or NO generator compounds stimulate dopamine release in a Ca2þ‐dependent fashion (West and Galloway, 1997). The NO system and excitatory amino acids may interact with dopamine neuronal firing to regulate dopamine release from presynaptic sites in the striatum.

8

Dopaminergic Innervations in the Central Nervous System

A great portion of dopaminergic neurons in the central nervous system is located in three dopaminergic nuclei, the substantia nigra pars compacta (area A9), the ventral tegmental area (area A10), and the arcuate nucleus. Pathway originating from the substantia nigra pars compacta projects to the caudate nucleus and putamen and it forms the nigrostriatal dopaminergic system. Neurons from the ventral tegmental area largely project to the limbic structures like nucleus accumbens, prefrontal cortex, and cingulate cortex. These projections are designated as mesocortical/mesolimbic dopaminergic pathways. From the hypothalamic arcuate nucleus, dopaminergic neurons project to the pituitary gland, this system is designated as the tuberoinfundibular dopaminergic system. The tuberoinfundibular intermediate‐length dopaminergic system controls prolactin release from the anterior pituitary and its blockade by antipsychotics leads to

165

166

7

Dopamine and the dopaminergic systems of the brain

neuroendocrine effects (hyperprolactinemia or lactation) characteristic for this class of drugs. Other dopaminergic neurons distribute in the retina (ultrashort dopaminergic system) and other areas like the olfactory bulb and the lemniscal area (Cooper et al., 1996).

8.1 Dopamine in the Striatum In the neostriatum, D1 receptors are predominantly present in the GABAergic striatonigral GABAergic neurons, which contain substance P and dynorphin as cotransmitters, and project to the substantia nigra. D2 receptors occur mainly in striatopallidal GABAergic neurons, these neurons also contain enkephalins. Dopamine exerts inhibitory action on corticostriatal glutamatergic axon terminals, and this effect is mediated by D2 receptors (Harsing and Vizi, 1991). Striatal dopamine release controls movement patterns in physiological conditions and voluntary movement is impaired when dopaminergic tone is impaired by dopamine cell loss. Dopamine acting on D1 and D2 receptors influences two opposite types of synaptic plasticity, in the striatum, the long‐term potentiation and long‐term depression. Such plasticity within the striatum may be involved in acquisition of complete motor skills. The nigrostriatal neurons are the neural substrate for antiparkinsonian drugs and antipsychotic agents induce extrapyramidal side effects.

8.2 Dopamine in the Cerebral Cortex Cortical pyramidal neurons in layers V and VI receive dopaminergic influence from the ventral tegmental area and the released dopamine acts on D1/D5 receptors on apical dendrites of the pyramidal neurons. Dopamine terminals in the prefrontal cortex do not contain dopamine transporters (Lewis and Gonzales‐ Burgos, 2006). As a consequence, dopamine released from these sites would be free to diffuse to a much greater extent. The mesocortical and mesolimbic dopaminergic neurons are involved in cognitive and emotive functions and in the pathophysiology of various forms of psychosis. These dopaminergic systems are widely used to explain the mode of action of antipsychotic drugs and chemicals inducing drug abuse. The reward pathway is also well characterized: dopaminergic connection between the ventral tegmental areas and the nucleus accumbens mediating reinforcing properties of drugs of abuse. Stress causes dopamine release in the amygdala and lesion of amygdala tends to block stress‐induced increases in dopamine levels in the prefrontal cortex.

9

Altered Dopaminergic Neurotransmission

9.1 Neurotoxins Used for Destruction of Dopaminergic Neurons The dopamine analog 6‐hydroxydopamine, when it is administered directly into brain tissue, is selectively transported into dopaminergic axon terminals via high affinity uptake system. 6‐hydroxydopamine is then readily oxidizes to form of a series of cytotoxic compounds, 6‐hydroxydopamine‐p‐quinone, hydrogen  peroxide (H2O2), superoxide anion (O 2 ), and hydroxyl radical (OH ) (Zigmond et al., 1992). The accumulation of potentially cytotoxic compounds destroys dopaminergic elements by lipid peroxidation and protein and DNA damage. Dopamine uptake inhibitors can suspended the neurotoxic effect of 6‐hydroxydopamine whereas the MAO inhibitor pargyline potentiates it. Neurochemical changes after 6‐hydroxydopamine‐induced lesion involve increase dopamine release, synthesis and turnover rate, and reduced dopamine uptake (Zigmond, 1990; Juranyi et al., 2004). The 6‐hydroxydopamine‐induced destruction of dopamine neurons leads to severe neurological deficits, such as akinesia, reduce food and water intake, and lack of response to sensory stimuli. Dopamine agonists or muscarinic antagonists can reverse akinesia.

Dopamine and the dopaminergic systems of the brain

7

1‐Methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine (MPTP) is a selective neurotoxin for dopaminergic neurons, which is commonly used to induce experimental Parkinson’s disease in mice and monkeys (Heikkila and Sonsalla, 1987). MPTP enters the brain and B form of MAO converts it to 1‐methyl‐4‐ phenylpyridinium (MPPþ) (Heikkila et al., 1984). This product is then taken up into dopaminergic nerve terminals by dopamine transporters and it either inhibits complex II of the respiratory chain or generate oxygen‐reactive species within the nerve endings (Marsden, 2006). MPTP‐induced degeneration of dopaminergic neurons can be suspended with inhibition of MAO‐B by L‐deprenyl or by selective dopamine uptake blockers, mazidol or nomifensine (Javitch et al., 1985). Knockout mice lacking dopamine transporter express no sensitivity to the neurotoxin MPTP.

9.2 Dopaminergic Neurotransmission in Knockout Mice Tyrosine hydroxylase knockout mouse is not viable, whereas mice lacking dopamine decarboxylase exhibit hypersensitive to amphetamine. It was reported that MAO and catecol‐O‐methyl‐transferase knockout mice show aggression and altered anxiety behavior. D2 receptor‐deficient animals showed reduced dopamine autoreceptor‐mediated cell firing inhibition. Absence of dopamine transporter in mice induces hyperactivity, which may be the consequence of hyperdopaminergic state (Gainetdinov et al., 2002). In dopamine transporter knockout mice, there is an increase in extracellular dopamine levels and delayed clearance of dopamine released, and increased dopamine synthesis was observed. Because there is a loss of autoreceptor‐ mediated tone, tyrosine hydroxylase is disinhibited due to a lack of intraneuronal dopamine and dopamine turnover is markedly increased. These changes in dopaminergic neurotransmission in knockout mice are similar to the normal function of prefrontal dopaminergic neurons.

10 Conclusions and Future Avenues The success in dopamine research is due to the fact that the pathology of a series of psychiatric and neurological disorders can be explained based upon their dopamine theory. The positive symptoms of schizophrenia (hallucination, delusion, thought disorder) and cognitive deficits characteristics for this disorder can satisfyingly be explained by hyperfunctionality of subcortical dopaminergic systems and a deficit in cortical dopaminergic neurotransmission (Lewis and Gonzales‐Burgos, 2006). Both the first and the second generations of antipsychotic drugs exert potent antagonistic effects on D2 dopamine receptors although the ratio of 5‐HT2A versus D2 receptor antagonism is more pronounced for the second generation antischizophrenic agents. Currently used antipsychotic agents exhibit a wild range of side effects due to the broad range of receptors on which these agents act. Drugs acting preferentially at D3 binding site (S33138, Millan et al., 2002) or as D2/D3 receptor antagonists/partial agonists (RGH‐188, Kiss et al., 2006) or D2/D3 antagonist with D4 partial agonistic effect (F15063, Newman‐Tancredi et al., 2006) represent a series of third generation antipsychotic compounds. Some of these drugs are now in Phase I/Phase II human clinical trials. It is strongly believed that compounds with these receptor‐binding profiles will lead to antipsychotic activity associated with lower incident of adverse side effects. Of the neurodegenerative disorders, Parkinson’s disease is characterized with loss of dopaminergic neurons in the substantia nigra pars compacta. The reduced dopaminergic tone and the consequent disbalance between the direct and indirect GABAergic projection neurons in the striatum may satisfyingly explain the symptoms of Parkinson’s disease (tremor at rest, bradykinesia, and muscle rigidity). The impaired dopaminergic influence may be enhanced by addition of the dopamine precursor L‐dopa in combination with the peripheral dopa decarboxylase inhibitor carbidopa. Besides supplementary therapy, inhibition of dopamine breakdown can also be of therapeutic value: the B type MAO inhibitor L‐deprenyl is used in the therapy of Parkinson’s disease for this purpose. Rasagiline, another B type MAO inhibitor, has also been shown to be effective in treatment of Parkinson’s disease (Siderowf, 2002).

167

168

7

Dopamine and the dopaminergic systems of the brain

Acknowledgement This work was supported in part by grants of the Hungarian Research Foundation (OTKA T‐43511) and Hungarian Medical Research Council (ETT‐482/2003). The author acknowledges the editorial work of Ms. Judit Puskas.

References Bagdy E, Harsing LG Jr. 1995. The role of various calcium and potassium channels in the regulation of somatodendritic serotonin release. Neurochem Res 20: 1409-1415. Bergquist F, Nissbrandt H. 2005. Dopamine release in substantia nigra: Release mechanisms and physiological function in motor control. Dendritic Neurotransmitter Release. Ludwig M, editor. New York: Springer; pp. 85-99. Cooper JR, Bloom FE, Roth RH. 1996. Dopamine. The Biochemical Basis of Neuropharmacology. Oxford University Press; New York: pp. 293-351. Dahlstrom A, Fuxe K. 1964. Evidence for the existence of monoamine containing neurons in the central nervous system. 1. Demonstration of monoamines in the cell bodies of brain stem neurons. Acta Physiol Scand Suppl 232: 1-55. de Langen CDJ, Mulder AH. 1979. Compartmental analysis of the accumulation of 3H‐dopamine in synaptosomes from rat corpus striatum. Naunyn Schmiedebergs Arch Pharmacol 308: 31-39. Demarest KT, Smith DJ, Azzaro AJ. 1980. The presence of the type A form of monoamine oxidase within nigrostriatal dopamine‐containing neurons. J Pharmacol Exp Ther 215: 461-468. Deutch AY, Roth RH. 2004. Pharmacology and biochemistry of synaptic transmission: Classic transmitters. From Molecules to Network. Byrne JH, Roberts JL, editors. New York: Elsevier Academic Press; pp. 245-278. Gainetdinov RR, Sotnikova TD, Caron MG. 2002. Monoamine transporter pharmacology and mutant mice. Trends Pharmacol Sci 23: 367-373. Gether U, Andersen PH, Larsson OM, Schousboe A. 2006. Neurotransmitter transporters: Molecular function of important drug targets. Trends Pharmacol Sci 27: 375-383. Giros B, Sokoloff P, Martres MP, Riou JF, Emorine LJ, et al. 1989. Alternative splicing directs the expression of two D2 dopamine receptor isoforms. Nature 342: 923-926. Glover V, Sandler M, Owen F, Riley GJ. 1977. Dopamine is a monoamine oxidase B substrate in man. Nature 265: 80-81. Gnegy ME. 2003. The effect of phosphorylation on amphetamine‐mediated outward transport. Eur J Pharmacol 479: 83-91. Grace AA. 1991. Phasic versus tonic dopamine release and the modulation of dopamine system responsivity: A hypothesis for the etiology of schizophrenia. Neuroscience 41: 1-24.

Grace AA. 2002. Dopamine. Neuropsychopharmacology: The Fifth Generation of Progress. Davis KL, Charney D, Coyle JT, Nemeroff C, editors. New York: Lippincott Williams and Wilkins; pp. 119-132. Greengard P, Allen PB, Nairn AC. 1999. Beyond the dopamine receptor: The DARPP‐32/protein phosphatase‐1 cascade. Neuron 23: 435-447. Gresch PJ, Sved AF, Zigmond MJ, Finlay JM. 1995. Local influence of endogenous norepinephrine on extracellular dopamine in rat medial prefrontal cortex. J Neurochem 65: 111-116. Hammond C. 1996. Neurotransmitter release. Cellular and Molecular Neurobiology. Hammond C, editor. New York: Academic Press; pp. 188-214. Harsing LG Jr, Vizi ES. 1991. Alpha 2‐adrenoceptors are not involved in the regulation of striatal glutamate release: Comparison to dopaminergic inhibition. J Neurosci Res 28: 376-381. Harsing LG Jr, Sershen H, Vizi SE, Lajtha A. 1992. N‐type calcium channels are involved in the dopamine releasing effect of nicotine. Neurochem Res 17: 729-734. Harsing LG Jr, Zigmond MJ. 1997. Influence of dopamine on GABA release in striatum: Evidence for D1‐D2 interactions and non‐synaptic influences. Neuroscience 77: 419-429. Harsing LG Jr. 2006. The pharmacology of the neurochemical transmission in the midbrain raphe nuclei of the rat. Curr Neuropharmacol 4: 313-339. Heal DJ, Frankland AT, Buckett WR. 1990. A new and highly sensitive method for measuring 3‐methoxytyramine using HPLC with electrochemical detection. Studies with drugs which alter dopamine metabolism in the brain. Neuropharmacology 29: 1141-1150. Heikkila RE, Manzino L, Cabbat FS, Duvoisin RC. 1984. Protection against the dopaminergic neurotoxicity of 1‐methyl‐4‐phenyl‐1,2,5,6‐tetrahydropyridine by monoamine oxidase inhibitors. Nature 311: 467-469. Heikkila RE, Sonsalla PK. 1987. The use of the MPTP‐treated mouse as an animal model of parkinsonism. Can J Neurol Sci 14: 436-440. Javitch JA, D’Amato RJ, Strittmatter SM, Snyder SH. 1985. Parkinsonism‐inducing neurotoxin, N‐methyl‐4‐ phenyl‐1,2,3,6‐tetrahydropyridine: Uptake of the metabolite N‐methyl‐4‐phenylpyridine by dopamine neurons

Dopamine and the dopaminergic systems of the brain explains selective toxicity. Proc Natl Acad Sci USA 82: 2173-2177. Juranyi Z, Sziray N, Marko B, Levay G, Harsing LG Jr. 2004. AMPA receptor blockade potentiates the stimulatory effect of L‐DOPA on dopamine release in dopamine‐deficient corticostriatal slice preparation. Crit Rev Neurobiol 16: 129-139. Justice JB Jr, Nicolaysen LC, Michael AC. 1988. Modeling the dopaminergic nerve terminal. J Neurosci Methods 22: 239-252. Kebabian JW, Calne DB. 1979. Multiple receptors for dopamine. Nature 277: 93-96. Kiss B, Schmidt E, Horvath A, Orosz S, Laszlovszky I, et al. 2006. Neurochemical profile of RGH‐188, a novel dopamine D3/D2 antagonist/partial agonist. Soc Neurosci 36: Abstr 93.2. Kuhar MJ, Minneman K, Muly EC. 2006. Catecholamines. Basic Neurochemistry, Molecular, Cellular, and Medical Aspects. Siegel GJ, Albers RW, Brady ST, Price DL, editors. New York: Elsevier Academic Press; pp. 211-223. Leviel V. 2001. The reverse transport of DA, what physiological significance? Neurochem Int 38: 83-106. Lewis DA, Gonzalez‐Burgos G. 2006. Pathophysiologically based treatment interventions in schizophrenia. Nat Med 12: 1016-1022. Lonart G, Zigmond MJ. 1991. High glutamate concentrations evoke Ca(þþ)‐independent dopamine release from striatal slices: A possible role of reverse dopamine transport. J Pharmacol Exp Ther 256: 1132-1138. Marsden CA. 2006. Dopamine: The rewarding years. Br J Pharmacol 147 (Suppl 1): S136-S144. McMillen BA, German DC, Shore PA. 1980. Functional and pharmacological significance of brain dopamine and norepinephrine storage pools. Biochem Pharmacol 29: 3045-3050. McMillen BA. 1983. CNS stimulants: Two distinct mechanisms of action for amphetamine‐like drugs. Trends Pharmacol Sci 4: 429-432. Megyeri K, Marko B, Sziray N, Gacsalyi I, Juranyi Z, et al. 2007. Effects of 2,3‐benzodiazepine AMPA antagonists on dopamine turnover in the striatum of rats with experimental parkinsonism. Brain Res Bull 71: 501-507. Meller E, Bohmaker K, Goldstein M, Basham DA. 1993. Evidence that striatal synthesis‐inhibiting autoreceptors are dopamine D3 receptors. Eur J Pharmacol 249: R5-R6. Millan MJ, Cussac D, Newman‐Tancredi A, Gobert A, Lejeune F, et al. 2002. The novel benzopyrano‐pyrrole and antipsychotic, S33138, is a preferential antagonist at dopamine (DA) D3 vs D2 receptors. Am Soc Neurosci 32, Abstr 894.14. Nestler EJ, Hyman SE, Malenka RC. 2001. Molecular Neuropharmacology. A Foundation for Clinical Neuroscience. McGraw‐Hill; New York: pp. 115-137.

7

Newman‐Tancredi A, Assie M, Martel J, Cosi C, Heusler P, et al. 2006. F15063, an antipsychotic with D2/D3 antagonist, 5‐HT1A agonist, and D4 partial agonist properties: Activity in vitro and neurochemical profile in rodents. Soc Neurosci, 36: Abstr 93.6. Norregaard L, Gether U. 2001. The monoamine neurotransmitter transporters: Structure, conformational changes and molecular gating. Curr Opin Drug Discov Devel 4: 591-601. Ogawa N, Tanaka K, Asanuma M. 2000. Bromocriptine markedly suppresses levodopa‐induced abnormal increase of dopamine turnover in the parkinsonian striatum. Neurochem Res 25: 755-758. Roth RH. 1984. CNS dopamine autoreceptors: Distribution, pharmacology, and function. Ann N Y Acad Sci 430: 27-53. Siderowf A. 2002. A controlled trial of rasagiline in early Parkinson disease: The TEMPO study. Arch Neurol 59: 1937-1943. Snyder AM, Keller RW, Zigmond MJ. 1990. Dopamin efflux from striatal slices after intracerebral 6‐hydroxydopamine: Evidence for compensatory hyperactivity of residual terminals. J Pharm Exp Ther 253: 867-876. The IUPHAR Compendium of Receptor Characterization and Classification. 2000. 2nd edition, London: IUPHAR Media; pp. 170–181. Thony B, Auerbach G, Blau N. 2000. Tetrahydrobiopterin biosynthesis, regeneration and functions. Biochem J 347 Pt 1: 1-16. Vaughan RA. 2004. Phosphorylation and regulation of psychostimulant‐sensitive neurotransmitter transporters. J Pharmacol Exp Ther 310: 1-7. Vizi ES, Harsing LG Jr, Gaal J, Kapocsi J, Bernath S, et al. 1986. CH‐38083, a selective, potent antagonist of alpha‐2 adrenoceptors. J Pharmacol Exp Ther 238: 701-706. Vizi ES. 2000. Role of high‐affinity receptors and membrane transporters in nonsynaptic communication and drug action in the CNS. Pharmacol Rev 55: 8775-8779. Von Bohlen und Halbach O, Dermietzel R. 2002. Neurotransmitters and Neuromodulators. Wiley‐VCH Verlag GmbH; Weinheim: pp. 53-63. Walter DS, Flockhart IR, Haynes MJ, Howlett DR, Lane AC, et al. 1984. Effects of idazoxan on catecholamine systems in rat brain. Biochem Pharmacol 33: 2553-2557. Weihe E, Eiden LE. 2000. Chemical neuroanatomy of the vesicular amine transporters. FASEB J 14: 2435-2449. West AR, Galloway MP. 1997. Endogenous nitric oxide facilitates striatal dopamine and glutamate efflux in vivo: Role of ionotropic glutamate receptor‐dependent mechanisms. Neuropharmacology 36: 1571-1581. Westerink BH, Spaan SJ. 1982. On the significance of endogenous 3‐methoxytyramine for the effects of centrally acting

169

170

7

Dopamine and the dopaminergic systems of the brain

drugs on dopamine release in the rat brain. J Neurochem 38: 680-686. Wonnacott S, Gothert M, Chahl LA, Willow M, Nicholson GM. 1995. Modulation of neurotransmitter release by some therapeutic and socially used drugs. Neurotransmitter release and its Modulation. Powis DA, Bunn SJ, editors. Cambridge University Press; pp. 293–328.

Zigmond MJ. 1990. Compensations after lesions of central dopaminergic neurons: Some clinical and basic implications. Trends Neurosci 13: 290-296. Zigmond MJ, Hastings TG, Abercrombie ED. 1992. Neurochemical responses to 6‐hydroxydopamine and L‐dopa therapy: Implications for Parkinson’s disease. Ann N Y Acad Sci 648: 71-86.

8

5‐Hydroxytryptamine in the Central Nervous System

A. C. Dutton . N. M. Barnes

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

2

Neuroanatomy of the 5‐Hydroxytryptaminergic System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

3

Forebrain Projections of the Raphe Nuclei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

4

The Physiology of 5‐HT Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

5 5.1 5.1.1 5.1.2 5.1.3 5.1.4 5.1.5 5.2 5.2.1 5.2.2 5.2.3 5.3 5.4 5.5 5.6 5.7

The 5‐HT Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174 The 5‐HT1 Receptor Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174 The 5‐HT1A Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175 The 5‐HT1B Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 178 The 5‐HT1D Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 The 5‐ht1E Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 182 The 5‐HT1F Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 182 The 5‐HT2 Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 5‐HT2A Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 The 5‐HT2B Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185 The 5‐HT2C Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186 The 5‐HT3 Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 The 5‐HT4 Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 The 5‐ht5 Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 190 The 5‐HT6 Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 192 The 5‐HT7 Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194

6

The 5‐HT Transporter (SERT) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 196

7

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 198

#

2008 Springer ScienceþBusiness Media, LLC.

172

8

5‐Hydroxytryptamine in the central nervous system

Abstract: 5-Hydroxytryptamine (5-HT, serotonin), mediates numerous physiological processes in the CNS. The diversity of function is at least partly a consequence of the 14 distinct receptors evident in mammals. The present review describes the biochemistry, physiology and pharmacology of the 5-HT system in the brain. List of Abbreviations: AHP, afterhyperpolarization potential; APP, amyloid precursor protein; BDNF, brain‐derived neurotrophic factor; EPSCs, excitatory postsynaptic currents; GPCR, G‐protein‐coupled receptor; IPSCs, inhibitory postsynaptic currents; NREMS, Nonrapid eye movement sleep; OCD, obsessive compulsive disorder; PET, positron emission tomography; PLC, phospholipase C; PPI, prepulse inhibition; REM, rapid eye movement; SAR, structure–affinity relationship; SCN, suprachiasmatic nucleus

1

Introduction

5‐Hydroxytryptamine (5‐HT, serotonin) acts as a neurotransmitter within the central nervous system (CNS) and peripheral nervous system, in addition to mediating nonneuronal actions in numerous other tissues including the gastrointestinal tract and blood vessels. The actions of 5‐HT are, therefore, considered to be copious and diverse. With respect to its role as a neurotransmitter in the brain, which will be the primary focus of this chapter, 5‐HT is implicated in the processes of mood, sleep, aggression, cognition, memory, emesis, and feeding behavior, as well as the pathophysiology of disorders including major depression, schizophrenia, obsessive–compulsive disorder, and anxiety. The impact of this plethora of roles has consequently led to the actions of 5‐HT being extensively studied, leading to the development of many compounds of therapeutic value, including antidepressants, antipsychotics, and antiemetics. This chapter begins with an outline of the 5‐hydroxytryptaminergic system in the brain, including the origins and projections of 5‐HT‐containing neurons, subsequently focusing on the roles of the individual 5‐HT receptors (> Table 8-1) and the transporter in both physiological and pathophysiological processes.

2

Neuroanatomy of the 5‐Hydroxytryptaminergic System

The cell bodies of 5‐HT neurons are situated along the rostrocaudal midline of the brain stem, as first identified by Dahlstro¨m and Fuxe in 1964 (Dahlstro¨m and Fuxe, 1964; see Hornung, 2003, for an in‐depth review), and are categorized into nine anatomical groups, named B1–9. The dorsal raphe nucleus (B6, B7), median raphe nucleus (B8), and B9 contain
85% of the 5‐HT neurons found within the brain, and project extensively to widespread regions of the forebrain. The remaining raphe nuclei, B1–4, innervate primarily both the brain stem and spinal cord. Although the raphe nuclei contain predominantly 5‐HT neurons, it should be noted that other neuronal phenotypes are also apparent.

3

Forebrain Projections of the Raphe Nuclei

Raphe nuclei neurons project to widespread regions throughout the cerebral hemispheres (see Hensler, 2006, for review) via two main pathways, the dorsal periventricular pathway and the ventral tegmental radiation, which unite in the hypothalamus before continuing along the medial forebrain bundle. The hypothalamus, medial septum, and dorsal hippocampus receive predominant innervation from the median raphe nucleus, whereas the ventral hippocampus, amygdala, lateral septum, striatum, and prefrontal cortex (PFC) contain 5‐HT neuron terminals largely from the dorsal raphe nucleus. The cerebral cortex receives inputs from both subdivisions, though various regions are thought to receive different degrees of median and dorsal raphe neuron innervation.

4

The Physiology of 5‐HT Neurons

5‐HT neurons have distinctive electrophysiological properties (e.g., Aghajanian and Haigler, 1974), confirmed using a combination of electrophysiology and fluorescent immunohistochemistry, enabling the activity of 5‐HT‐immunopositive cells to be recorded (Beck et al., 2004). 5‐HT neurons appear to fire

8

5‐Hydroxytryptamine in the central nervous system . Table 8-1 Summary of the structure, pharmacology, and function of 5‐HT receptors Receptor Human gene

5‐HT1A 5q11.2–q13

5‐HT1B 6q13

5‐HT1D 1p34.3–36.3

Structure Transduction system

GPCR ↓cAMP G‐protein‐coupled‐Kþ current 8‐OH‐DPAT

GPCR ↓cAMP

GPCR ↓cAMP

Sumatriptan

Sumatriptan

(R)‐UH301

L 694247

PNU 109291

U92016A WAY 100635 (S)‐UH301 NAD299 (robalzotan) ↑Acetylcholine

GR 55562 SB 224289 SB 236057 ↓5‐HT

Agonists

5‐ht1E 6q14– q15 GPCR ↓cAMP

5‐HT1F 3q11 GPCR ↓cAMP

LY 344864 –

Antagonists

Effect on neurotransmission

Noradrenaline ↓Dopamine Therapeutic target

Depression Anxiety/stress/panic Aggression Cognition

L 694247

↑Acetylcholine ↑Glutamate ↓Dopamine Depression Anxiety Aggression Migraine Drug addiction

Receptor

5‐HT2A

5‐HT2B

5‐HT2C

Human gene

13q14–q21

2q36.3–2q37.1

Xq24

Structure Transduction system

GPCR ↑PLC

GPCR ↑PLC

Agonists

DOI

Antagonists

Ketanserin MDL 100907

GPCR ↑PLC DOI BW 723C86 Ro 600175 RS 127445 SB 200646 SB 204741

Effect on neurotransmission

↑Glutamate ↑Dopamine

Therapeutic target

Depression Anxiety Schizophrenia Cognition Eating disorders Sleep disorder?

LY 334370

BRL 15572 SB 714786

–

–

↑Glutamate

–

–

Migraine

–

Migraine

DOI Ro 600175 SB 242084 RS 102221

? ↓Dopamine

Depression Anxiety Sleep disorder? Migraine

Anxiety Obesity Cognition

5‐HT3 11q23.1–23.2 (A) 11q23.1 (B) 3q27 (C/D/E) LGIC Ion conductance (Kþ, Naþ, Ca2þ) 2‐Methyl 5‐HT SR 57227 m‐Chlorophenyl biguanide DOI Granisetron Ondansetron Tropisetron ↑5‐HT ↑Dopamine ↓Acetylcholine Emesis Anxiety Cognition Drug addiction Analgesia Chronic fatigue syndrome

173

174

8

5‐Hydroxytryptamine in the central nervous system

. Table 8-1 (Continued) Receptor Human gene

5‐HT4 5q31–q33

5‐ht5a 7q36

Structure

GPCR

Transduction system

↑cAMP

Agonists

Antagonists

Effect on neurotransmission

BIMU 8 RS 67506 ML 10302 GR 113808 SB 204070

5‐HT7 10q21–q24

GPCR

GPCR

Not known

↑cAMP

↑cAMP

5‐CT

5‐CT

–

8‐OH‐DPAT

? SB 699551‐ A

–

Ro 630563 SB 271046

SB 258719 SB 269970

SB 357134 ↓Acetylcholine

SB 656104

↑Dopamine ↓Glutamate Cognition Schizophrenia Depression

? ↑ ↓5‐HT

RS 100235 ↓Acetylcholine ↑Dopamine ↑5‐HT

Therapeutic target

5‐HT6 1p35–36

GPCR ↑cAMP ? Ca2þ mobilization ? Kþ current

5‐ht5B 2q11–13 (nonfunctional) GPCR

Cognition

Not known

Not known

Anxiety

Not known

Not known

Anxiety/stress Epilepsy

Depression Schizophrenia Sleep disorder Epilepsy Cognition

spontaneously with a slow, rhythmic activity, and exhibit a relatively long action potential in addition to a large afterhyperpolarization potential (AHP). The rhythmic activity is thought to be generated by a pacemaker cycle mediated by a calcium‐dependent potassium current (Aghajanian, 1990).

5

The 5‐HT Receptors

The ability of 5‐HT to mediate a diverse array of actions can be accounted for by the existence of an imposing number of 5‐HT receptors (Barnes and Sharp, 1999). Numerous 5‐HT receptor families and subtypes have been identified, particularly within the last two decades following the development of techniques in the field of molecular biology. Presently, there are 18 genes that give rise to 14 distinct mammalian 5‐HT receptor subtypes, divided into 7 families, the majority of which are members of the G‐protein‐coupled receptor (GPCR) superfamily, the sole exception being the 5‐HT3 receptor, a ligand‐ gated ion channel. Further, receptor multiplicity is generated through RNA editing (the 5‐HT2C receptor), alternative splicing (5‐HT3, 5‐HT4, and 5‐HT7 receptors), and the putative formation of homo‐ and heterodimers (5‐HT4 and the b2‐adrenoceptor; Berthouze et al., 2005).

5.1 The 5‐HT1 Receptor Family The 5‐HT1 receptor family contains five subtypes, 5‐HT1A, 5‐HT1B, 5‐HT1D, 5‐HT1E, and 5‐HT1F, each having distinct, but overlapping patterns of expression within the brain. This family can be characterized by its inhibitory effect on cellular cAMP levels, although the 5‐HT1A receptor can also activate a G‐protein‐ activated potassium channel independent of second‐messenger cascades involving cAMP.

5‐Hydroxytryptamine in the central nervous system

8

5.1.1 The 5‐HT1A Receptor In 1981, Pedigo et al. (Pedigo et al., 1981) identified the 5‐HT1A receptor‐binding site in rat brain, but the sequence encoding the receptor was not isolated until 1987 (Kobilka et al., 1987). The 5‐HT1A receptor inhibits adenylate cyclase activity through coupling to Gi/o proteins, which has been demonstrated both in native tissue (e.g., rat hippocampus) and recombinant cell systems (see Albert et al., 1996, for review). Such coupling has not, however, been observed in the raphe nuclei (Clarke et al., 1996), where receptor activation induces G‐protein‐modulated potassium current which is cAMP‐independent (Aghajanian, 1995). Expression of the 5-HT7 receptor within the brain region thought to control circadian rhythm, the suprachiasmatic nucleus (SCN), and the overlapping pharmacologies of the 5‐HT1A and 5‐HT7 receptors (e.g. activation by 8-OH-DPAT), complicates interpretation of results in this area of research. More recently, immunohistochemical studies using selective 5‐HT1A receptor antibodies have provided greater resolution of receptor expression through light and electron microscopy. Within the raphe nuclei, the 5‐HT1A receptor appears to be expressed somatodendritically by serotonergic neurons projecting to the forebrain, with dendritic receptors predominantly located in extrasynaptic regions (Kia et al., 1996b; Riad et al., 2000; > Figure 8-1a). The receptor is also found in many regions of the forebrain, including the frontal, piriform, and entorhinal cortices, the hippocampus, preoptic areas, lateral and medial septum, the diagonal band of Broca, hypothalamus, amygdala, and thalamic regions (Aznar et al., 2003). Within the isocortex, the receptor is expressed throughout all laminae (with the exception of layer I), where both glutamatergic pyramidal neurons and calbindin‐ and parvalbumin‐positive inhibitory g‐amino butyric acid (GABA)ergic interneurons express the receptor (Aznar et al., 2003). Within the hippocampus, granule and pyramidal cells are also believed to express the 5‐HT1A receptor on both the soma and dendrites (Riad et al., 2000; Aznar et al., 2003; > Figure 8-1b), particularly in the postsynaptic membrane, but also in nonsynaptic regions, of dendritic spines (Kia et al., 1996b). In the medial septum and diagonal band of Broca, the receptor appears to be expressed somatodendritically by cholinergic neurons (Kia et al., 1996a) and by inhibitory interneurons (Aznar et al., 2003). Given the apparent common subcellular location of the 5‐HT1A receptor, it has been suggested that a structural component of the protein may be responsible for targeting the receptor to a somatodendritic location (Darmon et al., 1998). Activation of the somatodendritic 5‐HT1A autoreceptor in the raphe nuclei induces membrane hyperpolarization, leading to reduced 5‐HT neuron excitability, firing, and ultimately a reduction in 5‐HT release in the raphe forebrain projection areas (Aghajanian, 1995; Sharp et al., 1996). 5‐HT1A receptor agonists also inhibit neuronal firing in forebrain regions, including the hippocampus (e.g., Sprouse and Aghajanian, 1988). The release of other neurotransmitters, including acetylcholine, noradrenaline, and dopamine, is thought to be regulated by 5‐HT1A receptor activation. For example, 8‐OH‐DPAT augments acetylcholine release in the hippocampus and cortex of guinea pigs (Bianchi et al., 1990; Wilkinson et al., 1994). The mechanism of this action is unclear, though more recent studies suggest that activation of noncortical presynaptic 5‐HT1A autoreceptors mediate the increase in acetylcholine release within the cortex (Millan et al., 2004), while locally administered 8‐OH‐DPAT‐induced elevation of acetylcholine levels in the rat dorsal hippocampus may be mediated by postsynaptic 5‐HT1A receptors (Nakai et al., 1998). In contrast, other reports suggest that 5‐HT1A receptor antagonists enhance acetylcholine release within the hippocampus by blocking tonically active inhibitory 5‐HT1A receptors on cholinergic cells (Millan et al., 2004), whereas noradrenaline levels in the hippocampus, ventral tegmental area (VTA), and hypothalamus increase following 5‐HT1A receptor activation (Done and Sharp, 1994; Chen and Reith, 1995; Suzuki et al., 1995). More recently, putative 5‐HT1A receptor‐mediated effects on dopamine release have been observed; whereby the 5‐HT1A receptor agonist BAYx3702 elevated dopamine release in both the VTA and medial PFC (mPFC), as well as increasing dopaminergic neuron activity in the VTA (Dı´az‐Mataix et al., 2005). 5‐HT1A receptors may also modulate glutamatergic neurotransmission. Indeed, 5‐HT1A receptor activation attenuates AMPA currents in pyramidal neurons of the PFC, potentially through a reduction in PKA‐dependent AMPA subunit phosphorylation (Cai et al., 2002). The activity of 5‐HT1A receptors may be involved in the pathogenesis and treatment of psychiatric disorders. One such condition that attracted much study is depression. It has been suggested that levels of 5‐HT1A receptors are altered in depressed subjects, although to date, studies to this effect remain

175

176

8

5‐Hydroxytryptamine in the central nervous system

. Figure 8-1 (a) Low‐power electron micrograph illustrating the somatodendritic localization of the 5‐HT1A immunoreactivity (immunoperoxidase labeling) in the NRD. Most of the field is occupied by the multipolar cell body of a presumed 5‐HT neuron (N1 in nucleus) showing strong immunolabeling of its perikaryon and emerging proximal dendrite (D). Smaller adjacent nerve cell bodies (N2, N3 in nuclei) are immunonegative. Numerous transversely sectioned dendritic branches of smaller calibre (d) are also immunoreactive in the surrounding neurophil, but myelinated axons and axon terminals are all immunonegative. Also note the immunonegativity of a nearby astrocyte (A), pericyte (P), and endothelial cell (E). In the labeled cell body and the largest of the dendritic branches, the diffusible immunoperoxidase precipitate is conspicuously concentrated on the inner face of the plasma membrane, as a rim almost 1 mm wide; in smaller dendritic processes, the confluence of these peripheral zones of 5‐HT1A immunoreactivity accounts for labeling of the entire sectional surface. This and the following electron micrograph were obtained from tissue fixed with acrolein plus paraformaldehyde. Scale bar ¼ 5 mm. (b) Immunoperoxidase labeling of several immunoreactive dendrites (d) in a small field from the stratum radiatum of CA3. The peroxidase immunoprecipitate tends to cluster near the plasma membrane and is also found in dendritic spines (arrows) seen to emerge from their parent dendritic branches in the upper left and lower right corners of the figure. Scale bar ¼ 1 mm (Taken from Riad et al., 2006, these figures are reproduced with permission from the authors)

5‐Hydroxytryptamine in the central nervous system

8

controversial. Some reports have suggested that the levels of the receptor in the hippocampus are reduced (Cheetham et al., 1990), whereas others detect no differences between suicide victims suffering from depression and control patients (Stockmeier et al., 1997). Alternatively, abnormally high levels of 5‐HT1A receptors were found in the PFC (Arango et al., 1995) and dorsal raphe nucleus (Stockmeier et al., 1998) of depressed suicide victims, whereas conversely other studies found no change in the cortex (e.g., Lowther et al., 1997) or reductions in the number of 5‐HT1A receptors in the dorsal raphe nucleus (Arango et al., 2001). Receptor levels aside, it has become apparent that compounds acting on the 5‐HT1A receptor may have therapeutic value in treating depression. The use of 5‐HT1A receptor antagonists in conjunction with selective serotonin reuptake inhibitors (SSRIs) has been suggested to reduce the onset of relief from depression (Artigas et al., 1994), although other reports have found no such benefits (Segrave and Nathan, 2005). The 5‐HT1A receptor has also been linked to depression through the ability of the 5‐HT1A receptor agonist, 8‐OH‐DPAT, following 5‐HT depletion, to induce granule cell proliferation within the dentate gyrus of the hippocampus (Huang and Herbert, 2005), a process thought to facilitate the treatment of depression (Malberg et al., 2000; Santarelli et al., 2003). Similarly, the 5‐HT1A receptor has been linked to suicidal behavior in addition to depression. Pitchot et al. (2005) have found that flesinoxan, a 5‐HT1A receptor agonist, attenuated cortisol and temperature responses in suicidal patients when compared with controls, but not in merely depressed patients, suggesting that reduced 5‐HT1A receptor sensitivity may be connected to suicidal tendencies. Initial reports using a 5‐HT1A receptor knockout mouse strain demonstrated various behavioral abnormalities compared with wild‐type mice, including increased levels of anxiety in the open‐field arena test and attenuated immobility in the forced‐swim test. This latter effect may represent either increased anxiety due to the inherent stress of the testing or an inhibition of behavioral despair, a measure of predisposition to depression (Parks et al., 1998; Ramboz et al., 1998). These observations correlate with the observation that 5‐HT1A receptor agonists are anxiolytic (Lucki et al., 1994). Indeed, buspirone, a partial 5‐HT1A receptor agonist, has therapeutic usage in the treatment of generalized anxiety disorder (Kapczinski et al., 2003), while exhibiting minimal side effects relative to those induced following administration of benzodiazepines for anxiety‐related disorders (Goa and Ward, 1986). Recently, new 5‐HT1A receptor ligands (oMPP derivatives), supposed postsynaptic antagonists and partial agonists, display anxiolytic activity when assessed in the Vogel conflict drinking test (Bojarski et al., 2006). 5‐HT1A receptor targeting, therefore, continues to provide a useful strategy in the search for novel treatments of anxiety and depression. In addition to anxiety, the 5‐HT1A receptor may also be involved in responses to chronic stress implicated in depression. It has been shown that chronic stress may lead to the downregulation of hippocampal 5‐HT1A receptors, potentially through the actions of glucocorticoids in suppressing gene transcription (Wissink et al., 2000). In addition, glucocorticoids may affect 5‐HT1A receptor sensitivity, as loss of glucocorticoids by adrenalectomy resulted in a leftward shift of the 5‐HT1A receptor concentration– response curve in the CA3 region of the hippocampus (Okuhara and Beck, 1998). The 5‐HT1A receptor is also thought to be involved in panic responses generated by the dorsal raphe nucleus–dorsal periaqueductal gray pathway. Pobbe and Zangrossi (2005), for example, demonstrated that injection of the 5‐HT1A receptor antagonist, WAY 100635, into the dorsal raphe nucleus impaired escape in the elevated T‐maze, indicating a reduced level of panic, presumably resulting from the inhibition of tonic 5‐HT1A autoreceptor activity. 5‐HT1A receptor activity may be involved in mediating aggressive behavior. Agonists at the 5‐HT1A receptor, for example, have been shown to inhibit aggressive behavior in rodents and humans, though they also sedate, which complicates interpretation (de Boer and Koolhaas, 2005). A recent study, however, used an atypical 5‐HT1A receptor ligand, S‐15535, which acts as an agonist at the somatodendritic 5‐HT1A receptor, while acting as an antagonist, or weak partial agonist, at postsynaptic receptors (de Boer and Koolhaas, 2005). This compound exhibited antiaggressive properties, suggesting that 5‐HT1A receptor agonism at the presynaptic level is responsible for the antiaggressive effects. Furthermore, administration of both S‐15535 and alnespirone (the latter being agonist at both pre‐ and postsynaptic 5‐HT1A receptors) showed an additive effect in reducing aggression rather than a predicted attenuation of the alnespirone‐ mediated effect if the actions of S‐15535 were postsynaptic.

177

178

8

5‐Hydroxytryptamine in the central nervous system

The control of rodent circadian rhythm may also be affected by 5‐HT1A receptor activation. Weber et al. (1998) demonstrated that compounds with 5‐HT1A receptor agonist activity were able to suppress light‐ induced phase shifts in the hamster circadian rhythm. In addition, Gannon and Millan (2006) showed that S‐15535 increased hamster light‐induced phase advances. Investigation of the 5‐HT1A receptor knockout mouse also discovered the potential role of this receptor subtype in sleep, specifically with respect to the paradoxical sleep phase. The knockout strain showed signs of increased periods of paradoxical sleep, while WAY 100635 induced similar effects in the wild‐type mouse (Boutrel et al., 2002). Expression of the 5‐HT7 receptor within the brain region thought to control circadian rhythm and the suprachiasmatic nucleus (SCN); however, the overlapping pharmacologies of the 5‐HT1A and 5‐HT7 receptors (e.g., activation by 8‐OH‐DPAT) has complicated this area of research. Studies with the 5‐HT1A receptor knockout mouse have led to suggestions that this subtype may take part in the processes of memory and learning. Sarnyai et al. (2000) investigated the performance of knockout mice in the Morris water maze and Y‐maze tests, and found knockout mice had lower levels of performance compared with wild‐type subjects, suggesting that the 5‐HT1A receptor has a positive effect on hippocampal‐dependent learning and memory. In contrast, studies with the 5‐HT1A receptor agonist 8‐OH‐DPAT suggest that activation of the receptor impairs working and spatial memory function. For example, systemic administration of 8‐OH‐DPAT impaired performance of rats in the radial maze, an effect blocked by the 5‐HT1A receptor antagonist, WAY 100635 (Helsley et al., 1998; Egashira et al., 2006). Egashira and colleagues further suggest that this effect of 8‐OH‐DPAT is mediated by postsynaptic 5‐HT1A receptors in the dorsal hippocampus as the effects of systemic administration were mimicked by local injection into the dorsal hippocampus, but not into other regions. It is possible, however, that 8‐OH‐ DPAT induces a biphasic response, as high doses impair memory and low doses have been shown to attenuate scopolamine‐ and tetrahydrocannabinol‐induced memory impairment, potentially through enhancing acetylcholine release in the dorsal hippocampus (Inui et al., 2004). Further evidence for 5‐HT1A receptor activation impairing cognitive processes has been provided in human studies, where administration of tandospirone, a 5‐HT1A receptor agonist, inhibited verbal memory (Yasuno et al., 2003), and psilocybin, a 5‐HT1A/2A receptor agonist, reduced performance in attentional tracking, but not in spatial working memory, in the presence of the 5‐HT2A receptor antagonist ketanserin (Carter et al., 2005). One recent development in our understanding of the role of the 5‐HT1A receptor in neurological disorders is the possible involvement of this subtype in Alzheimer’s disease. A recent positron emission tomography (PET) study of Alzheimer’s patients demonstrated reduced 5‐HT1A receptor‐binding sites in both the hippocampus and raphe nuclei, having accounted for a loss in neuronal volume, which was correlated with the severity of the observed clinical symptoms (Kepe et al., 2006). If future studies confirm the 5‐HT1A receptor is involved, then ligands at this receptor subtype may have therapeutic benefit in treating Alzheimer’s disease.

5.1.2 The 5‐HT1B Receptor The 5‐HT1B receptor‐binding site was initially distinguished from the 5‐HT1A receptor due to its low affinity for 8‐OH‐DPAT (Middlemiss and Fozard, 1983) and the rat receptor sequence was identified in 1991 by Voigt et al. (Voigt et al., 1991). During the cloning of the numerous 5‐HT1 receptor genes, there was some confusion as to whether newly identified rodent and human sequences simply represented species differences of the same subtype, or if they encoded distinct receptor subtypes. For instance, initially the human 5‐HT1B receptor sequence was classified as 5‐HT1Db, though it has now been reclassified (Hartig et al., 1996). The distribution of the 5‐HT1B receptor in the brain has been extensively characterized through receptor autoradiography (Pazos and Palacios, 1985) and immunohistochemistry (Sari et al., 1997, 1999), whereas the location of 5‐HT1B receptor mRNA has been determined by in situ hybridization (Boschert et al., 1994; Varnas et al., 2005). Given the similarity in pharmacological profile with the 5‐HT1D receptor, however, early studies reported that the detection of the 5‐HT1B receptor‐binding site may have been unable to distinguish between the two subtypes (see later). The 5‐HT1B receptor appears to

5‐Hydroxytryptamine in the central nervous system

8

be expressed at highest levels in the basal ganglia, particularly the globus pallidus and substantia nigra, with lower levels being found in the periaqueductal gray, superficial layer of the superior colliculus, cortex, amygdala, hypothalamus, hippocampus, cerebellum, and dorsal horn of the spinal cord (Bonaventure et al., 1997; Sari et al., 1999; Varnas et al., 2001). A more detailed study of the ultrastructural receptor location suggests a predominant location on axons and axon terminals, though not on synaptic membranes, as has been demonstrated within the substantia nigra and globus pallidus (Sari et al., 1997, 1999). Correspondingly, the distribution of receptor transcripts does not completely match the location of the receptor‐ binding sites, as 5‐HT1B mRNA has been identified in the raphe nuclei, striatum, hippocampus, cortex, and thalamus (Varnas et al., 2005). Transcripts are notably absent from the substantia nigra and globus pallidus, which display the strongest levels of binding sites. The 5‐HT1B receptor is thought to act as an auto‐ and heteroreceptor on 5‐HT and non‐5‐HT neurons, respectively. 5‐HT1B receptor activation has been shown to mediate inhibition of 5‐HT release in the forebrain, including the frontal cortex and hippocampus (Sleight et al., 1989; Bosker et al., 1995; Trillat et al., 1997). In addition, studies with the 5‐HT1B knockout mouse detected lower levels of 5‐HT within the nucleus accumbens and locus coeruleus (Ase et al., 2000). With respect to an impact on other neurotransmitter systems, the 5‐HT1B receptor inhibits acetylcholine release in the hippocampus from septal afferents (Cassel et al., 1995), suppresses glutamatergic transmission in the subiculum (Sari, 2004), and inhibits GABA release in the substantia nigra (Stanford and Lacey, 1996). The 5‐HT1B receptor knockout mouse also showed reduced basal dopamine levels in the nucleus accumbens, indicative of increased dopamine turnover (Ase et al., 2000). The 5‐HT1B receptor has been suggested to play a role in many physiological and pathophysiological processes. Firstly, this receptor subtype may contribute to the generation of anxious states; selective 5‐HT1B receptor agonists have been demonstrated to be anxiogenic (Lin and Parsons, 2002), potentially acting through receptors in the hippocampus and amygdala. In support of these findings, it has been shown that overexpression of the 5‐HT1B receptor in the dorsal raphe nucleus elevates stress‐induced anxiety (Clark et al., 2002). The 5‐HT1B receptor may also have a role in the generation or treatment of depression, illustrated by potential interactions between antidepressant treatment and 5‐HT1B receptor function. Chronic treatment with SSRIs, for example, downregulates or desensitizes 5‐HT1B receptors in the SCN (O’Connor and Kruk, 1994), and reversibly reduces levels of 5‐HT1B receptor transcripts in the dorsal raphe nucleus (Anthony et al., 2000), while receptor antagonism augments SSRI‐induced increases in frontal cortex 5‐HT levels (Gobert et al., 1997). In addition, a behavioral study has demonstrated that 5‐HT1B receptor activation reduces immobility in the mouse forced‐swim test, most likely via heteroreceptors, which forward a therapeutic strategy for the treatment of depression (Tatarczynska et al., 2005). Further supporting evidence comes from a recent study by Svenningsson et al. (2006). Using the yeast‐two‐hybrid system, they identified a molecular interaction between the 5‐HT1B receptor and p11, an S100 EF‐hand protein. Interestingly, p11 expression is reduced in both animal models of depression and in the human brains of depressed patients, whereas expression is upregulated on pharmacological (e.g., imipramine) or electroconvulsant therapy (> Figure 8-2). Furthermore, p11 knockout mice display symptoms of depression. It appears that p11 has a role in targeting the 5‐HT1B receptor to the cell surface, as well as facilitating receptor transduction. It is possible, therefore, that loss of 5‐HT1B receptor function through defective p11 expression may contribute to the pathogenesis of depression. Like the 5‐HT1A receptor, it has been suggested that the 5‐HT1B receptor is involved in regulating aggressive behavior. The 5‐HT1B receptor knockout mouse shows signs of increased aggression in resident– intruder paradigms (Saudou et al., 1994). Likewise, 5‐HT1B agonists, including CP 93129 and CGS 12066B, appear to reduce levels of aggression (de Boer and Koolhaas, 2005). It is unclear, however, whether these effects are mediated by presynaptic auto‐ or heteroreceptors (de Boer and Koolhaas, 2005), although the anterior hypothalamus has been forwarded as a putative location for serotonergic modulation of aggressive behavior (Ferris et al., 1997). The putative ability of the 5‐HT1B receptor to modulate responses to addictive drugs has also great therapeutic potential. For example, 5‐HT1B receptor activation reduces the self‐administration of alcohol in rats (Tomkins and O’Neill, 2000) and 5‐HT1B receptor knockout mice consume more alcohol (Crabbe et al., 1996). This 5‐HT1B receptor activity may be mediated by its regulation of dopaminergic neurotransmission,

179

180

8

5‐Hydroxytryptamine in the central nervous system

. Figure 8-2 Regulation of p11 expression by antidepressant treatments and in depression‐like states. In situ hybridization illustrating an upregulation of p11 mRNA in the forebrain following (a) repeated treatment with imipramine [10 mg/kg per day, intraperitoneally (i.p.) for 14 days] in mice (n ¼ 8 per group) and (b) electroconvulsive therapy (ECT) for 10 days in rats (n ¼ 5 per group). Conversely, p11 mRNA was downregulated in (c) the forebrain in helpless H/Rouen versus nonhelpless NH/Rouen mice (n ¼ 10 per group), and (d) in patients who suffered from unipolar major depression (n ¼ 15 per group). Data from the anterior (a; b; c, left; d) and posterior (c, right) cingulated cortices were normalized to the corresponding controls and represent means  SEM. *p < 0.05, ***p < 0.001 versus control by student’s t test (This figure is reproduced with permission from the authors (Svenningsson et al., 2006))

as demonstrated by Yan et al. (2005) by showing that ethanol‐induced increases in dopamine neuron activity in the VTA were suppressed by the selective 5‐HT1B receptor antagonist, SB 216641, and enhanced by the 5‐HT1B receptor agonist, CP 94253. Evidence has also been presented for the involvement of the 5‐ HT1B receptor in the reinforcing properties of cocaine. Studies using the 5‐HT1B receptor knockout mouse demonstrated elevated dopamine levels in the nucleus accumbens, both basally and following cocaine administration (Shippenberg et al., 2000).

5‐Hydroxytryptamine in the central nervous system

8

Modulation of central dopamine function may also be relevant for the 5‐HT1B receptor’s role in locomotor activity demonstrated by the ability of the 5‐HT1B receptor agonist, RU 24969, to increase spontaneous movement and the increased exploratory behavior in 5‐HT1B receptor knockout mice (Ramboz et al., 1996; Brunner et al., 1999). The circadian rhythm in rodents may also be modulated by the actions of the 5‐HT1B receptor. The 5‐HT1B receptor has been identified within the SCN of the hypothalamus, a key region involved in controlling circadian rhythm. In 2000, Garabette et al. (Garabette et al., 2000) demonstrated that local administration of the 5‐HT1B receptor agonist, RU 24969 attenuated 5‐HT release in the SCN. In addition, 5‐HT1B receptor agonists block glutamatergic excitatory postsynaptic currents (EPSCs) within this region (Smith et al., 2001), and reduce the frequency of inhibitory postsynaptic currents (IPSCs) in GABAergic SCN neurons (Bramley et al., 2005). Study of the 5‐HT1B receptor knockout mouse identified an inability of this strain to entrain to unnatural photic stimuli (i.e., those not based on a 24‐h cycle), suggesting that the absence of this receptor subtype affects the ability of the circadian rhythm to respond to changes in light (Sollars et al., 2006). At present, perhaps the most important therapeutic application of 5‐HT1B receptor ligands involves the treatment of migraine, which is both a neural and vascular disorder. The receptor inhibits trigeminal sensory nerve activity with a subsequent reduction in relevant neuropeptide release. In addition, the 5‐HT1B receptor is expressed by the endothelium of meningeal vessels, which undergo an inflammatory response during migraine, and hence regulate the contractile activity of these vessels. The strategic development of 5‐HT1B receptor agonists (e.g., sumatriptan, zolmitriptan, and naratriptan) as antimigraine agents has been vindicated and made an important impact on the treatment of migraine. Unlike other 5‐HT receptors, the 5‐HT1B receptor appears to be modulated by an endogenous tetrapeptide (Leu–Ser–Ala–Leu), named 5‐HT moduline. This peptide, first isolated from rat and bovine brain in 1996 (Rousselle et al., 1996), appears to inhibit 5‐HT binding to the receptor through an allosteric site distinct from the ligand‐binding domain, and can be released, Ca2þ/Kþ dependently, from rat cortical synaptosomes (Massot et al., 1996). Furthermore, there appears to be a functional consequence to this interaction since 5‐HT moduline has a 5‐HT1B receptor antagonist‐like effect in the mouse social‐ interaction test and the actions of the 5‐HT1B receptor agonist, RU 24969, are inhibited (Massot et al., 1996). In addition, presumed block of 5‐HT moduline action with an antibody reduced measures of anxiety in the open‐field test (Grimaldi et al., 1999). The receptor interaction of this endogenous peptide may, therefore, offer a therapeutic strategy for the treatment of anxiety, depression, and aggressive behavior.

5.1.3 The 5‐HT1D Receptor The human 5‐HT1D receptor (formally known as the 5‐HT1Da receptor) was initially cloned in 1991 by Hamblin and Metcalf (Hamblin and Metcalf, 1991). The 5‐HT1D receptor is relatively less well understood in comparison with the 5‐HT1A and 5‐HT1B receptors, primarily due to the difficulties in separating 5‐HT1B and 5‐HT1D receptors pharmacologically. Consequently, selective identification of 5‐HT1D receptor‐binding sites has been difficult. Receptor autoradiography using, for instance, [125I]GTI with CP 93129 to mask 5‐HT1B receptor sites, identified 5‐HT1D‐like binding sites in the rat basal ganglia (globus pallidus, substantia nigra, and caudate putamen), and the hippocampus and cortex (Bruinvels et al., 1993). Studies with human brain tissue exploited the 15–30‐fold greater selectivity of ketanserin for the 5‐HT1D receptor and determined ketanserin‐sensitive [3H]sumatriptan‐binding sites (Castro et al., 1997), where the distribution of 5‐HT1D‐like sites was comparable with rat brain. In situ hybridization studies in rat and primate brain localized transcripts to the dorsal raphe nucleus, locus coeruleus, nucleus accumbens, olfactory cortex, and caudate putamen (Bruinvels et al., 1994). Similar to the 5‐HT1B receptor, mRNA was not identified in the globus pallidus and substantia nigra, suggesting that the receptor protein may be found on axon terminals projecting from other regions (caudate putamen?). The 5‐HT1D receptor inhibits adenylate activity in recombinant cell systems (Hamblin and Metcalf, 1991). With respect to physiological functions, the nature of receptor location suggests that it could act as an auto‐ or heteroreceptor. It is generally accepted, however, that at least the principle terminal 5‐HT

181

182

8

5‐Hydroxytryptamine in the central nervous system

autoreceptor, in terms of function, is the 5‐HT1B receptor due to sensitivity to the relatively selective 5‐HT1B receptor antagonist, SB 216641, but not the 5‐HT1D receptor antagonist, BRL 15572 (Schlicker et al., 1997). However, some evidence for a heteroreceptor function is available. For instance, Maura et al. (1998) demonstrated a proposed 5‐HT1D receptor‐mediated inhibition of glutamate release in the human cortex. The performance of this study, however, predated the availability of selective ligands and hence a definitive central role for the 5‐HT1D receptor remains to be determined. This 5‐HT receptor subtype may play a role in the treatment of migraine; however, the efficacious antimigraine ‘‘triptans’’ are 5‐HT1D receptor agonists (as well as 5‐HT1B receptor agonists). To further support this premise, the 5‐HT1D receptor is expressed in the trigeminal ganglia (Hou et al., 2001). The recent discovery of a highly selective 5‐HT1D receptor antagonist, SB 714786, should aid the investigation of the physiological functions of the 5‐HT1D receptor (Ward et al., 2005).

5.1.4 The 5‐ht1E Receptor The 5‐ht1E receptor remains the only member of the 5‐HT1 receptor family that retains the lower class appellation of a protein with unidentified function. The human gene was isolated in 1992 (McAllister et al., 1992), but there was a subsequent lack of rodent 5‐ht1E cloned sequences. Recently, however, the guinea pig 5‐ht1E receptor gene was identified, whereas the mouse genome has been found to contain no 5‐ht1E receptor sequence (Bai et al., 2004). 5‐ht1E receptor distribution within the brain has been documented, with transcripts found in the cortex, caudate putamen, and amygdala of human tissue (Bruinvels et al., 1994). Attempts to map the receptor‐binding sites have exploited the low affinity of the 5‐ht1E receptor for 5‐CT to distinguish it from most other 5‐HT1 receptors (Miller and Teitler, 1992). Putative 5‐ht1E receptor‐binding sites were found in the cerebral cortex, caudate putamen, and claustrum, with lower levels of binding in the hippocampus and amygdala. The apparent colocalization of mRNA and receptor suggests a postsynaptic distribution for this receptor subtype. Similar to other members of the 5‐HT1 family, within recombinant cell systems the 5‐ht1E receptor couples negatively to adenylate cyclase (Levy et al., 1992). Study of this potential receptor is hindered by the lack of selective ligands, although structure–affinity relationship (SAR) studies have been reported (e.g., Dukat et al., 2004), which demonstrate a similar pharmacology to the 5‐HT1F receptor, although a few notable distinctions are apparent (see later). Hence, such studies indicate that selective ligands for the 5‐ht1E receptor may be identifiable.

5.1.5 The 5‐HT1F Receptor The 5‐HT1F receptor was initially identified in the mouse genome, but was named 5‐ht1Eb due to its similar pharmacological to the 5‐ht1E receptor (Amlaiky et al., 1992), though localization of 5‐HT1F receptor transcripts clearly showed a distinct pattern of expression when compared with the 5‐ht1E receptor. Receptor autoradiography with [3H]sumatriptan in the presence of 5‐CT to block 5‐HT1B/1D receptor‐binding sites demonstrates that the 5‐HT1F receptor is located within the guinea pig hippocampus, cortex, claustrum, and caudate nucleus (Waeber and Moskowitz, 1995a, b). More recently, the selective ligand [3H]LY 334370 confirmed largely the distribution of the 5‐HT1F receptor in rodent brain (Lucaites et al., 2005). Despite the identification of two 5‐HT1F receptor selective ligands, LY 344864 and LY 334370, little information has been published concerning the physiological function(s) of this receptor. One potential role is an involvement in the treatment of migraine; forwarded initially due to the relatively high affinity of sumatriptan and naratriptan. In support of this hypothesis, the 5‐HT1F receptor has been localized on glutamatergic neurons within the trigeminal ganglia (Ma, 2001); activation of these neurons are thought to induce dural protein extravasation, a potential contributor to the generation of migraine, and the 5‐HT1F receptor agonist, LY 344864, inhibits this process (Phebus et al., 1997).

5‐Hydroxytryptamine in the central nervous system

8

5.2 The 5‐HT2 Receptors The 5‐HT2 receptor family consists of three receptor subtypes: 5‐HT2A, 5‐HT2B, and 5‐HT2C receptors (the latter was previously known as 5‐HT1C receptor, but was appointed to the 5‐HT2 receptor family on the basis of its structure, mode of signal transduction, and to some extent pharmacology; Hoyer et al., 1994). Until fairly recently, it has been difficult to distinguish the three subtypes pharmacologically, but now selective antagonists including MDL 100907 (5‐HT2A), SB 204741 (5‐HT2B), and SB 242084 (5‐HT2C) have become available to allow responses to be assigned to individual receptors with more confidence.

5.2.1 5‐HT2A Receptor The 5‐HT2A receptor was initially identified as a binding site in rat cortical membranes (Peroutka and Snyder, 1979), with subsequent identification of the rat sequence a decade later (Pritchett et al., 1988; Julius et al., 1990). The distribution of the 5‐HT2A receptor in the brain has been well characterized. Receptor autoradiography with selective ligands, such as [3H]‐MDL 100907, has shown high levels of expression in human and rodent forebrain, including the neocortex, entorhinal and piriform cortices, hippocampus, caudate nucleus, nucleus accumbens, and olfactory tubercle (Lopez‐Gimenez et al., 1997). The location of 5‐HT2A mRNA corresponds well to receptor distribution (Burnet et al., 1995), generally following the distribution of 5‐HT neuron innervation, implying that the receptor has a postsynaptic location. The cellular expression of the 5‐HT2A receptor protein appears to be predominantly neuronal, both on GABAergic interneurons in the cortex and glutamatergic pyramidal cells within the cortex and hippocampus (Pompeiano et al., 1994; Burnet et al. 1995; Willins et al., 1997; Jakab and Goldman‐Rakic, 1998). A detailed study of the subcellular location of 5‐HT2A receptors in rat PFC (Miner et al., 2003) observed expression on both the shafts and spines of proximal and distal pyramidal dendrites, reinforcing the likely postsynaptic location of the receptor. In addition to the 5‐HT2A receptor being expressed on the plasma membrane (Willins et al., 1997), it also exhibits a degree of intracellular localization, which may be indicative of a high rate of receptor turnover (Martin‐Ruiz et al., 2001; Cornea‐Hebert et al., 2002). The precise ultrastructural positioning of the receptor is supported by the work of Cornea‐Hebert et al. (2002), who demonstrated that the 5‐HT2A receptor physically interacts with the cytoskeletal protein, MAP1A, suggesting that the receptor may regulate neuronal development or dendritic plasticity. The 5‐HT2A receptor is coupled to the activation of phospholipase C (PLC), inducing the mobilization of intracellular Ca2þ stores, in both recombinant systems and native tissue (Conn and Sanders‐Bush, 1984; Pritchett et al., 1988). Additionally, the 5‐HT2A receptor may activate second‐messenger cascades responsible for the regulation of brain‐derived neurotrophic factor (BDNF) levels, as 5‐HT2A receptor agonists reduce levels of BDNF in the dentate gyrus of the hippocampus, while increasing levels in the neocortex, which has potentially profound effects on neuronal growth (Vaidya et al., 1997). 5‐HT2A receptor activation is also thought to mediate, at least in part, the 5‐HT‐induced attenuation of the slow after hyperpolarizing current observed in layer V pyramidal neurons in the cortex following a burst of spikes (Villalobos et al., 2005). The 5‐HT2A receptor regulates the release of many neurotransmitters, including glutamate, dopamine, and GABA. Within the forebrain, for example, the 5‐HT2A receptor increases both glutamate release from layer V pyramidal neurons in the PFC (see Aghajanian and Marek, 1999b, for review) and GABA release onto CA1 pyramidal neurons in the hippocampus (Shen and Andrade, 1998). The 5‐HT2A receptor also appears to have a regulatory effect on dopaminergic neuron firing, supported by receptor expression being associated with dopaminergic neurons within the VTA and substantia nigra (Ikemoto et al., 2000). Furthermore, 5‐HT2A receptor antagonism attenuates dopamine release in the VTA (De Deurwaerdere and Spampinato, 1999) and striatum (Lucas and Spampinato, 2000). It has been suggested that the 5‐HT2A may be involved in the pathogenesis of depression and mediate some of the effects of antidepressant treatment. Correspondingly, the selective knockout of 5‐HT2A receptor expression through injection of antisense oligonucleotides evokes antidepressant‐like effects in the mouse forced‐swim test (Sibille et al., 1997), and also reduced anxiety in the elevated plus maze paradigm

183

184

8

5‐Hydroxytryptamine in the central nervous system

(Cohen, 2005). Indeed, it has been suggested that levels of 5‐HT2A receptor expression are altered in the ‘‘depressed’’ brain. One PET study, for example, identified reductions in 5‐HT2A receptor levels within the frontal, occipital, temporal, and cingulate cortices of drug‐naı¨ve depressed patients relative to control subjects (Messa et al., 2003). The evaluation of drug‐naı¨ve depressed patients here is likely to be important. For instance, the many studies assessing 5‐HT2A receptor levels in human postmortem tissue from depressed or suicide victims have generally reported conflicting results, with some papers finding elevated 5‐HT2A receptor levels (e.g., Pandey et al., 2002), whereas others are unable to identify any differences. The complication of therapy impacting on 5‐HT2A receptor expression is difficult to control in these postmortem studies. Relevant to potential alterations in the expression of the 5‐HT2A receptor, studies of the physiological actions of the 5‐HT2A receptor have postulated a therapeutic role in the treatment of depression, as 5‐HT2A receptor activation has a negative feedback on raphe neuron firing (Boothman et al., 2003). It has, therefore, been suggested that antagonism of the 5‐HT2A receptor might accelerate the rate of onset of antidepressant drugs, similar to 5‐HT1A receptor antagonism. Consequently, Boothman et al. (2006) reported an elevation in hippocampal 5‐HT release following administration of the SSRI citalopram in combination with a 5‐HT2A receptor antagonist. The authors postulated that this effect could be due to either a block of terminal 5‐HT2A receptor‐mediated negative feedback to the raphe neurons, or through inhibiting GABAergic transmission by interneurons expressing the 5‐HT2A receptor in the raphe nucleus itself. The 5‐HT2A receptor may also contribute to depression through modification of BDNF levels, which increase as a consequence of antidepressant treatment (Nibuya et al., 1995). Furthermore, expression of BDNF and the 5‐HT2A receptor appear to be linked, as a mutant mouse with low levels of postnatal BDNF expression displayed a significant deficit in 5‐HT2A receptor expression in both the dorsal raphe nucleus and PFC, and exhibited a corresponding reduction in the 5‐HT2A receptor‐mediated postsynaptic response in the mPFC (Rios et al., 2006; > Figure 8-3). The 5‐HT2A receptor is also thought to play an integral role in the treatment of schizophrenia, in addition to mediating the effects of hallucinogenic drugs, both of which may have similar underlying mechanisms (Vollenweider and Geyer, 2001). The most compelling evidence that the 5‐HT2A receptor is involved in the pathogenesis and/or the treatment of schizophrenia has come from the study of atypical antipsychotic pharmacology, as most, but not all, of these compounds have a high affinity for the 5‐HT2A receptor (for review, see Meltzer et al., 2003). The most likely region to generate both of these actions is the . Figure 8-3 5‐HT2A‐mediated postsynaptic responses in the prefrontal cortex. (a) Cumulative probability curves showing that serotonin elicits a significant increase in sEPSC frequency in neurons from wild‐type mice (K‐S test; p < 0.001) but not BDNF‐mutant (p ¼ NS) mice. (b) Bar graph summarizes data (mean  SE) for WT and BDNF‐mutant mice. The data are based on recordings from wild‐type cells (n ¼ 5; two mice) and cells from BDNF2L/2LCamKCse93‐mutant mice (n ¼ 9; four mice); t test, p < 0.001 (This figure is reproduced with permission from the authors (Rios et al., 2006))

5‐Hydroxytryptamine in the central nervous system

8

cortex, where glutamatergic pyramidal neurons have been shown to express the 5‐HT2A receptor, and to be regulated by its activity (Aghajanian and Marek, 1999a, b). Furthermore, some studies have shown a reduction in both 5‐HT2A receptor transcripts and binding sites in the cortex and parahippocampal gyrus of schizophrenic patients (Burnet et al., 1996), although a PET study appears to contradict these findings (Okubo et al., 2000). Learning and memory may be affected by 5‐HT2A receptor activation, given the high levels of receptor expression within the hippocampus and cortex, both key regions in mnemonic function. Of more direct evidence for instance, 5‐HT2A receptor agonists have been shown to improve rabbit eyeblink response, a measure of associative learning, though the compounds used are also 5‐HT2C receptor agonists (Welsh et al., 1998; see Harvey, 2003, for review). A role for the 5‐HT2A receptor in working memory has also been postulated (Williams et al., 2002), due to reduced performances of primates in delayed response tasks following administration of selective 5‐HT2A receptor antagonists, while agonists induced a modest improvement in performance. Interestingly, working memory is thought to be deficient in schizophrenia, further implying that this receptor subtype is involved in the pathogenesis of this disorder. Secretion of hormones from the hypothalamus, including ACTH, corticosterone, oxytocin, prolactin, and rennin, has been found to be under the control of 5‐HT, probably via the 5‐HT2A receptor, since the response is mimicked by the 5‐HT2A/2C receptor agonist DOI and antagonized by the selective 5‐HT2A receptor antagonist, MDL 100907 (Van de Kar et al., 2001). Anorexia and bulimia nervosa are psychological conditions thought to involve the 5‐HT system, due to the occurrence of abnormalities in mood, feeding, and impulsiveness, in addition to the ability of drugs acting on the 5‐HT system being only the compounds to date displaying efficacy to treat these conditions. Little other direct evidence has been found, although Audenaert et al. (2003) have shown an apparent reduction in 5‐HT2A receptor binding in the frontal, parietal, and occipital cortices of patients with eating disorders using SPECT brain imaging and the selective ligand, [123I]‐5‐I‐R91150. The 5‐HT2A receptor may also be involved in the regulation of a particular phase of sleep, termed nonrapid eye movement sleep (NREMS). The relatively nonselective 5‐HT2 receptor antagonist, ritanserin, increases NREMS in human subjects (Idzikowski et al., 1991), while blocking the 5‐HT2A receptor in mice has the same effect and this response was not evident in 5‐HT2A receptor knockout strains (Popa et al., 2005). Further investigation of the role of the 5‐HT2A receptor in sleep is clearly required.

5.2.2 The 5‐HT2B Receptor The 5‐HT2B receptor (previously known as the 5‐HT2F receptor) was first identified through its mediation of rat stomach fundus contraction, from where its sequence was subsequently cloned (Foguet et al., 1992). The 5‐HT2B receptor, in comparison with other members of the 5‐HT2 receptor family, has a limited distribution within the brain, with 5‐HT2B receptor‐like immunoreactivity localized on neuronal cells within the cerebellum, lateral septum, dorsal hypothalamus, and medial amygdala (Duxon et al., 1997a). The putative functions of this receptor may include a role in anxiety‐like behaviors, as the marginally selective 5‐HT2B receptor agonist, BW 723C86, is anxiolytic following direct intra‐amygdala injection (Duxon et al., 1997b). In addition, this agonist increased the number of punishments received in the rat Vogel drinking conflict test, an effect blocked by the selective 5‐HT2B receptor antagonist, SB 215505 (Kennett et al., 1998). These studies support a role for the 5‐HT2B receptor in anxiety and this subtype may therefore be a therapeutic target for the treatment of depression and anxiety‐like disorders. Certain phases of sleep may also be regulated by the actions of the 5‐HT2B receptor. For example, administration of the 5‐HT2B receptor antagonist, SB 215505, at the beginning of the light phase, increases the waking period, with a corresponding loss in paradoxical sleep (Kantor et al., 2004). One study has demonstrated that activation of the 5‐HT2B receptor induces hyperphagia (Kennett et al., 1997a), implicating the 5‐HT2B receptor in the regulation of feeding, despite most attention being focused on the role of the 5‐HT2C receptor in feeding (see later). The 5‐HT2B receptor is 5‐HT receptor that may be involved in the generation of migraines. The 5‐HT2B/ 2C agonist, m‐CPP, is known to trigger migraine‐like pain in some individuals, and has the ability to induce

185

186

8

5‐Hydroxytryptamine in the central nervous system

plasma protein extravasation in the guinea pig; an effect blocked by the selective 5‐HT2B receptor antagonist, LY 202146 (Johnson et al., 2003). Furthermore, this receptor has been shown to be expressed by endothelial cells within meningeal blood vessels, the potential site responsible for migraine generation (Schmuck et al., 1996).

5.2.3 The 5‐HT2C Receptor The 5‐HT2C receptor‐binding site was originally identified in the choroid plexus, and displayed high affinity for [3H]5‐HT (Pazos et al., 1984), resulting in the initial classification within the 5‐HT1 receptor family; high affinity for 5‐HT being a key characteristic to guide classification at the time. The 5‐HT2C receptor sequence was subsequently identified in the rat (Julius et al., 1988). In contrast to the 5‐HT2B receptor, the 5‐HT2C receptor has a widespread distribution throughout the brain. Receptor autoradiographical and immunohistochemical studies have complemented each other, identifying putative sites of receptor expression in the choroid plexus, cortex, amygdala, hippocampus, substantia nigra, caudate nucleus, and cerebellum (e.g., Abramowski et al., 1995). Generally, in situ hybridization has colocalized 5‐HT2C receptor transcripts with the binding sites, suggesting that the receptor is postsynaptic, with the exception of a potentially presynaptic receptor localization in the medial habenula. Activation of the 5‐HT2C receptor is thought to induce membrane depolarization, and may mediate some of the excitatory effects of 5‐HT, for instance in piriform cortical pyramidal neurons (Sheldon and Aghajanian, 1991) and neurons nigral neurons (Rick et al., 1995). More recently, attention has been focused on the role of the 5‐HT2C receptor in regulating the firing activity of dopaminergic neurons, particularly within the VTA and substantia nigra. On the whole, evidence suggests that the 5‐HT2C receptor has a tonic inhibitory control on the firing activity of mesolimbic and mesostriatal dopaminergic neurons. For instance, acute administration of the 5‐HT2B/2C receptor antagonist, SB 206553, increased the rate of firing of neurons in the VTA and substantia nigra, resulting in elevated dopamine release in the nucleus accumbens and striatum (Di Giovanni et al., 1999; Alex et al., 2005). The role of the receptor to modulate nigral–striatal dopamine neurons is controversial, however, since the selective 5‐HT2C receptor antagonist, SB 242084, while increasing firing in the VTA, did not alter nigral neuron activity, and the 5‐HT2C receptor agonist, Ro 60‐0175, decreased basal firing of dopaminergic neurons in the VTA again the nigral neurons were not affected (Di Matteo et al., 1999). A major functional role of the 5‐HT2C receptor is postulated to be the control of feeding, which has incited interest in this subtype as a therapeutic target for antiobesity drugs. The first indication of such a role arose from observations that 5‐HT2C receptor knockout mice were abnormally overweight (Tecott et al., 1995). In addition, the 5‐HT2B/C agonist, mCPP, can induce hypophagia, which is prevented by the 5‐HT2C receptor antagonist, SB 242084 (Kennett et al., 1997b). The anxiolytic properties of 5‐HT2C receptor antagonists suggest a role for this receptor subtype in anxiety (Kennett et al., 1996, 1997b). A more recent report of a new antidepressant compound, agomelatine, which is an agonist of melatonin receptors MT1 and MT2, but also has 5‐HT2C antagonist activity (Den Boer et al., 2006), suggests that blockade of the 5‐HT2C receptor may also have therapeutic benefit in the treatment of depression. Some studies suggest that RNA editing of the 5‐HT2C receptor may be relevant in schizophrenic, depressed, and suicidal patients. This process is known to produce three amino acid alterations within the intracellular regions of the receptor, which may affect the efficiency of the receptor couples with G‐proteins (Burns et al., 1997), and may also influence pharmacology (Niswender et al., 2001). This latter study, however, was unable to detect changes in RNA editing in schizophrenic or depressed patients, though they did identify increased editing at one site in suicide victims. Interestingly, RNA editing appears to be regulated by 5‐HT levels in the brain; perhaps logically depletion attenuates editing, resulting in higher levels of isoforms with enhanced G‐protein‐coupling efficacy (Gurevich et al., 2002). The activity of the 5‐HT2C receptor may, therefore, be altered in diseased states where 5‐HT levels are abnormal, and may lead to further disruption of normal brain functions.

5‐Hydroxytryptamine in the central nervous system

8

Similar to the other 5‐HT2 receptor subtypes, the 5‐HT2C receptor may be involved in the regulation of sleep. Antagonists for this receptor promote slow‐wave sleep (Sharpley et al., 1994), though in contrast a more recent study demonstrated that the 5‐HT2C receptor antagonist, SB 242084, increased wakefulness at the expense of deep slow‐wave sleep (Kantor et al., 2005). Clarification of the role of the 5‐HT2C receptor in this respect is clearly required. Due to expression within the hippocampus, the 5‐HT2C receptor has also been postulated to have a role in memory and learning. Of relevance here, a significant impairment in the generation of perforant path‐dentate gyrus LTP, a process considered to be the most likely candidate for the molecular basis of memory, was observed in 5‐HT2C receptor knockout mice (Tecott et al., 1998). In addition, these mice showed impaired spatial learning in the Morris water maze. Finally, activation of the 5‐HT2C receptor may regulate locomotor activity, as the selective agonist, WAY 161503, suppresses measures of spontaneous movement (Mosher et al., 2005).

5.3 The 5‐HT3 Receptor The 5‐HT3 receptor is the only 5‐HT receptor that is a member of the cys–cys loop ligand‐gated ion channel family. The channel is thought to be pentameric (Boess et al., 1995), and may be formed by a combination of up to five different subunits, named 5‐HT3A–E, although at present only the 5‐HT3A and 5‐HT3B subunits have been demonstrated to be incorporated into functional channels. The receptor complex is a nonselective ion channel, permeable to Ca2þ, Naþ, and Kþ ions, that mediates fast synaptic neurotransmission in the brain, via membrane depolarization and is prone to rapid desensitization. Recent attention has focused on the combination of subunits forming the functional channel in native tissue, with current opinion favoring a combination of 5‐HT3A and 5‐HT3B subunits. Expression of only the 5‐HT3A receptor subunit in recombinant systems does not generate a high single‐channel conductance receptor that is evident in some populations of native neuronal receptors, whereas coexpression with the 5‐HT3B receptor modifies the biophysics of the receptor to more resemble some populations of native receptors (Davies et al., 1999; Dubin et al., 1999; > Figure 8-4). In addition to 5‐HT, 5‐HT3 receptor action is modulated, allosterically, by volatile anaesthetics and alcohols (Machu and Harris, 1994; Parker et al., 1996; Suzuki et al., 2002), though the actions of these compounds may depend on the subunit composition of the receptor (Stevens et al., 2005). Within the CNS, the 5‐HT3 receptor is expressed at highest density in the brain stem nuclei, encompassing the chemoreceptor trigger zone; dorsal motor nucleus of the vagus nerve, area postrema, and nucleus tractus solitarius (Pratt et al., 1990). The 5‐HT3 receptor is also found, albeit at much lower levels, in human forebrain regions including the hippocampus, amygdala, and caudate–putamen (Barnes et al., 1989a, b). Interestingly, this latter region in other species (rodents, nonhuman primates) does not display readily detectable expression of 5‐HT3 receptor‐binding sites. A comprehensive understanding of the differential distribution of the individual receptor subunits has not yet been reached, but may be achieved in future studies by the use of selective antibodies recognizing distinct epitopes within the individual subunits. Activation of the 5‐HT3 receptor is believed to modulate the transmission of various neurotransmitters. For example, in the hippocampus, frontal cortex, and hypothalamus, activation of the 5‐HT3 receptor enhances 5‐HT release (e.g., Martin et al., 1992), although the receptor is not thought to be expressed by 5‐HT neurons. In addition, this receptor is thought to have a facilitatory effect on dopamine release. For instance, electrical stimulation of dorsal raphe neurons increases dopamine release in the nucleus accumbens, an effect inhibited by the 5‐HT3 receptor antagonists, ondansetron and zacopride (De Deurwaerdere et al., 1998). Conversely, the 5‐HT3 receptor has an inhibitory effect on acetylcholine release in the cortex (Barnes et al., 1989b) that is likely to be mediated via GABAergic interneurons (Morales and Bloom, 1997; Diez‐Ariza et al., 2002). The primary therapeutic benefit of 5‐HT3 receptor ligands (e.g., ondansetron, granisetron, tropisetron) is their ability to relieve often very severe nausea and vomiting induced by aggressive anticancer chemo‐ and

187

188

8

5‐Hydroxytryptamine in the central nervous system

. Figure 8-4 The pharmacological and biophysical properties of homomeric and heteromeric 5‐HT3 receptors. (a) Concentration‐dependent activation of currents by 5‐HT recorded from HEK‐293 cells transfected with 5‐HT3A cDNA alone (filled circles) or in combination with 5‐HT3B cDNA (open circles). Data points represent mean current amplitudes recorded from at least four cells normalized to the maximum current amplitude. (b) Concentration‐dependent inhibition by metoclopramide (squares) and tubocurarine (circles) of currents mediated by 5‐HT3A (filled symbols) and heteromeric (open symbols) receptors. (c) Current–voltage relationships for responses evoked by 10 mM 5‐HT, recorded from cells expressing 5‐HT3A (filled circles) and heteromeric (open circles) receptors. Data points represent mean current amplitudes recorded from at least four cells and normalized to the amplitude of the current recorded at 80 mV. Data points for heteromeric receptors were fitted with a linear function. (d) Representative low‐gain d.c. and high‐gain a.c.‐coupled records of an inward current response to 5‐HT (1 mM) recorded at a holding potential of 60 mV from an HEK‐293 cell expressing heteromeric 5‐HT3 receptors. The relationship between membrane current variance and mean current amplitude (1‐s period) was fitted by linear regression for five cells, to yield a single‐channel amplitude (i) of 0.65  0.02 pA and an elementary conductance (g) of 11.7  0.3 pS. (e) Single‐channel recordings from outside‐out patches containing 5‐HT3A (top panel) and heteromeric (lower panel) 5‐HT3 receptors. The conductance (16 pS) of channels mediated by heteromeric receptors was derived from the linear fit to the current–voltage relationship obtained from three excised patches (Data are reproduced with permission from the authors (Davies et al., 1999))

radiotherapy (e.g., Ikeda et al., 2005) and also emesis occurring postoperatively, particularly evident following procedures involving the abdomen (Du Pen et al., 1992). Less well understood is the potential efficacy of 5‐HT3 receptor antagonists to reduce anxiety and other symptoms associated with the forebrain. Despite studies showing that 5‐HT3 receptor agonists may increase anxiety levels in the mouse (Costall et al., 1989), in addition to antagonists having anxiolytic properties, and the 5‐HT3A receptor knockout mouse exhibiting reduced levels of anxiety in the elevated plus maze and

5‐Hydroxytryptamine in the central nervous system

8

social interaction tests (Kelley et al., 2003), 5‐HT3 receptor antagonists do not appear to evoke a robust anxiolytic response in patients. The 5‐HT3 receptor may also be involved in cognitive processes, such as learning and memory (Barnes et al., 1990), which may be mediated through the indirect effects of the 5‐HT3 receptor on acetylcholine release. Also of relevance here, activation of the 5‐HT3 receptor has been demonstrated to inhibit long‐term potentiation in the rat hippocampus (Passani et al., 1994); this process being a well‐recognized physiological phenomenon believed to represent memory. Consistent with these findings, 5‐HT3 receptor antagonists have been forwarded for the treatment of various neurological disorders in which memory deficit is concerned, including Alzheimer’s disease, although the clinical responses are not encouraging. Other potential therapeutic roles of 5‐HT3 receptor antagonists include the relief of alcohol addiction (Dawes et al., 2005), the treatment of tardive dyskinesia (Sirota et al., 2000), and the treatment of pain. The role of the 5‐HT3 receptor in this latter process is currently ambiguous due to conflicting reports of its activation being pro‐ or antinociceptive. For example, the selective 5‐HT3 receptor antagonist, ondansetron, inhibits the formalin‐induced response in dorsal horn neurons (Green et al., 2000), while use of the 5‐HT3 receptor agonist 2‐methyl 5‐HT reduces the behavioral response to formalin (Sasaki et al., 2001). The 5‐HT3 receptor antagonists have also been found useful in the treatment of fibromyalgia (e.g., Fa¨rber et al., 2000), a disorder characterized by chronic, widespread pain. In addition, 5‐HT3 receptor antagonists, granisetron, tropisetron, and ondansetron, have been proposed to be beneficial in treating chronic fatigue syndrome (Spa¨th et al., 2000; The et al., 2003).

5.4 The 5‐HT4 Receptor Although only one 5‐HT4 receptor gene has been identified, the arising mRNA can be alternatively spliced within the corresponding C‐terminal region to produce nine isoforms, 5‐HT4A–H and 5‐HT4HB, although it is possible that more may become apparent. Most isoform transcripts studied appear to be expressed in the brain, with the exception of 5‐HT4D, which currently has been located only in the gut. The isoforms do not appear to differ pharmacologically, though they may vary in G‐protein‐coupling efficiency (Mialet et al., 2000), perhaps not surprising given the putative role of the C terminus of GPCRs to interact with G‐protein subunits. Various studies have investigated the location of the 5‐HT4 receptor within the brain, the binding sites and mRNA colocalize, indicating a probable postsynaptic location. Receptor autoradiography using human tissue has identified 5‐HT4 receptors with highest levels in the basal ganglia, including the substantia nigra, globus pallidus, caudate nucleus, putamen, nucleus accumbens, hippocampus (CA1 and subiculum), and cortex (Varnas et al., 2003). The 5‐HT4 receptor is positively coupled to adenylate cyclase, and enhances neuronal excitability (Gerald et al., 1995). Consistent with this role, the 5‐HT4 receptor increases the release of acetylcholine in the rat frontal cortex, which is activated using the selective 5‐HT4 receptor agonists, BIMU1 and BIMU8 (Consolo et al., 1994). Although antagonists block this response, they have no effect on acetylcholine release alone, suggesting that the 5‐HT4 receptor has little tonic control of basal cholinergic transmission. The 5‐HT4 receptor has also been shown to have a facilitatory effect on striatal dopaminergic transmission (Steward et al., 1996). Likewise, in 1993, Benloucif et al. (Benloucif et al., 1993) demonstrated that the nonselective 5‐HT4 receptor agonist, 5‐MT, could increase dopamine release in the striatum. More recently, the morphine‐induced increase in striatal dopamine release was inhibited by two selective 5‐HT4 receptor antagonists, GR 125487 and SB 204070. These compounds exerted no effect in the absence of neuronal excitation, or following amphetamine‐induced dopamine release, suggesting that only this receptor subtype has a modulatory influence during dopaminergic neuron activity (Porras et al., 2002). Interestingly, the 5‐HT4 receptor also facilitates nigral dopamine release (Thorre´ et al., 1998). 5‐HT release in the hippocampus (Ge and Barnes, 1996) also appears to be enhanced by the 5‐HT4 receptor, with the receptor being tonically active in freely moving rats. It has been suggested that this receptor can regulate the firing activity of serotonergic neurons originating in the dorsal raphe nucleus (Lucas et al., 2005). Such manipulation of the central 5‐HT system may forward this receptor as a target for antidepressant therapy.

189

190

8

5‐Hydroxytryptamine in the central nervous system

The 5‐HT4 receptor is believed to have a major role in mediating the neuronal basis of learning and memory. Ligands acting on this receptor subtype, therefore, have been studied to identify any cognitive‐ enhancing properties that may prove beneficial in the treatment of conditions such as Alzheimer’s disease. Moreover, there has been a report of reduced 5‐HT4 receptor density in the postmortem brains of Alzheimer’s disease sufferers (Reynolds et al., 1995), implying that the receptor may be involved in the pathogenesis of this disorder. Many studies have shown that 5‐HT4 receptor activation enhances performance in numerous behavioral paradigms of cognitive function. For example, the 5‐HT4 receptor agonist, RS 17017, improved primate performance in the delayed matching task (Terry et al., 1998), whereas an alternative 5‐HT4 receptor agonist, RS 67333, enhanced learning in the Morris water maze (Lelong et al., 2001). The partial 5‐HT4 receptor agonist, SL65.0155, also facilitated retention during an object recognition task, which was antagonized by the 5‐HT4 receptor antagonist, SDZ 205557 (Moser et al., 2002). The same report described a synergistic effect on memory enhancement when combining SL65.0155 and the cholinesterase inhibitor, rivastigmine, further implying that use of 5‐HT4 receptor agonists may have beneficial effects in treating Alzheimer’s disease. These positive effects of 5‐HT4 receptor activation may be mediated by its regulation of acetylcholine release in the cortex. An alternative possibility is the interaction of this receptor with amyloid precursor protein (APP) metabolism. 5‐HT4 receptor agonists appear to increase the secretion of sAPPa (> Figure 8-5), a neuroprotective peptide which counteracts the cellular toxicity generated by the overactivity of glutamatergic transmission, promoting neuronal growth. Indeed, some studies suggest that this polypeptide can enhance memory functions in behavioral paradigms. 5‐HT4 receptor activation may, therefore, improve cognitive functions by facilitating the release of this neuroprotective peptide (Lezoualc’h and Robert, 2003). In addition, the 5‐HT4 receptor may enhance cognitive performance through potentiating hippocampal LTP, the cellular mechanism proposed to underlie memory. Indeed, it has been demonstrated that activation of the 5‐HT4 receptor induces depolarization of pyramidal cells within the CA1 field (Chapin et al., 2002). The 5‐HT4 receptor may also have a role in the generation of anxiety. 5‐HT4 receptor antagonists, for example, have been shown to exhibit anxiolytic properties (Kennett et al., 1997c), whereas the 5‐HT4 receptor knockout mouse exhibited abnormal responses to stress, whereby stress‐induced hypophagia was attenuated in the knockout mouse compared with the wild‐type strain (Compan et al., 2004). Further investigation of 5‐HT4 receptor antagonists may, therefore, identify new compounds to treat anxiety.

5.5 The 5‐ht5 Receptors The 5‐ht5 subfamily contains the 5‐ht5A and 5‐ht5B receptors and despite nearly 15 years since their discovery, they remain the least well understood 5‐HT receptor subtypes with no conclusive evidence as to how they elicit second‐messenger responses despite their structural classification as GPCRs. Physiological roles for these receptor subtypes have not been identified, hence they retain lower‐case appellation to emphasize their current status as gene products, contrasting with the upper‐case notation of a receptor with known cellular functions. Of the two subtypes, the 5‐ht5A receptor has been the focus of most investigations. Currently, opinion favors negative coupling of the 5‐ht5A receptor to adenylate cyclase within recombinant cell systems (Francken et al., 1998; Hurley et al., 1998), though other reports have detected no such response (Grailhe et al., 2001). It has also been suggested that the 5‐ht5A receptor may induce intracellular Ca2þ mobilization (Noda et al., 2003) or couple to an inwardly rectifying potassium channel (Grailhe et al., 2001). In 2000, Oliver et al. (Oliver et al., 2000) conducted the first extensive study of 5‐ht5A receptor protein expression in the rat brain using immunohistochemistry. Immunoreactivity appeared to be associated with neurons, and was most strongly identified in the hypothalamus, raphe nuclei, locus coeruleus, horizontal nucleus of the diagonal band, and amygdala, with moderate staining in many regions of the cortex (particularly entorhinal cortex), the hippocampus, lateral habenula, substantia nigra, VTA, pons, and cerebellum. In situ hybridization using human brain tissue has demonstrated 5‐ht5A receptor transcripts in the cortex, hippocampus, amygdala, and cerebellum (Pasqualetti et al., 1998).

5‐Hydroxytryptamine in the central nervous system

8

. Figure 8-5 Activation of the h5‐HT4(g) receptor increases the release of nonamyloidogenic sAPPa. (a and b) Representative immunoblots showing the effects of increasing concentrations of 5‐HT (a) and prucalopride (b) on the cellular release of sAPPa in CHO cells stably coexpressing the h5‐HT4(g) receptor and human APP695. After incubating the cells with the indicated concentrations of ligands for 30 min, secreted sAPPa was measured by Western blot using the monoclonal antibody 6E10. A 110‐kDa molecular weight marker is indicated at the right. (c) Representative immunoblot showing the blocking effect of the selective 5‐HT4 antagonist, GR 133808, on 5‐HT and pruclapride‐induced sAPPa release. CHO cells were preincubated with 1 mM concentration of GR113808 10 min before treatment with 5‐HT4 ligands. After an additional 30‐min period, sAPPa was detected in the culture medium. Immunoblots were performed as in (a) and (b). CT, untreated control cells. Experiments were repeated at least three times with similar results (Data are reproduced with permission from the authors (Lezoualc’h and Robert, 2003))

Although no definitive role for the 5‐ht5A receptor has yet been elucidated, hindered greatly by the lack of an available selective ligand although this position appears to be changing (Corbett et al., 2005; Thomas, 2006), a few studies have suggested putative functions. The 5‐ht5A receptor knockout mouse, for example, exhibited increased levels of exploratory behavior in response to a novel environment as well as an attenuated response to the nonselective 5‐HT receptor ligand, LSD, suggesting that the 5‐ht5A receptor may mediate some of the behavioral effects elicited by this drug of abuse (Grailhe et al., 1999). The 5‐ht5A receptor may also be involved in the regulation of circadian rhythm in rats, although overlapping pharmacology with the 5‐HT7 receptor complicates interpretation (Sprouse et al., 2004). The development of compounds with selectivity for the 5‐ht5A receptor will greatly facilitate the elucidation of a physiological role for this receptor subtype. One such compound reported recently, SB 699551‐A (Corbett et al., 2005; Thomas, 2006), exhibits a 30‐fold selectivity for the human 5‐ht5A receptor over other 5‐HT receptor subtypes and other neurotransmitter receptors, aside from the serotonin transporter, for which it has only a

191

192

8

5‐Hydroxytryptamine in the central nervous system

tenfold selectivity (Thomas et al., 2006). Unfortunately, it appears that unlike the human and guinea pig receptors, this compound has a low affinity for the mouse and rat 5‐ht5A receptors (pKi ¼ 6.3), which limits the value of this compound in elucidating 5‐ht5A receptor function through common rodent paradigms. The 5‐ht5B receptor has attracted even less attention than the 5‐ht5A receptor, no doubt due to the discovery that the human 5‐ht5B gene sequence contains stop codons within its open reading frame, and therefore is not expected to encode a functional protein (Grailhe et al., 2001). The rat and mouse 5‐ht5B receptors, however, appear to be expressed and may be functional, though no evidence has been presented to support this latter notion. Identification of 5‐ht5B mRNA in the rat brain demonstrated expression in the hippocampus, habenula, entorhinal and piriform cortices, and the olfactory bulb (Matthes et al., 1993).

5.6 The 5‐HT6 Receptor The sequence for the 5‐HT6 receptor was first identified in 1993 (Monsma et al., 1993; Ruat et al., 1993) yet in terms of functional significance, this receptor remains relatively poorly understood. Investigations into the location of the receptor have identified receptor transcripts in the striatum, nucleus accumbens, hippocampus, and olfactory tubercles of rat and guinea pig brain tissue (Ruat et al., 1993; Ward et al., 1995). Correspondingly, the 5‐HT6 receptor protein has been identified using selective antibodies in the cortex (frontal, entorhinal, and piriform), nucleus accumbens, cerebellum, caudate putamen, substantia nigra, hippocampus (in particular, the dentate gyrus and CA1), and olfactory tubercles (Gerard et al., 1997; Hamon et al., 1999). Higher resolution studies using electron microscopy have identified the ultrastructural distribution of the protein, which is expressed postsynaptically by dendrites in the hippocampus and striatum (Hamon et al., 1999). It should be noted that while rat and human tissue readily express the 5‐HT6 receptor, the mouse brain appears to express very low levels of the receptor, thus most studies attempting to define functions for the 5‐HT6 receptor have used rat models (Hirst et al., 2003). Although the 5‐HT6 receptor is found predominantly in the CNS, it has also been detected nonneuronally within the thymus, spleen, and lymphocytes (Stefulj et al., 2000). The 5‐HT6 receptor has been demonstrated to be positively coupled to adenylate cyclase in both recombinant systems and in pig striatal tissue (Monsma et al., 1993; Schoeffter and Waeber, 1994). The receptor is thought to modulate the activity of various neurotransmitter systems, including acetylcholine, dopamine, noradrenaline, and glutamate, although some of the evidence currently appears to be conflicting. Inhibition of the 5‐HT6 receptor appears to facilitate cholinergic transmission, with evidence arising from studies using antisense oligonucleotides to knock down receptor expression, which produced a behavioral syndrome consisting of yawning, chewing, and stretching (Bourson et al., 1995). These behavioral responses were blocked with the muscarinic acetylcholine receptor antagonist, atropine. Reassuringly, a similar behavioral syndrome was induced by the 5‐HT6 receptor antagonist, Ro 04‐6790, which was also blocked by the muscarinic acetylcholine receptor antagonists (Bentley et al., 1999), implicating the central acetylcholine system in the response (Bentley et al., 1999). 5‐HT6 receptor blockade may also facilitate dopaminergic transmission. Despite some studies finding that 5‐HT6 receptor antagonists have no effect on dopaminergic transmission (Dawson et al., 2001), others have found that the administration of a 5‐HT6 receptor antagonist appears to regulate an enhanced dopaminergic system, as occurs following amphetamine administration (Pullagurla et al., 2004). In addition, increased levels of extracellular dopamine occur in the rat mPFC following administration of the 5‐HT6 receptor antagonist, SB 271046 (Lacroix et al., 2004). The 5‐HT6 receptor also regulates glutamatergic and GABAergic neurotransmission. Thus extracellular levels of glutamate in the frontal cortex and dorsal hippocampus increase following SB 271046 administration (Dawson et al., 2001), while the 5‐HT6 receptor agonist, WAY 446, elevates GABA levels in the cortex and hippocampus (Schechter et al., 2004). These findings are consistent with the identified cellular expression of the receptor (Hamon et al., 1999). The putative ability of the 5‐HT6 receptor to modulate acetylcholine release has attracted attention due to the role of this system in cognitive processes (see Mitchell and Neumaier, 2005, for a review). Generally, it is believed that 5‐HT6 receptor antagonists have a positive effect on cognition. For example, it has been

5‐Hydroxytryptamine in the central nervous system

8

shown that antagonists can relieve the amnesia induced by an anticholinergic drug (e.g., Woolley et al., 2003), though other reports have not been able to identify this response (Lindner et al., 2003). Additional studies support the positive effects of 5‐HT6 receptor antagonists on cognitive processes. For example, the 5‐HT6 receptor antagonist, Ro 04‐6790, enhanced the retention of spatial learning in the Morris water maze (Woolley et al., 2001) and consolidation in the novel‐object discrimination task (King et al., 2004), while the selective antagonists, SB 357134 and SB 399885, improved memory consolidation in an autoshaping learning task (Perez‐Garcia and Meneses, 2005). These studies suggest that 5‐HT6 receptor antagonists may have therapeutic value in reversing cognitive deficits and the results of clinical studies are eagerly awaited. A role for the 5‐HT6 receptor in the treatment of schizophrenia has also been suggested, largely due to the observation that some atypical antipsychotic compounds, including olanzapine and clozapine, have appreciable affinity for the 5‐HT6 receptor. Indeed, chronic administration of clozapine has been shown to reduce 5‐HT6 receptor mRNA levels in the rat hippocampus (Frederick and Meador‐Woodruff, 1999; > Figure 8-6). Action of these drugs at this subtype may, therefore, be responsible for some of the

. Figure 8-6 Effect of clozapine and haloperidol on 5‐HT6 mRNA expression in hippocampus. There was a significant effect of treatment. Post hoc analysis revealed that the treatment effect was entirely due to clozapine (clozapine vs. control, p < 0.01; haloperidol vs. control, p > 0.15) (Data are reproduced with permission from the authors (Frederick and Meador‐Woodruff, 1999))

therapeutic effects, though it may be argued that such interaction may mediate some of the adverse effects of the drug. In further support of a role for the 5‐HT6 receptor in the pathogenesis of schizophrenia, levels of mRNA appear to be reduced in the hippocampus of schizophrenic patients, though the density of 5‐HT6 receptor‐binding sites appears not to vary (East et al., 2002a, b). In addition, a study of Ro 04‐6790 on prepulse inhibition (PPI) in rats, a measure known to be attenuated in schizophrenia, showed no effect, hence failing to offer support that this receptor is involved in schizophrenia (Leng et al., 2003). Many association studies have been performed, analyzing putative links between polymorphisms in the 5‐HT6 receptor gene and schizophrenia. No clear association has been identified as the results have been conflicting (e.g., Shinkai et al., 1999; Tsai et al., 1999), which is often the case in such studies. The 5‐HT6 receptor has also been associated with the pathogenesis or treatment of anxiety and depression. For instance, treatment with antisense oligonucleotides resulted in a loss of 5‐HT6 receptor expression in the nucleus accumbens, in addition to enhanced anxiety levels evident in the elevated plus maze and social interaction tests (Hamon et al., 1999), which may be indicative of 5‐HT6 receptor

193

194

8

5‐Hydroxytryptamine in the central nervous system

activation having an antidepressant‐like effect. Furthermore, 5‐HT6 receptor expression may be regulated by stress hormones like corticosteroids. Thus prevention of corticosteroid release increases 5‐HT6 receptor mRNA levels in the hippocampus (Yau et al., 1997), suggesting that 5‐HT6 receptor activity may fall in response to stress, and hence might contribute to the generation of depression. Other potential therapeutic benefits of 5‐HT6 receptor ligands may include the control of seizures, as the receptor antagonist, SB 271046, displays anticonvulsant properties, though this effect was small in comparison with the efficacy of better recognized antiepileptic drugs (Routledge et al., 2000). Additionally, the 5‐HT6 receptor may be a target for antiobesity drugs, as both antisense oligonucleotide knock down of the receptor and the 5‐HT6 receptor antagonist, Ro 04‐6790, reduced rat body weight (Woolley et al., 2001).

5.7 The 5‐HT7 Receptor Unlike the other, more recently identified 5‐HT receptors, the 5‐HT7 receptor has been the subject of numerous investigations providing evidence as to its function. Although there is only one 5‐HT7 receptor gene, four distinct isoforms are generated through alternative splicing generating variations within the C terminus of the polypeptide sequence (Heidmann et al., 1997). Hence, it is possible that the different isoforms have differing abilities to couple to second‐messenger systems, though one report suggests that the three human isoforms have indistinguishable pharmacological and coupling characteristics (Krobert et al., 2001). In general, the distribution of 5‐HT7 receptor transcripts and receptor‐binding sites are similar, indicating a postsynaptic expression. A receptor autoradiographical study with human brain tissue demonstrated the distribution of 5‐HT7 receptor‐binding sites using the [3H] derivative of the selective 5‐HT7 receptor antagonist, SB 269970 (Varnas et al., 2004). 5‐HT7 receptor expression appears to be the highest in the thalamus and dentate gyrus of the hippocampus, with lower expression in the substantia nigra, VTA, dorsal raphe nucleus, Ammon’s horn of the hippocampus, cingulate cortex, amygdala, and hypothalamus. Low levels of expression have also been found in the cortex, subiculum, and parahippocampal gyrus. In the human brain, mRNA levels of 5‐HT7(a) and 5‐HT7(b) are comparable in the caudate and hippocampus, while the 5‐HT7(a) isoform is the predominant species in rat. In contrast, 5‐HT7(c) (rat) and 5‐HT7(d) (human) are expressed at relatively low levels (Heidmann et al., 1998). The 5‐HT7 receptor has been demonstrated to be positively coupled to adenylate cyclase (Bard et al., 1993), and modulates neuronal activity in various brain regions. For example in the hippocampus, activation of the 5‐HT7 receptor is believed to increase neuronal activity. For instance, the nonselective 5‐HT7 receptor agonist, 5‐CT, in the presence of WAY 100635 to block action at 5‐HT1A receptors, mediates an increase in the amplitude of the population spike (Tokarski et al., 2003). In addition, recoding from individual pyramidal neurons, 5‐HT7 receptor activation inhibits the slow AHP (sAHP) in both CA1 and CA3 cells (Bacon and Beck, 2000; Tokarski et al., 2003). A 5‐HT7 receptor‐mediated inhibition of the sAHP has also been identified in the thalamus (Goaillard and Vincent, 2002), where the 5‐HT7 receptor is also thought to increase neuronal excitability through modification of Ih, the hyperpolarization‐activated nonselective cation current (Chapin and Andrade, 2001). Within the DRN, the 5‐HT7 receptor may have a negative influence on the firing of 5‐HT neurons (Harsing et al., 2004). These authors suggested that this effect was due to the expression of 5‐HT7 heteroreceptors on the terminals of glutamatergic cortico‐raphe neurons. Alternatively, the 5‐HT7 receptor may facilitate raphe 5‐HT neuron activity via GABAergic interneurons. Roberts et al. (2004) demonstrated that the 5‐HT efflux in the guinea pig DRN was inhibited by the selective 5‐HT7 receptor antagonist, SB 269970‐A; the action was blocked by the GABAA receptor antagonist, bicuculline. They postulated that activation of the 5‐HT7 receptor induces a reduction in GABA release, hence facilitating serotonergic neuron activity. The 5‐HT7 receptor, probably along with the 5‐HT1A receptor, mediates the hypothermia induced by the nonselective agonists, 5‐CT and 8‐OH‐DPAT, since the response can be antagonized by the selective 5‐HT7 receptor antagonist, SB 269970 (Hagan et al., 2000). The 5‐HT7 receptor also induces phase shifts in the circadian rhythm of neurons within the SCN, where the receptor is expressed (Duncan et al., 1999). These 8‐OH‐DPAT‐induced phase shifts are antagonized by SB 269970 (Duncan et al., 2004). The 5‐HT7

5‐Hydroxytryptamine in the central nervous system

8

receptor may also regulate various neuroendocrine functions, such as the release of vasopressin and oxytocin (Jorgensen et al., 2003). In addition to circadian rhythm, it is also likely that the 5‐HT7 receptor regulates certain sleep phases, thus the 5‐HT7 receptor knockout mouse displays reduced levels of rapid eye movement (REM) sleep (Hedlund et al., 2005; > Figure 8-7), consistent with a reduction in REM sleep evident following administration of a selective 5‐HT7 receptor antagonist (Thomas et al., 2003). A recent study has investigated this action further by directly administering the antagonist SB 269970 to the DRN, which reduced REM sleep as a whole and also the number of REM periods (Monti and Jantos, 2006). The ability of the GABAA receptor

. Figure 8-7 Time courses of wakefulness, slow‐wave sleep, and rapid eye movement sleep over a 24‐h period. Both 5‐HT7þ/þ (open square) and 5‐HT7/ (filled square) mice showed a normal circadian sleep pattern. During the light period, 5‐HT7/ mice spent less time in rapid eye movement sleep. There was no difference between the genotypes in wakefulness or slow‐wave sleep. Values are expressed as mean  SEM. n ¼ 10 animals/group, *p < .05, two‐way repeated measures analysis of variance followed by a Bonferroni test (Data are reproduced with permission from the authors (Hedlund et al., 2005))

195

196

8

5‐Hydroxytryptamine in the central nervous system

agonist, muscimol, to prevent this response led the authors to suggest that the effects were mediated via GABAergic transmission within the DRN. The 5‐HT7 receptor is a putative therapeutic target for various other CNS disorders, including depression. Studies with the 5‐HT7 receptor knockout mouse have demonstrated a reduced immobility in forced‐swim tests and tail suspension tests, similar to behaviors exhibited by wild‐type mice following treatment with antidepressant agents (Guscott et al., 2005; Hedlund et al., 2005). Also of potential relevance, chronic treatment with the SSRI antidepressant compound, fluoxetine, and the tricyclic antidepressant drug, imipramine, reduces 5‐HT7 receptor‐binding sites in the hypothalamus (Sleight et al., 1995; Mullins et al., 1999). Additionally, treatment with citalopram and imipramine can reduce the excitatory effect of 5‐HT7 receptor activation on hippocampal bursting frequency (Tokarski et al., 2005). These studies suggest that antidepressants may exert beneficial effects through a potential downregulation of 5‐HT7 receptors. The evidence for a potential role for the 5‐HT7 receptor in schizophrenia is much less persuasive, being solely based on the affinity of some antipsychotic compounds having a high affinity for the 5‐HT7 receptor. Evidence against such a role has been provided by East et al. (1999), who could not find any difference in hippocampal 5‐HT7 receptor mRNA levels between control and schizophrenic tissue. Conversely, putative 5‐HT7 receptor involvement in epilepsy appears to be more promising. A rat model of absence epilepsy, WAG/Rij, displayed reduced epileptic activity following administration of the selective 5‐HT7 receptor antagonist, SB 258719 (Graf et al., 2004). The high level of 5‐HT7 receptor expression in the hippocampus has led to the suggestion that this receptor subtype may have a role in learning and memory. Interestingly, the 5‐HT7 receptor knockout mouse displays an impaired response to contextual fear conditioning (Roberts et al., 2004). The same report also demonstrated a reduced occurrence of LTP in the CA1 field of 5‐HT7 receptor knockout mice. Furthermore, 5‐HT7 receptor mRNA expression is increased in the hippocampus and PFC of rodents having undertaken autoshaping–training, a model of learning (Perez‐Garcia et al., 2006). Further investigation of the role of the 5‐HT7 receptor in cognitive processes may lead to the development of new clinically effective compounds in the treatment of neurological disorders.

6

The 5‐HT Transporter (SERT)

The 5‐HT transporter (SERTor 5‐HTT) is a member of the Naþ/Cl‐dependent biogenic amine transporter family, which includes the dopamine (DAT) and noradrenaline (NET) transporters (see Masson et al., 1999, for review). SERT plays a vital role within the 5‐HT system, limiting 5‐HT neurotransmission by removing the neurotransmitter through transport across the presynaptic membrane (Rudnick and Clark, 1993). Following the sequencing of rat SERT (Blakely et al., 1991), subsequent studies have suggested that SERT may exist as an oligomer in vivo. Ramamoorthy et al. (1993), for example, demonstrated that the predicted molecular weight of the transporter is
70 kDa, which contrasted with the 300 kDa estimated from the purified human placental protein. Furthermore, the transporter forms homooligomers in vitro (Kilic and Rudnick, 2000). Within the brain, SERT is located presynaptically on 5‐HT neurons, and displays central distribution closely matching the regions receiving 5‐HT neuron innervation. In situ hybridization studies in the rat demonstrated that SERT transcript was present in 5‐HT neuron cell bodies of most of the raphe nuclei in the hindbrain (Fujita et al., 1993). Correspondingly, human postmortem brain tissue autoradiography using the [3H] derivative of the tricyclic antidepressant, imipramine exhibited highest levels of binding in the raphe nuclei and midline thalamic nuclei, with weaker signals in the substantia nigra, locus coeruleus, nucleus interpeduncularis, nucleus nervi hypoglossi, nucleus nervi facialis, mammillary bodies and other regions of the hypothalamus, and low levels in the cortex, hippocampus, and amygdala (Cortes et al., 1988). Use of the SSRI, [3H]paroxetine yielded binding at highest levels in the substantia nigra, hypothalamus, and hippocampus, with lower levels in the basal ganglia and thalamus (Laruelle et al., 1988). Immunohistochemical investigation of SERT protein location in the rat brain has identified expression in the raphe nuclei, B9 and throughout the forebrain, including the hypothalamus, hippocampus, substantia nigra,

5‐Hydroxytryptamine in the central nervous system

8

amygdala, and cortex (Qian et al., 1995), in addition to the spinal cord (Sur et al., 1996). The location of SERT in these studies corresponded with 5‐HT immunoreactivity. The subcellular location of SERT appears to be dendritic and axonal, in addition to somal in the raphe nuclei (Qian et al., 1995), though some studies have been unable to demonstrate perikaryal labeling of neurons within this region (Yamamoto et al., 1998). Furthermore, electron microscopy has confirmed the predicted presynaptic location for the transporter in the rat cortex (Yamamoto et al., 1998). Despite exhibiting a predominant neuronal localization, it has been postulated that SERT may be expressed by glial cells (Lawrence et al., 1995a, b), though other reports have been unable to identify such expression (Fujita et al., 1993; Qian et al., 1995). A recent study has suggested that more than one form of SERT protein is present in vivo. Shigematsu et al. (2006) conducted immunohistochemical studies on the mouse brain with two selective antibodies, one raised against an epitope within the C terminus, the other against part of the N terminus. They observed that immunoreactivity with the N‐terminal antibody was absent in some regions, notably within the CA3 field of the hippocampus, where the C‐terminal antibody was observed to indicate SERT expression. This implies that SERT may contain variable N‐terminal domains, potentially through alternative splicing of exon 1. Interestingly, despite SERT being found predominantly, if not totally, on serotonergic neurons in the adult brain, expression of the transporter has been found to occur transiently in glutamatergic thalamocortical afferents in the developing mouse brain (Lebrand et al., 1996; Bruning and Liangos, 1997). These neurons are also immunopositive for 5‐HT, and since they are unable to synthesize this neurotransmitter, it has been suggested SERT sequesters 5‐HT, enabling the afferents to mediate serotonergic transmission during brain development. Expression of SERT within these neurons is believed to be maintained for several postnatal weeks (D’Amato et al., 1987). The actions of antidepressant drugs, in particular, the SSRIs, have led to extensive research attempting to elucidate the physiological roles of SERT in the brain. It seems indisputable that the transporter is involved in depression, though its precise mechanism of action is presently unclear. Many studies have investigated the genetic variation that occurs upstream of the SERT‐coding sequence, within the region known as 5‐HTT gene‐linked polymorphic region (5‐HTTLPR), to link the transporter with the occurrence of depression. This sequence consists of a series of repeated units, and acts as a promoter region to regulate levels of SERT expression (Lesch et al., 1996; Greenberg et al., 1999). A common polymorphism within this region is a 44‐base pair deletion, which gives rise to a short form (S), and two variations of a long form (LG and LA). The presence of the short‐form allele reduces SERT expression and activity in vitro, in contrast to cells homozygous for the long allele (Lesch et al., 1996), while a heterozygous genotype is associated with reduced SERT mRNA levels compared with a long allele genotype in the human brain (Little et al., 1998). More recently, it has been demonstrated that LG also results in levels of SERT expression comparable with the short‐form variant (Hu et al., 2006). It has been suggested that individuals carrying at least one short‐ form allele are predisposed to depressive episodes. Hariri et al. (2002) demonstrated increased neuronal activation in the human amygdala in response to fear when the short‐form allele was present. Furthermore, Caspi et al. (2003) monitored the occurrence of stressful life events in young adults and any subsequent depressive episodes, and found that the presence of the short allele was associated with a much greater incidence of depression when stressful life events were experienced. In conjunction, reduced levels of SERT binding in the brains of living depressed patients have been detected in the brain stem (Malison et al., 1998), amygdala, and midbrain (Parsey et al., 2006b), when compared with control subjects using SPECT and PET. Perhaps importantly, the reduction in SERT levels was greater in drug‐naı¨ve patients. In contrast, it does not appear that the short allele is associated with reduced SERT levels in the adult brain (Parsey et al., 2006a), though it cannot be ruled out that this polymorphism has an effect on SERT levels at earlier stages of life. Depression in monkeys, modeled through maternal separation, has also been linked with a reduction in SERT‐binding potential within the raphe nuclei and numerous forebrain regions assessed using PET analysis, suggesting that stress in early stages of life may lead to alterations in adult levels of SERT (Ichise et al., 2006). Studies of SERT function through knockout mouse strategies have also detected behavioral abnormalities that may be related to depression and anxiety. In 2003, Lira et al. (Lira et al., 2003) were unable to detect any differences in anxiety levels between SERT knockout and wild‐type strains of mice using the elevated plus maze and open‐field tests, but did identify abnormalities in the behavioral responses during

197

198

8

5‐Hydroxytryptamine in the central nervous system

paradigms designed to detect the antidepressant actions of compounds; increased immobility in the forced‐ swim test, higher rates of escape failure in shock avoidance, and longer periods of novelty‐suppressed feeding all imply a depressive‐like state. These differences may be explained by a 50% reduction in DRN neuron number and a fourfold reduction in neuron firing rate in the knockout mice. A further study also suggests that SERT knockout strains exhibit depressive or despair‐like states determined by an increase in the immobility in the tail suspension test (Zhao et al., 2006). These authors also reported a potential increase in anxiety levels. Furthermore, Adamec et al. (2006) postulate that loss of SERT may render an individual vulnerable to posttraumatic stress disorder, as SERT knockout mice displayed increased anxiety in response to predator odor exposure. Aside from a potential role in depression, SERT may be involved in other psychological disorders, including suicidal states. However, various reports have detected increased, decreased, and unchanged levels of SERT in the brains of suicide victims using autoradiographical approaches, resulting in no current consensus as to whether SERT has a role in the pathogenesis of suicidal behavior (see Purselle and Nemeroff, 2003, for an extensive review). Likewise, attempts to link the long and short alleles of the 5‐ HTTLPR locus with suicide have been similarly contradictory (Purselle and Nemeroff, 2003), thus further study is required. SERT may also have a role in the pathogenesis or treatment of schizophrenia. It has been observed that administration of SSRIs to schizophrenic patients results in the improvement of negative symptoms (Silver et al., 2000), an effect apparently mediated independently of antidepressant mechanisms, as use of an antidepressant with no effect on serotonin reuptake (maprotiline) was ineffective (Silver and Shmugliakov, 1998). Investigations of SERT‐binding sites in postmortem tissue, however, have been unable to generate a consensus as to whether levels of the transporter are aberrant in schizophrenic patients. Within the frontal cortex, for example, one study has reported no differences in the density of 5‐HT uptake sites (Gurevich and Joyce, 1997). Other potential sites of SERT‐binding changes include the caudate, hippocampus, nucleus accumbens, and putamen, though further studies are necessary before drawing of definitive conclusions. One report employed the use of PET to determine changes in SERT levels of living schizophrenic patients, and demonstrated no differences in any region assessed (midbrain, thalamus, caudate, putamen, striatum, amygdala, entorhinal cortex, and hippocampus), though the authors suggest that any changes in cortical SERT‐binding levels may be masked due to low expression levels in this region (Frankle et al., 2005). Another psychological disorder involving SERT activity may be obsessive compulsive disorder (OCD), as genetic studies have identified a potential link between OCD and the long‐form 5‐HTTLPR allele, LA (Hu et al., 2006). Recently, SERT has become an interesting potential therapeutic target in epilepsy research, as the use of two SSRIs, fluoxetine and citalopram, were shown to reduce seizures in nonsymptomatic epilepsy sufferers, completely abolishing seizures in approximately one‐third of the subjects (Albano et al., 2006). These results have tremendous potential, and study of putative SERT abnormalities in epileptic brains may identify a role for the transporter in the pathogenesis of seizures. In contrast with its role in treating various neurological disorders, SERT is also a target for various drugs of abuse, including MDMA (ecstasy) and cocaine. MDMA, for example, targets SERT, blocking 5‐HT reuptake and enhancing 5‐HT release (Pletscher et al., 1963; Rudnick and Wall, 1992). While cocaine is predominantly considered as acting on DAT, it is thought that SERT interaction contributes to the reward effects of cocaine (Rocha et al., 1998); hence, pharmacological manipulation of SERT activity may be exploited to prevent drug abuse through reward pathway inhibition.

7

Conclusions

The serotonergic system within the brain has been the subject of extensive research over the past five decades, resulting in a wealth of information regarding its anatomy, the receptors that mediate 5‐HT transmission, and the regulation of transmission through the 5‐HT reuptake transporter. Despite this intensity of activity, there remain numerous gaps in our understanding such as the possible functions of a number of the receptors (e.g., 5‐HT1D, 5‐ht1E, 5‐HT1F, 5‐ht5A, and 5‐ht5B receptors), the physiological significance of RNA editing of the 5‐HT2C receptor, alternative splicing of the 5‐HT4 and 5‐HT7 receptors,

5‐Hydroxytryptamine in the central nervous system

8

and the putative existence of homo‐ and heteromeric dimers in vivo – all have the potential to generate a vast diversity of responses to 5‐HT in the brain, and may with time, allow further manipulation of the central 5‐HT system for therapeutic benefit. The potential of each 5‐HT receptor subtype to mediate the pathophysiology of disease and psychological disorders has been discussed in this chapter, and it is already clear that the central 5‐HT system provides a number of important (potential) drug targets ranging from those to treat affective disorders such as anxiety and depression, to those relieving the symptoms of psychosis, Alzheimer’s disease, and emesis. It is perhaps not surprising, therefore, that the pharmaceutical industry maintains an intense interest in the actions of 5‐HT, and we await the development of novel therapeutics resulting from this activity.

References Abramowski D, Rigo M, Duc D, Hoyer D, Staufenbiel M. 1995. Localization of the 5‐hydroxytryptamine 2C receptor protein in human and rat brain using specific antisera. Neuropharmacology 34: 1635-1645. Adamec R, Burton P, Blundell J, Murphy DL, Holmes A. 2006. Vulnerability to mild predator stress in serotonin transporter knockout mice. Behav Brain Res 170: 126-140. Adham N, Kao HT, Schechter LE, Bard J, Olsen M, et al. 1993. Cloning of another human serotonin receptor (5‐HT1F): A 5th 5‐HT1 receptor subtype coupled to the inhibition of adenylate cyclase. PNAS 90: 408-412. Aghajanian GK. 1990. Use of Brain Slices in the Study of Serotoninergic Pacemaker Neurons of the Brainstem Raphe Nuclei. New York: John Wiley & Sons. Aghajanian GK. 1995. Electrophysiology of serotonin receptor subtypes cpathways. Bloom FR, Kupfer DJ, editors. Psychopharmacology: The Fourth Generation of Progress. Raven; New York: pp. 1451-1459. Aghajanian GK, Haigler HJ. 1974. L‐Tryptophan as a selective histochemical marker for serotonergic neurons in single‐ cell recording studies. Brain Res 81: 364-372. Aghajanian GK, Marek GJ. 1999a. Serotonin and hallucinogens. Neuropsychopharmacology 21: S16-S23. Aghajanian GK, Marek GJ. 1999b. Serotonin–glutamate interactions: A new target for antipsychotic drugs. Neuropsychopharmacology 21: S122-S133. Albano C, Cupello A, Mainardi P, Scarrone S, Favale E. 2006. Successful treatment of epilepsy with serotonin reuptake inhibitors: Proposed mechanism. Neurochem Res 31: 509-514. Albert PR, Lembo P, Storring JM, Charest A, Saucier C. 1996. The 5‐HT1A receptor: Signaling, desensitization, and gene transcription. Neuropsychopharmacology 14: 19-25. Alex KD, Yavanian GJ, McFarlane HJ, Pluto CP, Pehek EA. 2005. Modulation of dopamine release by striatal 5‐HT2C receptors. Synapse 55: 242-251. Amlaiky N, Ramboz S, Boschert U, Plassat JL, Hen R. 1992. Isolation of a mouse ‘‘5HT1E‐like’’ serotonin receptor

expressed predominantly in hippocampus. J Biol Chem 267: 19761-19764. Anthony JP, Sexton TJ, Neumaier JF. 2000. Antidepressant‐ induced regulation of 5‐HT(1b) mRNA in rat dorsal raphe nucleus reverses rapidly after drug discontinuation. J Neurosci Res 61: 82-87. Arango V, Underwood MD, Boldrini M, Tamir H, Kassir SA, et al. 2001. Serotonin 1A receptors, serotonin transporter binding and serotonin transporter mRNA expression in the brainstem of depressed suicide victims. Neuropsychopharmacology 25: 892-903. Arango V, Underwood MD, Gubbi AV, Mann JJ. 1995. Localized alterations in pre‐ and postsynaptic serotonin binding sites in the ventrolateral prefrontal cortex of suicide victims. Brain Res 688: 121-133. Artigas F, Perez V, Alvarez E. 1994. Pindolol induces a rapid improvement of depressed patients with serotonin reuptake inhibitors. Arch Gen Psychiat 51: 248-251. Ase AR, Reader TA, Hen R, Riad M, Descarries L. 2000. Altered serotonin and dopamine metabolism in the CNS of serotonin 5‐HT(1A) or 5‐HT(1B) receptor knockout mice. J Neurochem 75: 2415-2426. Audenaert K, Laere Van K, Dumont F, Vervaet M, Goethals I, et al. 2003. Decreased 5‐HT2a receptor binding in patients with anorexia nervosa. J Nucl Med 44: 163-169. Aznar S, Qian Z, Shah R, Rahbek B, Knudsen GM. 2003. The 5‐HT1A serotonin receptor is located on calbindin‐ and parvalbumin‐containing neurons in the rat brain. Brain Res 959: 58-67. Bacon WL, Beck SG. 2000. 5‐Hydroxytryptamine7 receptor activation decreases slow afterhyperpolarization amplitude in CA3 hippocampal pyramidal cells. J Pharmacol Exp Ther 294: 672-679. Bai F, Yin T, Johnstone EM, Su C, Varga G, et al. 2004. Molecular cloning and pharmacological characterization of the guinea pig 5‐HT1E receptor. Eur J Pharmacol 484: 127-139. Baker LP, Nielson MD, Impey S, Hacker BM, Poser SW, et al. 1999. Regulation and immunohistochemical localization of

199

200

8

5‐Hydroxytryptamine in the central nervous system

betagamma‐stimulated adenylyl cyclases in mouse hippocampus. J Neurosci 19: 180-192. Bard JA, Zgombick J, Adham N, Vaysse P, Branchek TA, et al. 1993. Cloning of a novel human serotonin receptor (5‐HT7) positively linked to adenylate cyclase. J Biol Chem 268: 23422-23426. Barnes JM, Barnes NM, Costall B, Ironside JW, Naylor RJ. 1989a. Identification and characterisation of 5‐HT3 recognition sites in human brain tissue. J Neurochem 53: 1787-1793. Barnes JM, Barnes NM, Costall B, Naylor RJ, Tyers MB. 1989b. 5‐HT3 receptors mediate inhibition of acetylcholine release in cortical tissue. Nature 338: 762-763. Barnes JM, Costall B, Coughlan J, Domeney AM, Gerrard PA, et al. 1990. The effects of ondansetron, a 5‐HT3 receptor antagonist, on cognition in rodents and primates. Pharmacol Biochem Behav 35: 955-962. Barnes NM, Sharp T. 1999. A review of central 5‐HT receptors and their function. Neuropharmacology 38: 1083-1152. Beck SG, Pan, YZ, Akanwa AC, Kirby LG. 2004. Median and dorsal raphe neurons are not electrophysiologically identical. J Neurophysiol 91: 994-1005. Benloucif S, Keegan MJ, Galloway MP. 1993. Serotonin‐facilitated dopamine release in vivo‐pharmacological characterisation. J Pharmacol Exp Ther 265: 373-377. Bentley JC, Bourson A, Boess FG, Fone KCF, Marsden CA, et al. 1999. Investigation of stretching behaviour induced by the selective 5‐HT6 receptor antagonist, Ro 04‐6790, in rats. Brit J Pharmacol 126: 1537-1542. Berthouze M, Ayoub M, Russo O, Rivail L, Sicsic S, et al. 2005. Constitutive dimerization of human serotonin 5‐HT4 receptors in living cells. FEBS Lett 579: 2973-2980. Bianchi C, Siniscalchi A, Beani L. 1990. 5‐HT1A agonists increase and 5‐HT3 agonists decrease acetylcholine efflux from the cerebral cortex of freely‐moving guinea‐pigs. Brit J Pharmacol 101: 448-452. Blackburn TP, Minabe Y, Middlemiss DN, Shirayama Y, Hashimoto K, et al. 2002. Effect of acute and chronic administration of the selective 5‐HT2C receptor antagonist SB‐243213 on midbrain dopamine neurons in the rat: An in vivo extracellular single cell study. Synapse 46: 129-139. Blakely RD, Berson HE, Fremeau RTJ, Caron MG, Peek MM, et al. 1991. Cloning and expression of a functional serotonin transporter from rat brain. Nature 354: 66-70. Boess FG, Beroukhim R, Martin IL. 1995. Ultrastructure of the 5‐hydroxytryptamine(3) receptor. J Neurochem 64: 1401-1405. Bojarski AJ, Paluchowska MH, Duszynska B, Bugno R, Klodzinska A, et al. 2006. Structure–intrinsic activity relationship studies in the group of 1‐imido/amido substituted 4‐(4‐arylpiperazin‐1‐yl) cyclohexane derivatives; new,

potent 5‐HT1A receptor agents with anxiolytic‐like activity. BioorganMed Chem 14: 1391-1402. Bonaventure P, Schotte A, Cras P, Leysen JE. 1997. Autoradiographic mapping of 5‐HT1B‐ and 5‐HT1D receptors in human brain using [3H]alniditan, a new radioligand. Receptor Channel 5: 225-230. Boothman LJ, Allers KA, Rasmussen K, Sharp T. 2003. Evidence that central 5‐HT2A and 5‐HT2B/C receptors regulate 5‐HT cell firing in the dorsal raphe nucleus of the anaesthetised rat. Brit J Pharmacol 139: 998-1004. Boothman LJ, Mitchell SN, Sharp T. 2006. Investigation of the SSRI augmentation properties of 5‐HT2 receptor antagonists using in vivo microdialysis. Neuropharmacology 50: 726-732. Boschert U, Amara DA, Segu L, Hen R. 1994. The mouse 5‐hydroxytryptamine1B receptor is localized predominantly on axon terminals. Neuroscience 58: 167-182. Bosker FJ, Esseveldt van KE, Klompmakers AA, Westenberg HG. 1995. Chronic treatment with fluvoxamine by osmotic minipumps fails to induce persistent functional changes in central 5‐HT1A and 5‐HT1B receptors, as measured by in vivo microdialysis in dorsal hippocampus of conscious rats. Psychopharmacology 117: 358-363. Bouhelal R, Smounya L, Bockaert J. 1988. 5‐HT1B receptors are negatively coupled with adenylate cyclase in rat substantia nigra. Eur J Pharmacol 151: 189-196. Bourson A, Borroni E, Austin RH, Monsma FJ, Sleight AJ. 1995. Determination of the role of the 5‐ht6 receptor in the rat brain: A study using antisense oligonucleotides. J Pharmacol Exp Ther 274: 173-180. Boutrel B, Monaca C, Hen R, Hamon M, Adrien J. 2002. Involvement of 5‐HT1A receptors in homeostatic and stress‐ induced adaptive regulations of paradoxical sleep: Studies in 5‐HT1A knock‐out mice. J Neurosci 22: 4686-4692. Bramley JR, Sollars PJ, Pickard GE, Dudek FE. 2005. 5‐HT1B receptor‐mediated presynaptic inhibition of GABA release in the suprachiasmatic nucleus. J Neurophysiol 93: 3157-3164. Bruinvels AT, Landwehrmeyer B, Gustafson EL, Durkin MM, Mengod G, et al. 1994. Localization of 5‐HT1B, 5‐HT1Da, 5‐HT1E and 5‐HT1F, receptor messenger RNA in rodent and primate brain. Neuropharmacology 33: 367-386. Bruinvels AT, Palacios JM, Hoyer D. 1993. Autoradiographic characterisation and localisation of 5‐HT1D compared to 5‐HT1B binding sites in rat brain. N‐S Arch Pharmacol 347: 569-582. Bruning G, Liangos O. 1997. Transient expression of the serotonin transporter in the developing mouse thalamocortical system. Acta Histochem 99: 117-121. Brunner D, Buhot MC, Hen R, Hofer M. 1999. Anxiety, motor activation, and maternal–infant interactions in 5‐HT1B knockout mice. Behav Neurosci 113: 587-601.

5‐Hydroxytryptamine in the central nervous system Burnet PWJ, Eastwood SL, Harrison PJ. 1996. 5‐HT1A and 5‐HT2A receptor mRNAs and binding site densities are differentially altered in schizophrenia. Neuropsychopharmacology 15: 442-455. Burnet PWJ, Eastwood SL, Lacey K, Harrison PJ. 1995. The distribution of 5HT1A and 5‐HT2A receptor mRNA in human brain. Brain Res 676: 157-168. Burns CM, Chu H, Rueter SM, Hutchinson LK, Canton H, et al. 1997. Regulation of serotonin‐2C receptor G‐protein coupling by RNA editing. Nature 387: 303-308. Cai X, Gu Z, Zhong P, Ren Y, Yan Z. 2002. Serotonin 5‐HT1A receptors regulate AMPA receptor channels through inhibiting Ca2þ/calmodulin‐dependent kinase II in prefrontal cortical pyramidal neurones. J Biol Chem 277: 3655336562. Carter OL, Burr DC, Pettigrew JD, Wallis GM, Hasler F, et al. 2005. Using psilocybin to investigate the relationship between attention, working memory, and the serotonin 1A and 2A receptors. J Cognitive Neurosci 17: 1497-1508. Caspi A, Sugden K, Moffitt TE, Taylor A, Craig IW, et al. 2003. Influence of life stress on depression: Moderation by a polymorphism in the 5‐HTT gene. Science 301: 386-389. Cassel JC, Jeltsch H, Neufang B, Lauth D, Szabo B, et al. 1995. Downregulation of muscarinic‐ and 5‐HT1B‐mediated modulation of [3H]acetylcholine release in hippocampal slices of rats with fimbria–fornix lesions and intrahippocampal grafts of septal origin. Brain Res 704: 153-166. Castro ME, Pascual J, Romon T, del Arco C, del Olmo E, et al. 1997. Differential distribution of [3H]sumatriptan binding sites (5‐HT1B, 5‐HT1D and 5‐HT1F receptors) in human brain: Focus on brainstem and spinal cord. Neuropharmacology 36: 535-542. Chapin EM, Andrade R. 2001. A 5‐HT7 receptor‐mediated depolarization in the anterodorsal thalamus. ii. involvement of the hyperpolarization‐activated current Ih. J Pharmacol Exp Ther 297: 403-409. Chapin EM, Haj‐Dahmane S, Torres G, Andrade R. 2002. The 5‐HT(4) receptor‐induced depolarization in rat hippocampal neurons is mediated by cAMP but is independent of I(h). Neurosci Lett 324: 1-4. Cheetham SC, Crompton MR, Katona CLE, Horton RW. 1990. Brain 5‐HT1 binding sites in depressed suicides. Psychopharmacology 102: 544-548. Chen NH, Reith ME. 1995. Monoamine interactions measured by microdialysis in the ventral tegmental area of rats treated systemically with (9)‐8‐hydroxy‐2‐(di‐n‐propylamino)tetralin. J Neurochem 64: 1585-1597. Clark MS, Sexton TJ, McClain M, Root D, Kohen R, et al. 2002. Overexpression of 5‐HT1B receptor in dorsal raphe nucleus using herpes simplex virus gene transfer increases anxiety behavior after inescapable stress. J Neurosci 22: 4550-4562.

8

Clarke WP, Yocca FD, Maayani S. 1996. Lack of 5‐hydroxytryptamine1A‐mediated inhibition of adenylyl cyclase in dorsal raphe of male and female rats. J Pharmacol Exp Ther 277: 1259-1266. Cohen H. 2005. Anxiolytic effect and memory improvement in rats by antisense oligodeoxynucleotide to 5‐hydroxytryptamine‐2A precursor protein. Depress Anxiety 22: 84-93. Compan V, Zhou M, Grailhe R, Gazzara RA, Martin R, et al. 2004. Attenuated response to stress and novelty and hypersensitivity to seizures in 5‐HT4 receptor knock‐out mice. J Neurosci 24: 412-419. Conn PJ, Sanders‐Bush E. 1984. Selective 5‐HT2 antagonists inhibit serotonin stimulated phosphatidyl inositol metabolism in cerebral cortex. Neuropharmacology 23: 993-996. Consolo S, Arnaboldi S, Giorgi S, Russi G, Ladinsky H. 1994. 5‐HT4 receptor stimulation facilitates acetylcholine release in rat frontal cortex. NeuroReport 5: 1230-1232. Corbett DF, Heightman TD, Moss SF, Bromidge SM, Coggon SA, et al. 2005. Discovery of a potent and selective 5‐ht5A receptor antagonist by high‐throughput chemistry. Bioorgan Med Chem Lett 15: 4014-4018. Cornea‐Hebert V, Watkins KC, Roth BL, Kroeze WK, Gaudreau P, et al. 2002. Similar Ultrastructural distribution of the 5‐HT2A serotonin receptor and microtubule‐associated protein MAP1A in cortical dendrites of adult rat. Neuroscience 113: 23-35. Cortes R, Soriano E, Pazos A, Probst A, Palacios JM. 1988. Autoradiography of antidepressant binding sites in the human brain: Localization using [3H]‐imipramine and [3H]‐paroxetine. Neuroscience 27: 473-496. Costall B, Kelly ME, Naylor RJ, Onaivi ES, Tyers MB. 1989. Neuroanatomical sites of action of 5‐HT3 receptor agonists and antagonists for alteration of aversive behaviour in the mouse. Brit J Pharmacol 96: 325-332. Crabbe JC, Phillips TJ, Feller DJ, Hen R, Wenger CD, et al. 1996. Elevated alcohol consumption in null mutant mice lacking 5‐HT1B serotonin receptors. Nat Genet 14: 98-101. D’Amato RJ, Blue ME, Largent BL, Lynch DR, Ledbetter DJ, et al. 1987. Ontogeny of the serotonergic projection to rat neocortex: Transient expression of a dense innervation to primary sensory areas. PNAS 84: 4322-4326. Dahlstro¨m A, Fuxe K. 1964. Evidence for the existence of monoamine containing neurons in the central nervous system. I. Demonstration of monoamines in cell bodies of brain neurons. Acta physiol Scand 62 (Suppl 232): 1-55. Darmon M, Langlois X, Suffisseau L, C‐M, Fattaccini Hamon M. 1998. Differential membrane targeting and pharmacological characterisation of chimeras of rat serotonin 5‐HT1A and 5‐HT1B receptors expressed in epithelial LLC‐PK1 cells. J Neurochem 71: 2294-2303.

201

202

8

5‐Hydroxytryptamine in the central nervous system

Davies PA, Pistis M, Hanna MC, Peters JA, Lambert JL, et al. 1999. The 5‐HT3B subunit is a major determinant of serotonin‐receptor function. Nature 397: 359-363. Dawes MA, Johnson BA, Ait‐Daoud N, Ma JZ, Cornelius JR. 2005. A prospective, open‐label trial of ondansetron in adolescents with alcohol dependence. Addict Behav 30: 1077-1085. Dawson LA, Nguyen HQ, Li P. 2001. The 5‐HT6 receptor antagonist SB‐271046 selectively enhances excitatory neurotransmission in the rat frontal cortex and hippocampus. Neuropsychopharmacology 25: 662-668. de Boer SF, Koolhass JM. 2005. 5‐HT1A and 5‐HT1B receptor agonists and aggression: A pharmacological challenge of the serotonin deficiency hypothesis. Eur J Pharmacol 526: 125-139. De Deurwaerdere P, Spampinato U. 1999. Role of serotonin2A and serotonin2B/2C receptor subtypes in the control of accumbal and striatal dopamine release elicited in vivo by dorsal raphe nucleus electrical stimulation. J Neurochem 73: 1033-1042. De Deurwaerdere P, Stinus L, Spampinato U. 1998. Opposite change of in vivo dopamine release in the rat nucleus accumbens and striatum that follows electrical stimulation of dorsal raphe nucleus: Role of 5‐HT3 receptors. J Neurosci 18: 6528-6538. Den Boer JA, Bosker FJ, Meesters Y. 2006. Clinical efficacy of agomelatine in depression: The evidence. Int Clin Psychopharmacol 21: S21-S24. Di Giovanni G, De Deurwaerdere P, Di Mascio M, Di Matteo V, Esposito E, et al. 1999. Selective blockade of serotonin‐ 2C/2B receptors enhances mesolimbic and mesostriatal dopaminergic function: A combined in vivo electrophysiological and microdialysis study. Neuroscience 91: 587-597. Di Matteo V, Di Giovanni G, Di Mascio M, Esposito E. 1999. SB 242 084, a selective serotonin2C receptor antagonist, increases dopaminergic transmission in the mesolimbic system. Neuropharmacology 38: 1195-1205. Dı´az‐Mataix L, Scorza MC, Bortolozzi A, Toth M, Celada P, et al. 2005. Involvement of 5‐HT1A receptors in prefrontal cortex in the modulation of dopaminergic activity: Role in atypical antipsychotic action. J Neurosci 25: 1083110843. Diez‐Ariza M, Garcia‐Alloza M, Lasheras B, Del Rio J, Ramirez MJ. 2002. GABA(A) receptor antagonists enhance cortical acetylcholine release induced by 5‐HT(3) receptor blockade in freely moving rats. Brain Res 956: 81-85. Done CJ, Sharp T. 1994. Biochemical evidence for the regulation of central noradrenergic activity by 5‐HT1A and 5‐HT2 receptors: Microdialysis studies in the awake and anaesthetized rat. Neuropharmacology 33: 411-421. Du Pen S, Scuderi P, Wetchler B, Sung YF, Mingus M, et al. 1992. Ondansetron in the treatment of postoperative

nausea and vomiting in ambulatory outpatients: A dose‐comparative, stratified, multicentre study. Eur J Anaesthesiol 9: 55-62. Dubin AE, Huvar R, D’Andrea MR, Pyati J, Zhu JY, et al. 1999. The pharmacological and functional characteristics of the serotonin 5‐HT3A receptor are specifically modified by a 5‐HT3B receptor subunit. J Biol Chem 274: 3079930810. Dukat M, Smith C, Herrick‐Davis K, Teitler M, Glennon RA. 2004. Binding of tryptamine analogs at h5‐HT1E receptors: A structure–affinity investigation. Bioorgan Med Chem 12: 2545-2552. Duncan MJ, Grear KE, Hoskins MA. 2004. Aging and SB‐ 269970‐A, a selective 5‐HT7 receptor antagonist, attenuate circadian phase advances induced by microinjections of serotonergic drugs in the hamster dorsal raphe nucleus. Brain Res 1008: 40-48. Duncan MJ, Short J, Wheeler DL. 1999. Comparison of the effects of aging on 5‐HT7 and 5‐HT1A receptors in discrete regions of the circadian timing system in hamsters. Brain Res 829: 39-45. Duxon MS, Flanigan TP, Reavley AC, Baxter GS, Blackburn TP, et al. 1997a. Evidence for the expression of the 5‐hydroxytryptamine2B receptor protein in the rat central nervous system. Neuroscience 76: 323-329. Duxon MS, Kennett GA, Lightowler S, Blackburn TP, Fone KC. 1997b. Activation of 5‐HT2B receptors in the medial amygdala causes anxiolysis in the social interaction test in the rat. Neuropharmacology 36: 601-608. East SZ, Burnet PW, Kerwin RW, Harrison PJ. 2002b. An RT‐ PCR study of 5‐HT(6) and 5‐HT(7) receptor mRNAs in the hippocampal formation and prefrontal cortex in schizophrenia. Schizoph Res 57: 15-26. East SZ, Burnet PWJ, Harrison PJ. 1999. Expression of the 5‐HT7 receptor in schizophrenia. Schizophr Res 36: 71. East SZ, Burnet PWJ, Leslie RA, Roberts JC, Harrison PJ. 2002a. 5‐HT6 receptor binding sites in schizophrenia and following antipsychotic drug administration: Autoradiographic studies with [125I]SB‐258585. Synapse 45: 191-199. Egashira N, Yano A, Ishigami N, Mishima K, Iwasaki K, et al. 2006. Investigation of mechanisms mediating 8‐OH‐DPAT‐ induced impairment of spatial memory: Involvement of 5‐ HT1A receptors in the dorsal hippocampus in rats. Brain Res 1069: 54-62. Fa¨rber L, Stratz T, Bruckle W, Spatz M, Pongratz D, et al. 2000. Efficacy and tolerability of tropisetron in primary fibromyalgia: A highly selective and competitive 5‐HT3 receptor antagonist. Scand J Rheumatol 113: 49-54. Federman AD, Conklin BR, Schrader KA, Reed RR, Bourne HR. 1992. Hormonal stimulation of adenylyl cyclase through Gi‐protein beta gamma subunits. Nature 356: 159-161.

5‐Hydroxytryptamine in the central nervous system Ferris CF, Melloni RH, Koppel G, Perry KW, Fuller RW, et al. 1997. Vasopressin/serotonin interaction in the anterior hypothalamus control aggressive behavior in golden hamsters. J Neurosci 17: 4331-4340. Fletcher A, Forster EA, Bill DJ, Brown G, Cliffe IA, et al. 1996. Electrophysiological, biochemical, neurohormonal and behavioural studies with WAY 100635, a potent, selective and silent 5‐HT1A receptor antagonist. Behav Brain Res 73: 337-353. Foguet M, Hoyer D, Pardo LA, Parekh A, Kluxen FW, et al. 1992. Cloning and functional characterization of the rat stomach fundus serotonin receptor. EMBO J 11: 34813487. Francken BJB, Jurzak M, Vanhauwe JFM, Luyten WHML, Leysen JE. 1998. The human 5‐ht5A receptor couples to Gi/Go proteins and inhibits adenylate cyclase in HEK 293 cells. Eur J Pharmacol 361: 299-309. Frankle WG, Narendran R, Huang Y, Hwang DR, Lombardo I, et al. 2005. Serotonin Transporter availability in patients with schizophrenia: A positron emission tomography imaging study with [11C]DASB. Biol Psychiat 57: 1510-1516. Frederick JA, Meador‐Woodruff JH. 1999. Effects of clozapine and haloperidol on 5‐HT6 receptor mRNA levels in rat brain. Schizophr Res 38: 7-12. Fujita M, Shimada S, Maeno H, Nishimura T, Tohyama M. 1993. Cellular localization of serotonin transporter mRNA in the rat brain. Neurosci Lett 162: 59-62. Gannon RL, Millan MJ. 2006. Serotonin1A autoreceptor activation by S 15535 enhances circadian activity rhythms in hamsters: Evaluation of potential interactions with serotonin2A and serotonin2C receptors. Neuroscience 137: 287-299. Garabette ML, Martin KF, Redfern PH. 2000. Circadian variation in the activity of the 5‐HT1B autoreceptor in the region of the suprachiasmatic nucleus, measured by microdialysis in the conscious freely‐moving rat. Brit J Pharmacol 131: 1569-1576. Ge J, Barnes NM. 1996. 5‐HT4 receptor mediated modulation of 5‐HT release in the rat hippocampus in vivo. Brit J Pharmacol 17: 1474-1480. Gerald C, Adham A, Kao HT, Olsen MA, Laz TM, et al. 1995. The 5‐HT4 receptor: Molecular cloning and pharmacological characterisation of two splice variants. EMBO J 14: 2806-2815. Gerard C, Martres MP, Lefevre K, Miquel MC, Verge D, et al. 1997. Immuno‐localization of serotonin 5‐HT6 receptor‐ like material in the rat central nervous system. Brain Res 746: 207-219. Goa KL, Ward A. 1986. Buspirone. A preliminary review of its pharmacological properties and therapeutic efficacy as an anxiolytic. Drugs 32: 114-129.

8

Goaillard JM, Vincent P. 2002. Serotonin suppresses the slow afterhyperpolarization in rat intralaminar and midline thalamic neurones by activating 5‐HT7 receptors. J Physiol 541: 453-465. Gobert A, Rivet JM, Cistarelli L, Millan MJ. 1997. Potentiation of the fluoxetine‐induced increase in dialysate levels of serotonin (5‐HT) in the frontal cortex of freely moving rats by combined blockade of 5‐HT1A and 5‐HT1B receptors with WAY 100,635 and GR 127,935. J Neurochem 68: 1159-1163. Graf M, Jakus R, Kantor S, Levay G, Bagdy G. 2004. Selective 5‐HT1A and 5‐HT7 antagonists decrease epileptic activity in the WAG/Rij rat model of absence epilepsy. Neurosci Lett 359: 45-48. Grailhe R, Grabtree GW, Hen R. 2001. Human 5‐HT5 receptors: The 5‐HT5A receptor is functional but the 5‐HT5B receptor was lost during mammalian evolution. Eur J Pharmacol 418: 157-167. Grailhe R, Waeber C, Dulawa SC, Hornung JP, Zhuang X, et al. 1999. Increased exploratory activity and altered response to LSD in mice lacking the 5‐HT5A receptor. Neuron 22: 581-591. Green GM, Scarth J, Dickenson A. 2000. An excitatory role for 5‐HT in spinal inflammatory nociceptive transmission; state‐dependent actions via dorsal horn 5‐HT3 receptors in the anaesthetized rat. Pain 89: 81-88. Greenberg BD, Tolliver TJ, Huang SJ, Li Q, Bengel D, et al. 1999. Genetic variation in the serotonin transporter promoter region affects serotonin uptake in human blood platelets. Am J Med Genet 88: 83-87. Grimaldi B, Bonnin A, Fillion MP, Prudhomme N, Fillion G. 1999. 5‐HT moduline: A novel endogenous peptide involved in the control of anxiety. Neuroscience 93: 1223-1225. Groenink L, Mos J, der Gugten Van J, Olivier B. 1996. The 5‐HT1A receptor is not involved in emotional stress‐induced rises in stress hormones. Pharmacol Biochem Behav 55: 303-308. Gurevich EV, Joyce JN. 1997. Alterations in the cortical serotonergic system in schizophrenia: A postmortem study. Biol Psychiat 42: 529-545. Gurevich I, Englander MT, Adlersberg M, Siegal NB, Schmauss C. 2002. Modulation of serotonin 2C receptor editing by sustained changes in serotonergic neurotransmission. J Neurosci 22: 10529-10532. Guscott M, Bristow LJ, Hadingham K, Rosahl TW, Beera MS, et al. 2005. Genetic knockout and pharmacological blockade studies of the 5‐HT7 receptor suggest therapeutic potential in depression. Neuropharmacology 48: 492-502. Hagan JJ, Price GW, Jeffrey P, Deeks NJ, Stean T, et al. 2000. Characterization of SB‐269970‐A, a selective 5‐HT(7) receptor antagonist. Brit J Pharmacol 130: 539-548.

203

204

8

5‐Hydroxytryptamine in the central nervous system

Hamblin MW, Metcalf MA. 1991. Primary structure and functional characterization of a human 5‐HT1D‐type serotonin receptor. Mol Pharmacol 40: 143-148. Hamon M, Doucet E, Lefe`vre K, Miquel MC, Lanfumey L, et al. 1999. Antibodies antisense oligonucleotide for probing the distribution and putative functions of central 5‐HT6 receptors. Neuropsychopharmacology 21: 68S-76S. Hariri AR, Mattay VS, Tessitore A, Kolachana B, Fera F, et al. 2002. Serotonin transporter genetic variation and the response of the human amygdala. Science 297: 400-403. Harsing LG, Prauda I, Barkoczy J, Matyus P, Juranyi Z. 2004. A 5‐HT7 heteroreceptor‐mediated inhibition of [3H]serotonin release in raphe nuclei slices of the rat: Evidence for a serotonergic–glutamatergic interaction. Neurochem Res 29: 1487-1497. Hartig PR, Hoyer D, Humphrey PPA, Martin GR. 1996. Alignment of receptor nomenclature with the human genome: Classification of 5‐HT1B and 5‐HT1D receptor subtypes. Trends Pharmacol Sci 17: 103-105. Harvey JA. 2003. Role of the serotonin 5‐HT2A receptor in learning. Learn Memory 10: 355-362. Hedlund PB, Huitron‐Resendiz S, Henriksen SJ, Sutcliffe JG. 2005. 5‐HT7 receptor inhibition and inactivation induce antidepressant‐like behavior and sleep pattern. Biol Psychiat 58: 831-837. Hedlund PB, Kelly L, Mazur C, Lovenberg T, Sutcliffe JG, et al. 2004. 8‐OH‐DPAT acts on both 5‐HT1A and 5‐HT7 receptors to induce hypothermia in rodents. Eur J Pharmacol 487: 125-132. Heidmann DEA, Metcalf MA, Kohen R, Hamblin MW. 1997. Four 5 hydroxytryptamine7 (5‐HT7) receptor isoforms in human and rat produced by alternative splicing: Species differences due to altered intron–exon organization. J Neurochem 68: 1372-1381. Heidmann DEA, Szot P, Kohen R, Hamblin MW. 1998. Function and distribution of three rat 5‐hydroxytryptamine7 (5‐HT7) receptor isoforms produced by alternative splicing. Neuropharmacology 37: 1621-1632. Helsley S, Siegel TL, Fiorella D, Rabin RA, Winter JC. 1998. WAY‐100635 reverses 8‐OH‐DPAT‐induced performance impairment in the radial maze. Prog Neuropsychopharmacol Biol Psychiat 22: 1179-1184. Hensler JG. 2006. Serotonergic modulation of the limbic system. Neurosci Biobehav Rev 30: 203-214. Hirst WD, Abrahamsen B, Blaney FE, Calver AR, Aloj L, et al. 2003. Differences in the central nervous system distribution and pharmacology of the mouse 5‐hydroxytryptamine‐6 receptor compared with rat and human receptors investigated by radioligand binding, site‐directed mutagenesis, and molecular modeling. Mol Pharmacol 64: 1295-1308.

Hjorth S, Sharp T. 1991. Effect of the 5‐HT1A receptor agonist 8‐OH‐DPAT on the release of 5‐HT in dorsal and median raphe innervated rat brain regions as measured by in vivo microdialysis. Life Sci 48: 1779-1786. Hornung JP. 2003. The human raphe nuclei and the serotonergic system. J Chem Neuroanatomy 26: 331-343. Hou M, Kanje M, Longmore J, Tajti J, Uddman R, et al. 2001. 5‐HT1B and 5‐HT1D receptors in the human trigeminal ganglion: Co‐localization with calcitonin gene‐related peptide, substance P and nitric oxide synthase. Brain Res 909: 112-120. Hoyer D, Clarke DE, Fozard JR, Hartig PR, Martin GR, et al. 1994. International Union of Pharmacology classification of receptors for 5‐hydroxytryptamine (serotonin). Pharmacol Rev 46: 157-204. Hu XZ, Lipsky RH, Zhu G, Akhtar LA, Taubman J, et al. 2006. Serotonin transporter promoter gain‐of‐function genotypes are linked to obsessive–compulsive disorder. Am J Human Genet 78: 815-826. Huang GJ, Herbert J. 2005. The role of 5‐HT1A receptors in the proliferation and survival of progenitor cells in the dentate gyrus of the adult hippocampus and their regulation by corticoids. Neuroscience 135: 803-813. Hurley PT, McMahon RA, Fanning P, O’Boyle KM, Rogers M, et al. 1998. Functional coupling of a recombinant human 5‐HT5A receptor to G‐proteins in HEK‐293 cells. Brit J Pharmacol 124: 1238-1244. Ichise M, Vines DC, Gura T, Anderson GM, Suomi SJ, et al. 2006. Effects of early life stress on [11C]DASB positron emission tomography imaging of serotonin transporters in adolescent peer‐ and mother‐reared rhesus monkeys. J Neurosci 26: 4638-4643. Idzikowski C, Mills FJ, James RJ. 1991. A dose–response study examining the effects of ritanserin on human slow wave sleep. Brit J Clin Pharmacol 31: 193-196. Ikeda M, Yasui M, Fukunaga H, Seshimo I, Takayama O, et al. 2005. Clinical usefulness of oral granisetron hydrochloride for alleviation of delayed nausea and vomiting induced by CPT‐11. Eur J Cancer Care 14: 435-439. Ikemoto K, Nishimura A, Okado N, Mikuni M, Nishi K, et al. 2000. Human midbrain dopamine neurons express serotonin 2A receptor: An immunohistochemical demonstration. Brain Res 853: 377-380. Inui K, Egashira N, Mishima K, Yano A, Matsumoto Y, et al. 2004. The serotonin1A receptor agonist 8‐OH‐DPAT reverses D9‐tetrahydrocannabinol‐induced impairment of spatial memory and reduction of acetylcholine release in the dorsal hippocampus in rats. Neurotox Res 6: 153-158. Jakab R, Goldman‐Rakic PS. 1998. 5‐Hydroxytryptamine2A serotonin receptors in the primate cerebral cortex: Possible

5‐Hydroxytryptamine in the central nervous system site of action of hallucinogenic and antipsychotic drugs in pyramidal cell apical dendrites. PNAS 95: 735-740. Johnson KW, Nelson DL, Dieckman DK, Wainscott DB, Lucaites VL, et al. 2003. Neurogenic dural protein extravasation induced by meta‐chlorophenylpiperazine (mCPP) involves nitric oxide and 5‐HT2B receptor activation. Cephalalgia 23: 117-123. Jorgensen H, Riis M, Knigge U, Kjaer A, Warberg J. 2003. Serotonin receptors involved in vasopressin and oxytocin secretion. J Neuroendocrinol 15: 242-249. Julius D, Huang KN, Livelli TJ, Axel R, Jessell TM. 1990. The 5HT2 receptor defines a family of structurally distinct but functionally conserved serotonin receptors. PNAS 87: 928-932. Julius D, Mac Dermott AB, Axel R, Jessell TM. 1988. Molecular characterisation of a functional cDNA encoding the serotonin1C receptor. Science 244: 558-564. Kantor S, Jakus R, Balogh B, Benko A, Bagdy G. 2004. Increased wakefulness, motor activity and decreased theta activity after blockade of the 5‐HT2B receptor by the subtype‐selective antagonist SB‐215505. Brit J Pharmacol 142: 1332-1342. Kantor S, Jakus R, Molnar E, Gyongyosi N, Toth A, et al. 2005. Despite similar anxiolytic potential, the 5‐hydroxytryptamine 2C receptor antagonist SB‐242084 [6‐chloro‐5‐ methyl‐1‐[2‐(2‐methylpyrid‐3‐yloxy)‐pyrid‐5‐yl carbamoyl] indoline] and chlordiazepoxide produced differential effects on electroencephalogram power spectra. J Pharmacol Exp Ther 315: 921-930. Kapczinski F, Lima MS, Souza JS, Cunha A, Schmitt R. 2003. Antidepressants for generalized anxiety disorder. The Cochrane Database of Systematic Reviews Issue 2: Article No.: CD003592. Kelley SP, Bratt AM, Hodge CW. 2003. Targeted gene deletion of the 5‐HT3A receptor subunit produces an anxiolytic phenotype in mice. Eur J Pharmacol 461: 19-25. Kennett GA, Ainsworth K, Trail B, Blackburn TP. 1997a. BW 723C86, a 5‐HT2B receptor agonist, causes hyperphagia and reduced grooming in rats. Neuropharmacology 36: 233-239. Kennett GA, Bright F, Trail B, Blackburn TP, Sanger GJ. 1997c. Anxiolytic‐like actions of the selective 5‐HT4 receptor antagonists, SB204070A and SB207266A in rats. Neuropharmacology 36: 707-712. Kennett GA, Trail B, Bright F. 1998. Anxiolytic‐like actions of BW 723C86 in the rat Vogel conflict test are 5‐HT2B receptor mediated. Neuropharmacology 37: 1603-1610. Kennett GA, Wood MD, Bright F, Cilia J, Piper DC, et al. 1996. In vitro and in vivo profile of SB 206553, a potent

8

5‐HT2C:5‐HT2B receptor antagonist with anxiolytic‐like properties. Brit J Pharmacol 117: 427-434. Kennett GA, Wood MD, Bright F, Trail B, Riley G, et al. 1997b. SB 242084, a selective and brain penetrant 5‐HT2C receptor antagonist. Neuropharmacology 36: 609-620. Kepe V, Barrio JR, Huang SC, Ercoli L, Siddarth P, et al. 2006. Serotonin 1A receptors in the living brain of Alzheimer’s disease patients. PNAS 103: 702-707. Kia HK, Brisorgueil MJ, Daval G, Langlois X, Hamon M, et al. 1996a. Serotonin 5‐HT1A receptors expressed by a subpopulation of cholinergic neurons in the rat medial septum and diagonal band of Broca – a double immunocytochemical study. Neuroscience 74: 143-154. Kia HK, Brisorgueil MJ, Hamon M, Calas A, Verge D. 1996b. Ultrastructural localization of 5‐hydroxytryptamine1A receptors in rat brain. J Neurosci Res 46: 697-708. Kilic F, Rudnick G. 2000. Oligomerization of serotonin transporter and its functional consequences. PNAS 97: 31063111. King MV, Sleight AJ, Woolley ML, Topham IA, Marsden CA, et al. 2004. 5‐HT6 receptor antagonists reverse delay‐ dependent deficits in novel object discrimination by enhancing consolidation – an effect sensitive to NMDA receptor antagonism. Neuropharmacology 47: 195-204. Kobilka BK, Frielle T, Collins S, Yang‐Feng T, Kobilka TS, et al. 1987. An intronless gene encoding a potential member of the family of receptors coupled to guanine nucleotide regulatory proteins. Nature 329: 75-77. Koyama S, Matsumoto N, Kubo C, Akaike N. 2000. Presynaptic 5‐HT3 receptor‐mediated modulation of synaptic GABA release in the mechanically dissociated rat amygdala neurons. J Physiol 529: 373-383. Krobert KA, Bach T, Syversveen T, Kvingedal AM, Levy FO. 2001. The cloned human 5‐HT7 receptor splice variants: A comparative characterization of their pharmacology, function and distribution. N‐S Arch Pharmacol 363: 620-632. Lacroix LP, Dawson LA, Hagan JJ, Heidbreder CA. 2004. 5‐HT6 receptor antagonist SB‐271046 enhances extracellular levels of monoamines in the rat medial prefrontal cortex. Synapse 51: 158-164. Laruelle M, Vanisberg MA, Maloteaux JM. 1988. Regional and subcellular localization in human brain of [3H]paroxetine binding, a marker of serotonin uptake sites. Biol Psychiat 24: 299-309. Lawrence JA, Charters AR, Butcher SP, Kelly JS, Olverman HJ. 1995a. Recognition of 5‐HT transporter by antipeptide antibodies. Biochem Soc Trans 23: 473. Lawrence JA, Charters AR, Butcher SP, Kelly JS, Olverman HJ. 1995b. 5‐HT transporter antibodies as a tool in serotonergic synaptosomal isolation. Biochem Soc Trans 23: 1155.

205

206

8

5‐Hydroxytryptamine in the central nervous system

Lebrand C, Cases O, Aldebrecht C, Doye A, Alvarez C, et al. 1996. Transient uptake and storage of serotonin in developing thalamic neurons. Neuron 17: 823-835. Lelong V, Dauphin F, Boulouard M. 2001. RS 67333 and D‐cycloserine accelerate learning acquisition in the rat. Neuropharmacology 41: 517-522. Leng A, Ouagazzal A, Feldon J, Higgins GA. 2003. Effect of the 5‐HT6 receptor antagonists Ro 04‐6790 and Ro 65‐7199 on latent inhibition and prepulse inhibition in the rat: Comparison to clozapine. Pharmacol Biochem Behav 75: 281-288. Lesch KP, Bengel D, Heils A, Sabol SZ, Greenberg BD, et al. 1996. Association of anxiety‐related traits with a polymorphism in the serotonin transporter gene regulatory region. Science 274: 1527-1531. Levy FO, Gudermann T, Birnbaumer M, Kaumann AJ, Birnbaumer L. 1992. Molecular cloning of a human gene (S31) encoding a novel serotonin receptor mediating inhibition of adenylyl cyclase. FEBS Lett 296: 201-206. Lezoualc’h F, Robert SJ. 2003. The serotonin 5‐HT4 receptor and the amyloid precursor protein processing. Exp Gerontol 38: 159-166. Li Q, Holmes A, Ma L, de Kar Van LD, Garcia F, et al. 2004. Medial hypothalamic 5‐hydroxytryptamine (5‐HT)1A receptors regulate neuroendocrine responses to stress and exploratory locomotor activity: Application of recombinant adenovirus containing 5‐HT1A sequences. J Neurosci 24: 10868-10877. Lin D, Parsons LH. 2002. Anxiogenic‐like effect of serotonin (1B) receptor stimulation in the rat elevated plus‐maze. Pharmacol Biochem Behav 71: 581-587. Lindner MD, Hodges DB, Hogan JB, Orie AF, Corsa JA, et al. 2003. An assessment of the effects of serotonin 6 (5‐HT6) receptor antagonists in rodent models of learning. J Pharmacol Exp Ther 307: 682-691. Lira A, Zhou M, Castanon N, Ansorge MS, Gordon JA, et al. 2003. Altered depression‐related behaviors and functional changes in the dorsal raphe nucleus of serotonin transporter‐deficient mice. Biol Psychiat 54: 960-971. Little KY, Mclaughlin DP, Zhang L, Livermore CS, Dalack GW, et al. 1998. Cocaine, ethanol, and genotype effects on human midbrain serotonin transporter binding sites and mRNA levels. Am J Psychiat 155: 207-213. Lopez‐Gimenez JF, Mengod G, Palacios JM, Vilaro MT. 1997. Selective visualisation of rat brain 5‐HT2A receptors by autoradiography with [3H]MDL 100 907. N‐S Arch Pharmacol 356: 446-454. Lowther S, De Paermentier F, Cheetham SC, Crompton MR, Katona CL, et al. 1997. 5‐HT1A receptor binding sites in post‐mortem brain samples from depressed suicides and controls. J Affect Disorders 42: 199-207.

Lucaites VL, Krushinski JH, Schaus JM, Audia JE, Nelson DL. 2005. [3H]LY334370, a novel radioligand for the 5‐HT1F receptor. II. Autoradiographic localization in rat, guinea‐ pig, monkey and human brain. N‐S Arch Pharmacol 371: 178-184. Lucas G, Compan V, Charnay Y, Neve RL, Nestler EJ, et al. 2005. Frontocortical 5‐HT4 receptors exert positive feedback on serotonergic activity: Viral transfections, subacute and chronic treatments with 5‐HT4 agonists. Biol Psychiat 57: 918-925. Lucas G, Spampinato U. 2000. Role of striatal serotonin2A and serotonin2C receptor subtypes in the control of in vivo dopamine outflow in the rat striatum. J Neurochem 74: 693-701. Lucki I, Singh A, Kreiss DS. 1994. Antidepressant‐like behavioral effects of serotonin receptor agonists. Neurosci Biobehav Rev 18: 85-95. Ma QP. 2001. Co‐localization of 5‐HT1B/1D/1F receptors and glutamate in trigeminal ganglia in rats. NeuroReport 12: 1589-1591. Machu TK, Harris RA. 1994. Alcohols and anesthetics enhance the function of 5‐hydroxytryptamine3 receptors expressed in Xenopus laevis oocytes. J Pharmacol Exp Ther 271: 898-905. Malberg JE, Eisch AJ, Nestler EJ, Duman RS. 2000. Chronic antidepressant treatment increases neurogenesis in adult rat hippocampus. J Neurosci 20: 9104-9110. Malison RT, Price LH, Berman R, Dyck van CH, Pelton GH, et al. 1998. Reduced brain serotonin transporter availability in major depression as measured by [123I]‐2 beta‐carbomethoxy‐3 beta‐(4‐iodophenyl)tropane and single photon emission computed tomography. Biol Psychiat 44: 10901098. Martin KF, Hannon S, Phillips I, Heal DJ. 1992. Opposing roles for 5‐HT1B and 5‐HT3 receptors in the control of 5‐HT release in rat hippocampus in vivo. Brit J Pharmacol 106: 139-142. Martin‐Ruiz R, Puig MV, Celada P, Shapiro DA, Roth BL, et al. 2001. Control of serotonergic function in medial prefrontal cortex by serotonin‐2A receptors through a glutamate dependent mechanism. J Neurosci 21: 9856-9866. Masson J, Sagne C, Hamon M, El Mestikawy S. 1999. Neurotransmitter transporters in the central nervous system. Pharmacol Rev 51: 439-464. Massot O, Rousselle JC, Fillion MP, Grimaldi B, Cloez‐ Tayarani I, et al. 1996. 5‐hydroxytryptamine‐moduline, a new endogenous cerebral tetrapeptide, controls the serotonergic activity via its specific interaction with 5‐ hydroxytryptamine1B/1D receptors. Mol Pharmacol 50: 752-762.

5‐Hydroxytryptamine in the central nervous system Matthes H, Boschert U, Amlaiky N, Grailhe R, Plassat JL, et al. 1993. Mouse 5‐hydroxytryptamine 5A and 5‐hydroxytryptamine 5B receptors define a new family of serotonin receptors: Cloning, functional expression, and chromosomal localization. Mol Pharmacol 43: 313-319. Maura G, Marcoli M, Tortarolo M, Andrioli GC, Raiteri M. 1998. Glutamate release in human cerebral cortex and its modulation by 5‐hydroxytryptamine acting at h5‐HT1D receptors. Brit J Pharmacol 123: 45-50. McAllister G, Charlesworth A, Snodin C, Beer MS, Noble AJ, et al. 1992. Molecular cloning of a serotonin receptor from human brain (5‐HT1E): A fifth 5‐HT1‐like subtype. PNAS 89: 5517-5521. Meltzer HY, Li Z, Kaneda Y, Ichikawa J. 2003. Serotonin receptors: Their key role in drugs to treat schizophrenia. Prog Neuropsychopharmacol Biol Psychiat 27: 1159-1172. Messa C, Colombo C, Moresco RM, Gobbo C, Galli L, et al. 2003. 5‐HT(2A) receptor binding is reduced in drug‐naive and unchanged in SSRI‐responder depressed patients compared to healthy controls: A PET study. Psychopharmacology 167: 72-78. Mialet J, Berque‐Bestel I, Sicsic S, Langlois M, Fischmeister R, et al. 2000. Pharmacological characterization of the human 5‐HT4(d) receptor splice variant stably expressed in Chinese hamster ovary cells. Brit J Pharmacol 131: 827-835. Middlemiss DN, Fozard JR. 1983. 8‐Hydroxy‐2‐(di‐n‐propylamino)‐tetralin discriminates between subtypes of the 5‐HT1 recognition site. Eur J Pharmacol 90: 151-153. Millan MJ, Gobert A, Roux S, Porsolt R, Meneses A, et al. 2004. The serotonin1A receptor partial agonist S15535 [4‐(benzodioxan‐5‐yl)1‐(indan‐2‐yl)piperazine] enhances cholinergic transmission and cognitive function in rodents: A combined neurochemical and behavioral analysis. J Pharmacol Exp Ther 311: 190-203. Miller KJ, Teitler M. 1992. Quantitative autoradiography of 5‐ CT sensitive (5‐HT1D) and 5‐CT insensitive (5‐HT1E) serotonin receptors in human brain. Neurosci Lett 136: 223-226. Miner LAH, Backstrom JR, Sanders‐Bush E, Sesack SR. 2003. Ultrastructural localization of serotonin2A receptors in the middle layers of the rat prelimbic prefrontal cortex. Neuroscience 116: 107-117. Mitchell ES, Neumaier JF. 2005. 5‐HT6 receptors: A novel target for cognitive enhancement. Pharmacol Ther 108: 320-333. Molliver ME. 1987. Serotonergic neuronal systems: What their anatomic organization tells us about function. J Clin Psychopharmacol 7: 3S-23S. Monsma FJ, Shen Y, Ward RP, Hamblin MW, Sibley DR. 1993. Cloning and expression of a novel serotonin receptor with

8

high affinity for tricyclic psychotropic drugs. Mol Pharmacol 43: 320-327. Monti JM, Jantos H. 2006. Effects of the 5‐HT7 receptor antagonist SB‐269970 microinjected into the dorsal raphe nucleus on REM sleep in the rat. Behav Brain Res 167: 245-250. Morales M, Bloom FE. 1997. The 5‐HT3 receptor is present in different subpopulations of GABAergic neurons in the rat telencephalon. J Neurosci 17: 3157-3167. Moser PC, Bergis OE, Jegham S, Lochead A, Duconseille E, et al. 2002. SL65.0155, a novel 5‐hydroxytryptamine4 receptor partial agonist with potent cognition‐enhancing properties. J Pharmacol Exp Ther 302: 731-741. Mosher T, Hayes D, Greenshaw A. 2005. Differential effects of 5‐HT2C receptor ligands on place conditioning and locomotor activity in rats. Eur J Pharmacol 515: 107-116. Mullins UL, Gianutsos G, Eison AS. 1999. Effects of antidepressants on 5‐HT7 receptor regulation in the rat hypothalamus. Neuropsychopharmacol 21: 352-367. Nakai K, Fujii T, Fujimoto K, Suzuki T, Kawashima K. 1998. Effect of WAY‐100135 on the hippocampal acetylcholine release potentiated by 8‐OH‐DPAT, a serotonin1A receptor agonist, in normal and p‐chlorophenylalanine‐ treated rats as measured by in vivo microdialysis. Neurosci Res 31: 23-29. Nibuya M, Morinobu S, Duman RS. 1995. Regulation of BDNF and trkB mRNA in rat brain by chronic electroconvulsive seizure and antidepressant drug treatments. J Neurosci 15: 7539-7547. Niswender CM, Herrick‐Davis K, Dilley GE, Meltzer HY, Overholser JC, et al. 2001. RNA editing of the human serotonin 5‐HT2C receptor: Alterations in suicide and implications for serotonergic pharmacotherapy. Neuropsychopharmacology 24: 478-491. Noda M, Yasuda S, Okada M, Higashida H, Shimada A, et al. 2003. Recombinant human serotonin 5A receptors stably expressed in C6 glioma cells couple to multiple signal transduction pathways. J Neurochem 84: 222-232. O’Connor JJ, Kruk ZL. 1994. Effects of 21 days treatment with fluoxetine on stimulated endogenous 5‐hydroxytryptamine overflow in the rat dorsal raphe and suprachiasmatic nucleus studied using fast cyclic voltammetry in vitro. Brain Res 640: 328-335. Okubo Y, Suhara T, Suzuki K, Kobayashi K, Inoue O, et al. 2000. Serotonin 5‐HT2 receptors in schizophrenic patients studied by positron emission tomography. Life Sci 66: 2455-2464. Okuhara DY, Beck SG. 1998. Corticosteroids alter 5‐hydroxytryptamine1A receptor‐effector pathway in hippocampal subfield CA3 pyramidal cells. J Pharmacol Exp Ther 284: 1227-1233.

207

208

8

5‐Hydroxytryptamine in the central nervous system

Oliver KR, Kinsey AM, Wainwright A, Sirinathsinghji DJ. 2000. Localization of 5‐ht(5A) receptor‐like immunoreactivity in the rat brain. Brain Res 867: 131-142. Pandey G, Dwivedi Y, Rizavi HS, Ren X, Pandey SC, et al. 2002. Higher expression of serotonin 5‐HT2A receptors in the postmortem brains of teenage suicide victims. Am J Psychiat 159: 419-429. Parker RM, Bentley KR, Barnes NM. 1996. Allosteric modulation of 5‐HT3 receptors: Focus on alcohols and anaesthetic agents. Trends Pharmacol Sci 17: 95-99. Parks CL, Robinson PS, Sibille E, Shenk T, Toth M. 1998. Increased anxiety of mice lacking the serotonin1A receptor. PNAS 95: 10734-10739. Parsey RV, Hastings RS, Oquendo MA, Hu X, Goldman D, et al. 2006a. Effect of a triallelic functional polymorphism of the serotonin‐transporter‐linked promoter region on expression of serotonin transporter in the human brain. Am J Psychiatr 163: 48-51. Parsey RV, Hastings RS, Oquendo MA, Huang YY, Simpson N, et al. 2006b. Lower serotonin transporter binding potential in the human brain during major depressive episodes. Am J Psychiat 163: 52-58. Pasqualetti M, Ori M, Nardi I, Castagna M, Cassano GB, et al. 1998. Distribution of the 5‐HT5A serotonin receptor mRNA in the human brain. Mol Brain Res 56: 1-8. Passani MB, Pugliese AM, Azzurrini M, Corradetti R. 1994. Effects of DAU 6125, a novel 5‐hydroxytryptamine3 (5‐HT3) antagonist on electrophysiological properties of the rat hippocampus. Brit J Pharmacol 112: 695-703. Pazos A, Hoyer D, Palacios JM. 1984. The binding of serotonergic ligands to the porcine choroid plexus: Characterisation of a new type of serotonin recognition site. Eur J Pharmacol 106: 539-546. Pazos A, Palacios JM. 1985. Quantitative autoradiographic mapping of serotonin receptors in the rat brain. I. Serotonin‐1 receptors. Brain Res 346: 205-230. Pedigo NW, Yamamura HI, Nelson DL. 1981. Discrimination of multiple [3H]5‐hydroxytrytamine binding sites by the neuroleptics spiperone in the rat brain. J Neurochem 36: 205-230. Perez‐Garcia G, Gonzalez‐Espinoa C, Meneses A. 2006. An mRNA expression analysis of stimulation and blockade of 5‐HT7 receptors during memory consolidation. Behav Brain Res 169: 83-92. Perez‐Garcia G, Meneses A. 2005. Oral administration of the 5‐HT6 receptor antagonists SB‐357134 and SB‐399885 improves memory formation in an autoshaping learning task. Pharmacol Biochem Behav 81: 673-682. Peroutka SJ, Snyder SH. 1979. Multiple serotonin receptors: Differential binding of [3H]5‐hydroxytryptamine, [3H] lysergic acid diethylamide and [3H]spiroperidol. Mol Pharmacol 16: 687-699.

Phebus LA, Johnson KW, Zgombick JM, Gilbert PJ, Belle Van K, et al. 1997. Characterisation of LY344864 as a pharmacological tool to study 5‐HTlF receptors: Binding affinities, brain penetration and activity in the neurogenic dural inflammation model of migraine. Life Sci 61: 2117-2126. Pitchot W, Hansenne M, Pinto E, Reggers J, Fuchs S, et al. 2005. 5‐Hydroxytryptamine 1A receptors, major depression, and suicidal behavior. Biol Psychiat 58: 854-858. Pletscher A, Burkard WP, Bruderer H, Gey KF. 1963. Decrease of cerebral 5‐hydroxytryptamine and 5‐hydroxyindolacetic acid by an arylalkylamine. Life Sci 2: 828-833. Pobbe RLH, Zangrossi H. 2005. 5‐HT1A and 5‐HT2A receptors in the rat dorsal periaqueductal gray mediate the antipanic‐like effect induced by the stimulation of serotonergic neurons in the dorsal raphe nucleus. Psychopharmacology 183: 314-321. Pompeiano M, Palacios JM, Mengod G. 1994. Distribution of the serotonin 5‐HT2 receptor family mRNAs: Comparision between 5‐HT2A and 5‐HT2C receptors. Mol Brain Res 23: 163-178. Popa D, Lena C, Fabre V, Prenat C, Gingrich J, et al. 2005. Contribution of 5‐HT2 receptor subtypes to sleep–wakefulness and respiratory control, and functional adaptations in knock‐out mice lacking 5‐HT2A receptors. J Neurosci 25: 11231-11238. Porras G, Di Matteo V, De Deurwaerdere P, Esposito E, Spampinato U. 2002. Central serotonin4 receptors selectively regulate the impulse dependent exocytosis of dopamine in the rat striatum: In vivo studies with morphine, amphetamine and cocaine. Neuropharmacology 43: 1099-1109. Pratt GD, Bowery NG, Kilpatrick GJ, Leslie RA, Barnes NM, et al. 1990. Consensus meeting agrees distribution of 5‐HT3 receptors in mammalian hindbrain. Trends Pharmacol Sci 11: 135-137. Pritchett DB, Bach AWJ, Wozny M, Taleb O, Dal Toso R, et al. 1988. Structure and functional expression of a cloned rat serotonin 5HT‐2 receptor. EMBO J 7: 4135-4140. Pullagurla M, Bondareva T, Young R, Glennon RA. 2004. Modulation of the stimulus effects of (þ)amphetamine by the 5‐HT6 antagonist MS‐245. Pharmacol Biochem Behav 78: 263-268. Purselle DC, Nemeroff CB. 2003. Serotonin transporter: A potential substrate in the biology of suicide. Neuropsychopharmacology 28: 613-619. Qian Y, Melikian HE, Rye DB, Levey AI, Blakely RD. 1995. Identification and characterization of antidepressant‐sensitive serotonin transporter proteins using site‐ specific antibodies. J Neurosci 15: 1261-1274. Ramamoorthy S, Leibach FH, Mahesh VB, Ganapathy V. 1993. Partial purification and characterization of the human placental serotonin transporter. Placenta 14: 449-461.

5‐Hydroxytryptamine in the central nervous system Ramboz S, Oosting R, Amara DA, Kung HF, Blier P, et al. 1998. Serotonin receptor 1A knockout: An animal model of anxiety‐related disorder. PNAS 95: 14476-14481. Ramboz S, Saudou F, Amara DA, Belzung C, Segu L, et al. 1996. 5‐HT1B receptor knock‐out mice. Behavioral consequences. Behav Brain Res 73: 305-312. Reynolds GP, Mason SL, Meldrum A, De Keczer S, Parnes H, et al. 1995. 5‐Hydroxytryptamine (5‐HT)4 receptors in post‐mortem human brain tissue: Distribution, pharmacology and effects of neurodegenerative diseases. Brit J Pharmacol 114: 993-998. Riad M, Garcia S, Watkins KC, Jodoin N, Doucet E, Langlois X, El Mestikawy S, Hamon M, Descarries L, 2000. Somatodendritic localization of 5‐HT1A and preterminal axonal localization of 5‐HT1B serotonin receptors in adult rat brain. J Comp Neurol 417: 181-194. Rick CE, Stanford IM, Lacey MG. 1995. Excitation of rat substantia nigra pars reticulata neurons by 5‐hydroxytryptamine in vitro: Evidence for a direct action mediated by 5‐hydroxytryptamine 2C receptors. Neuroscience 69: 903-913. Riemer C, Borroni E, Levet‐Trafit B, Martin JR, Poli S, et al. 2003. Influence of the 5‐HT6 receptor on acetylcholine release in the cortex: Pharmacological characterization of 4‐(2‐bromo‐6‐pyrrolidin‐1‐ylpyridine‐4‐sulfonyl)phenylamine, a potent and selective 5‐HT6 receptor antagonist. J Med Chem 46: 1273-1276. Rios M, Lambe EK, Liu R, Teillon S, Liu J, et al. 2006. Severe deficits in 5‐HT2A‐mediated neurotransmission in BDNF conditional mutant mice. J Neurobiol 66: 408-420. Roberts AJ, Krucker T, Levy CL, Slanina KA, Sutcliffe JG, et al. 2004. Mice lacking 5‐HT7 receptors show specific impairments in contextual learning. Eur J Neurosci 19: 19131922. Roberts C, Thomas DR, Bate ST, Kew JNC. 2004. GABAergic modulation of 5‐HT7 receptor‐mediated effects on 5‐HT efflux in the guinea‐pig dorsal raphe nucleus. Neuropharmacology 46: 935-941. Rocha BA, Fumagalli F, Gainetdinov RR, Jones SR, Ator R, et al. 1998. Cocaine self‐administration in dopamine transporter knockout mice. Nat Neurosci 1: 132-137. Rousselle JC, Massot O, Fillion MP, Delepierre D, Zifa E, et al. 1996. Isolation and characterization of an endogenous peptide from rat brain interacting specifically with the 5‐HT1B receptors. J Biol Chem 271: 726-735. Routledge C, Bromidge SM, Moss SF, Price GW, Hirst W, et al. 2000. Characterization of SB‐271046: A potent, selective and orally active 5‐HT(6) receptor antagonist. Brit J Pharmacol 130: 1606-1612. Ruat M, Traifford E, Arrang JM, Tardivel‐Lacombe J, Diaz J, et al. 1993. A novel rat serotonin (5‐HT6) receptor: Molecular cloning, localization and stimulation of cAMP

8

accumulation. Biochem Biophy Res Commun 193: 268-276. Rudnick G, Clark J. 1993. From synapse to vesicle: The reuptake and storage of biogenic amine neurotransmitters. Biochim Biophys Acta 1144: 249-263. Rudnick G, Wall SC. 1992. The molecular mechanism of ‘‘ecstasy’’ [3,4‐methylenedioxymethamphetamine (MDMA)]: Serotonin transporters are targets for MDMA‐induced serotonin release. PNAS 89: 1817-1821. Santarelli L, Saxe M, Gross C, Surget A, Battaglia F, et al. 2003. Requirement of hippocampal neurogenesis for the behavioural effects of antidepressants. Science 301: 805-809. Sari Y. 2004. Serotonin1B receptors: From protein to physiological function and behaviour. Neurosci Biobehav Rev 28: 565-582. Sari Y, Lefevre K, Bancila M, Quignon M, Miquel MC, et al. 1997. Light and electron microscopic immunocytochemical visualization of 5‐HT1B receptors in the rat brain. Brain Res 760: 281-286. Sari Y, Miquel MC, Brisorgueil MJ, Ruiz G, Doucet E, et al. 1999. Cellular and subcellular localization of 5‐hydroxytryptamine1B receptors in the rat central nervous system: Immunocytochemical, autoradiographic and lesion studies. Neuroscience 88: 899-915. Sarnyai Z, Sibille EL, Pavlides C, Fenster RJ, McEwen BS, et al. 2000. Impaired hippocampal‐dependent learning and functional abnormalities in the hippocampus in mice lacking serotonin1A receptors. PNAS 97: 14731-14736. Sasaki M, Ishizaki K, Obata H, Goto F. 2001. Effects of 5‐HT2 and 5‐HT3 receptors on the modulation of nociceptive transmission in rat spinal cord according to the formalin test. Eur J Pharmacol 424: 45-52. Saudou F, Amara DA, Dierich A, Le Meur M, Ramboz S, et al. 1994. Enhanced aggressive behavior in mice lacking 5‐ HT1B receptor. Science 265: 1875-1878. Schechter LE, Smith DL, Li P, Lin Q, Rosenzweig‐Lipson S, et al. 2004. WAY‐466: Pharmacological profile of a novel and selective 5‐HT6 receptor agonist. Society for Neuroscience Meeting, 394.11. Schlicker E, Fink K, Molderings GJ, Price GW, Duckworth M, et al. 1997. Effects of selective 5‐HT1B (SB‐216 641) and 5‐HT1D (BRL 15572) receptor ligands on guinea‐pig and human 5‐HT auto‐ and heteroceptors. N‐S Arch Pharmacol 356: 321-327. Schmuck K, Ullmer C, Kalkman HO, Probst A, Lubbert H. 1996. Activation of meningeal 5‐HT2B receptors: An early step in the generation of migraine headache? Eur J Neurosci 8: 959-967. Schoeffter P, Waeber C. 1994. 5‐Hydroxytryptamine receptors with a 5‐HT6 receptor‐like profile stimulating adenylyl cyclase activity in pig caudate membranes. N‐S Arch Pharmacol 350: 356-360.

209

210

8

5‐Hydroxytryptamine in the central nervous system

Segrave R, Nathan PJ. 2005. Pindolol augmentation of selective serotonin reuptake inhibitors: Accounting for the variability of results of placebo controlled double‐blind studies in patients with major depression. Hum Psychopharmacol 20: 163-174. Sharp T, Umbers V, Hjorth S. 1996. The role of 5‐HT1A autoreceptors and alpha1 adrenoceptors in the inhibition of 5‐HT release‐II NAN‐190 and SDZ 216‐525. Neuropharmacology 35: 735-741. Sharpley AL, Elliott JM, Attenburrow MJ, Cowen PJ. 1994. Slow wave sleep in humans: Role of 5‐HT2A and 5‐HT2C receptors. Neuropharmacology 33: 467-471. Sheldon PW, Aghajanian GK. 1991. Excitatory responses to serotonin (5‐HT) in neurons of the rat piriform cortex: Evidence for mediation by 5‐HT1C receptors in pyramidal cells and 5‐HT2 receptors in interneurons. Synapse 9: 208-218. Shen RY, Andrade R. 1998. 5‐Hydroxytryptamine2 receptor facilitates GABAergic neurotransmission in rat hippocampus. J Pharmacol Exp Ther 285: 805-812. Shigematsu N, Yamamoto K, Higuchi S, Fukuda T. 2006. Novel non‐uniform distribution of serotonin transporter in the mouse hippocampus and neocortex revealed by N‐ and C‐terminal domain‐specific immunohistochemistry. Brain Res 1075: 110-116. Shinkai T, Ohmori O, Kojima H, Terao T, Suzuki T, et al. 1999. Association study of the 5‐HT6 receptor gene in schizophrenia. Am J Med Genet 88: 120-122. Shippenberg TS, Hen R, He M. 2000. Region‐specific enhancement of basal extracellular and cocaine‐evoked dopamine levels following constitutive deletion of the serotonin1B receptor. J Neurochem 75: 258-265. Sibille E, Sarnyai Z, Benjamin D, Gal J, Baker H, et al. 1997. Antisense inhibition of 5‐hydroxytryptamine2a receptor induces an antidepressant‐like effect in mice. Mol Pharmacol 52: 1056-1063. Silver H, Barash I, Aharon N, Kaplan A, Poyurovsky M. 2000. Fluvoxamine augmentation of antipsychotics improves negative symptoms in psychotic chronic schizophrenic patients: A placebo‐controlled study. Int Clin Psychopharmacol 15: 257-261. Silver H, Shmugliakov N. 1998. Augmentation with fluvoxamine but not maprotiline improves negative symptoms in treated schizophrenia. J Clin Psychopharmacol 18: 208-211. Sirota P, Mosheva T, Shabtay H, Giladi N, Korczyn AD. 2000. Use of the selective serotonin 3 receptor antagonist ondansetron in the treatment of neuroleptic‐induced tardive dyskinesia. Am J Psychiat 157: 287-289. Sleight AJ, Carolo C, Petit N, Zwingelstein C, Bourson A. 1995. Identification of 5‐hydroxytryptamine7 receptor binding sites in rat hypothalamus: Sensitivity to chronic antidepressant treatment. Mol Pharmacol 47: 99-103.

Sleight AJ, Smith RJ, Marsden CA, Palfreyman MG. 1989. The effects of chronic treatment with amitriptyline and MDL 72394 on the control of 5‐HT release in vivo. Neuropharmacology 28: 477-480. Smith BN, Sollars PJ, Dudek FE, Pickard GE. 2001. Serotonergic modulation of retinal input to the mouse suprachiasmatic nucleus mediated by 5HT1B and 5HT7 receptors. J Biol Rhythms 16: 25-38. Sollars PJ, Ogilvie MD, Simpson AM, Pickard GE. 2006. Photic entrainment is altered in the 5‐HT1B receptor knockout mouse. J Biol Rhythms 21: 21-32. Spa¨th M, Welzel D, Fa¨rber L. 2000. Treatment of chronic fatigue syndrome with 5‐HT3 receptor antagonists – preliminary results. Scand J Rheumatol 29: 72-77. Sprouse J, Reynolds L, Braselton J, Schmidt A. 2004. Serotonin‐induced phase advances of SCN neuronal firing in vitro: A possible role for 5‐HT5A receptors? Synapse 54: 111-118. Sprouse JS, Aghajanian GK. 1988. Responses of hippocampal pyramidal cells to putative serotonin 5‐HT1A and 5‐HT1B agonists: A comparative study with dorsal raphe neurons. Neuropharmacology 27: 707-715. Stanford IM, Lacey MG. 1996. Differential actions of serotonin, mediated by 5‐HT1B and 5‐HT2C receptors, on GABA‐mediated synaptic input to rat substantia nigra pars reticulata neurons in vitro. J Neurosci 16: 7566-7573. Starkey SJ, Skingle M. 1994. 5‐HT1D as well as 5‐HT1A autoreceptors modulate 5‐HT release in the guinea‐pig dorsal raphe nucleus. Neuropharmacology 33: 393-402. Stefulj J, Jernej B, Cicin‐Sain L, Rinner I, Schauenstein K. 2000. mRNA expression of serotonin receptors in cells of the immune tissues of the rat. Brain Behav Immun 14: 219-224. Stevens R, Rusch D, Solt K, Raines DE, Davies PA. 2005. Modulation of human 5‐hydroxytryptamine type 3AB receptors by volatile anesthetics and n‐alcohols. J Pharmacol Exp Ther 314: 338-345. Steward LJ, Ge J, Stowe RL, Brown DC, Bruton RK, et al. 1996. Ability of 5‐HT4 receptor ligands to modulate rat striatal dopamine release in vitro and in vivo. Brit J Pharmacol 117: 55-62. Stockmeier CA, Dilley GE, Shapiro LA, Overholser JC, Thompson PA, et al. 1997. Serotonin receptors in suicide victims with major depression. Neuropsychopharmacology 16: 162-173. Stockmeier CA, Shapiro LA, Dilley GE, Kolli TM, Friedman L, et al. 1998. Increase in serotonin‐1A autoreceptors in the midbrain of suicide victims with major depression: Postmortem evidence for decreased serotonin activity. J Neurosci 18: 7394-7401. Sur C, Betz H, Schloss P. 1996. Localization of the serotonin transporter in rat spinal cord. Eur J Neurosci 8: 2753-2757.

5‐Hydroxytryptamine in the central nervous system Suzuki M, Matsuda T, Asano S, Somboonthum P, Takuma K, et al. 1995. Increase of noradrenaline release in the hypothalamus of freely moving rat by postsynaptic 5‐hydroxytryptamine1A receptor activation. Brit J Pharmacol 115: 703-711. Suzuki T, Koyama H, Sugimoto M, Uchida I, Mashimo T. 2002. The diverse actions of volatile and gaseous anesthetics on human‐cloned 5‐hydroxytryptamine3 receptors expressed in Xenopus oocytes. Anesthesiology 96: 699-704. Svenningsson P, Chergui K, Rachleff I, Flajolet M, Zhang X, et al. 2006. Alterations in 5‐HT1B receptor function by p11 in depression‐like states. Science 311: 77-80. Tatarczynska E, Antkiewicz‐Michaluk L, Klodzinska A, Stachowicz K, Chojnacka‐Wojcik E. 2005. Antidepressant‐ like effect of the selective 5‐HT1B receptor agonist CP 94253: A possible mechanism of action. Eur J Pharmacol 516: 46-50. Tecott LH, Logue SF, Wehner JM, Kauer JA. 1998. Perturbed dentate gyrus function in serotonin 5‐HT2C receptor mutant mice. PNAS 95: 15026-15031. Tecott LH, Sun LM, Akana SF, Strack AM, Lowenstein DH, et al. 1995. Eating disorder and epilepsy in mice lacking 5HT2C serotonin receptors. Nature 374: 542-546. Terry AV, Buccafusco JL, Jackson WJ, Prendergast MA, Fontana DJ, et al. 1998. Enhanced delayed matching performance in younger and older macaques administered the 5‐HT4 receptor agonist, RS17017. Psychopharmacology 135: 407-415. The GKH, Prins J, Bleijenberg G, der Meer van JWM, 2003. The effect of granisetron, a 5‐HT3 receptor antagonist, in the treatment of chronic fatigue syndrome patients – a pilot study. Neth J Med 61: 285-289. Thomas DR. 2006. 5‐ht5A receptors as a therapeutic target. Pharmacol Ther 111: 707-714. Thomas DR, Melotto S, Massagrande M, Gribble AD, Jeffrey P, et al. 2003. SB‐656104‐A, a novel selective 5‐HT7 receptor antagonist, modulates REM sleep in rats. Brit J Pharmacol 139: 705-714. Thomas DR, Soffin EM, Roberts C, Kew JN, de la Flor RM, et al. 2006. SB‐699551‐A (3‐cyclopentyl‐N‐[2‐(dimethylamino)ethyl]‐N‐[(40 ‐{[(2‐phenylethyl)amino]methyl}‐4‐ biphenylyl)methyl]propanamide dihydrochloride), a novel 5‐ht(5A) receptor‐selective antagonist, enhances 5‐HT neuronal function: Evidence for an autoreceptor role for the 5‐ht(5A) receptor in guinea pig brain. Neuropharmacology 51: 566-577. Thorre´ K, Ebinger G, Michotte Y. 1998. 5‐HT4 receptor involvement in the serotonin‐enhanced dopamine efflux from the substantia nigra of the freely moving rat: A microdialysis study. Brain Res 796: 117-124. Tokarski K, Zahorodna A, Bobula B, Grzegorzewska M, Pitra P, et al. 2005. Repeated administration of citalopram and

8

imipramine alters the responsiveness of rat hippocampal circuitry to the activation of 5‐HT7 receptors. Eur J Pharmacol 524: 60-66. Tokarski K, Zahorodna A, Bobula B, Hess G. 2003. 5‐HT7 receptors increase the excitability of rat hippocampal CA1 pyramidal neurons. Brain Res 993: 230-234. Tomkins DM, O’Neill MF. 2000. Effect of 5‐HT(1B) receptor ligands on self‐administration of ethanol in an operant procedure in rats. Pharmacol Biochem Behav 66: 129-36. Trillat AC, Malagie´ I, Scearce K, Pons D, Anmella MC, et al. 1997. Regulation of serotonin release in the frontal cortex and ventral hippocampus of homozygous mice lacking 5‐HT lB receptors: In vivo microdialysis studies. J Neurochem 69: 2019-2025. Tsai SJ, Chiu HJ, Wang YC, Hong CJ. 1999. Association study of serotonin‐6 receptor variant (C267T) with schizophrenia and aggressive behavior. Neurosci Lett 271: 135-137. Vaidya VA, Marek GJ, Aghajanian GK, Duman RS. 1997. 5‐HT2A receptor‐mediated regulation of brain‐derived neurotrophic factor mRNA in the hippocampus and the neocortex. J Neurosci 17: 2785-2795. de Kar Van LD, Javed A, Zhang y, Serres F, Raap D, et al. 2001. 5‐HT2A receptors stimulate ACTH, corticosterone, oxytocin, renin, and prolactin release and activate hypothalamic CRF and oxytocin‐expressing cells. J Neurosci 21: 35723579. Varnas K, Hall H, Bonaventure P, Sedvall G. 2001. Autoradiographic mapping of 5‐HT(1B) and 5‐HT(1D) receptors in the post mortem human brain using [(3)H]GR 125743. Brain Res 915: 47-57. Varnas K, Halldin C, Pike VW, Hall H. 2003. Distribution of 5‐HT4 receptors in the postmortem human brain – an autoradiographic study using [125I]SB 207710. Eur Neuropsychopharmacol 13: 228-234. Varnas K, Hurd YL, Hall H. 2005. Regional expression of 5‐HT1B receptor mRNA in the human brain. Synapse 56: 21-28. Varnas K, Thomas DR, Tupala E, Tiihonen J, Hakan H. 2004. Distribution of 5‐HT7 receptors in the human brain: A preliminary autoradiographic study using [3H]SB‐ 269970. Neurosci Lett 367: 313-316. Villalobos C, Beique JC, Gingrich JA, Andrade R. 2005. Serotonergic regulation of calcium‐activated potassium currents in rodent prefrontal cortex. Eur J Neurosci 22: 1120-1126. Voigt MM, Laurie DJ, Seeburg PH, Bach A. 1991. Molecular cloning and characterization of a rat brain cDNA encoding a 5‐hydroxytryptamine1B receptor. EMBO J 10: 4017-4023. Vollenweider FX, Geyer MA. 2001. A systems model of altered consciousness: Integrating natural and drug‐induced psychoses. Brain Res Bull 56: 495-507.

211

212

8

5‐Hydroxytryptamine in the central nervous system

Waeber C, Moskowitz MA. 1995a. Autoradiographic visualisation of [3H]5‐carboxamidotryptamine binding sites in the guinea pig and rat brain. Eur J Pharmacol 283: 31-46. Waeber C, Moskowitz MA. 1995b. [3H]sumatriptan labels both 5‐HT1D and 5‐HT1F receptor binding sites in the guinea pig brain: An autoradiographic study. N‐S Arch Pharmacol 352: 263-275. Ward RP, Hamblin MW, Lachowicz JE, Hoffman BJ, Sibley DR, et al. 1995. Localization of serotonin subtype 6 receptor messenger RNA in the rat brain by in situ hybridisation histochemistry. Neuroscience 64: 1105-1111. Ward SE, Harrington FP, Gordon LJ, Hopley SC, Scott CM, et al. 2005. Discovery of the first potent, selective 5‐hydroxytryptamine1D receptor antagonist. J Med Chem 48: 3478-3480. Weber ET, Gannon RL, Rea MA. 1998. Local administration of serotonin agonists blocks light‐induced phase advances of the circadian activity rhythm in the hamster. J Biol Rhythms 13: 209-218. Welsh SE, Kachelries WJ, Romano AG, Simansky KJ, Harvey JA. 1998. Effects of LSD, ritanserin, 8‐OH‐DPAT and lisuride on classical conditioning in the rabbit. Pharmacol Biochem Behav 59: 469-475. Wilkinson LO, Middlemiss DN, Hutson PH. 1994. 5‐HT1A receptor activation increases hippocampal acetylcholine efflux and motor activity in the guinea pig: Agonist efficacy influences functional activity in vivo. J Pharmacol Exp Ther 270: 656-664. Williams GV, Rao SG, Goldman‐Rakic PS. 2002. The physiological role of 5‐HT2A receptors in working memory. J Neurosci 22: 2843-2854. Willins DL, Deutch AY, Roth BL. 1997. Serotonin 5‐HT2A receptors are expressed on pyramidal cells and interneurons in the rat cortex. Synapse 27: 79-82. Wissink S, Meijer O, Pearce D, der Burg van B, der Saag van PT. 2000. Regulation of the rat serotonin‐1A receptor gene by corticosteroids. J Biol Chem 275: 1321-1326.

Woolley ML, Bentley JC, Sleight AJ, Marsden CA, Fone KCF. 2001. A role for 5‐ht6 receptors in retention of spatial learning in the Morris water maze. Neuropharmacology 41: 210-219. Woolley ML, Marsden CA, Sleight AJ, Fone KC. 2003. Reversal of a cholinergic‐induced deficit in a rodent model of recognition memory by the selective 5‐HT6 receptor antagonist, Ro 04‐6790. Psychopharmacology 170: 358-367. Yamamoto H, Fujimiya M, Shirai Y, Nakashita M, Oyasu M, et al. 1998. Immunohistochemical localization of serotonin transporter in normal and colchicine treated rat brain. Neurosci Res 32: 305-312. Yan QS, Zheng S‐Z, Feng MJ, Yan SE. 2005. Involvement of 5‐HT1B receptors within the ventral tegmental area in ethanol‐induced increases in mesolimbic dopaminergic transmission. Brain Res 1060: 126-137. Yasuno F, Suhara T, Nakayama T, Ichimiya T, Okubo Y, et al. 2003. Inhibitory effect of hippocampal 5‐HT1A receptors on human explicit memory. Am J Psychiat 160: 334-340. Yatham LN, Liddle PF, Dennie J, Shiah IS, Adam MJ, et al. 1999. Decrease in brain serotonin 2 receptor binding in patients with major depression following desipramine treatment: A positron emission tomography study with fluorine 18‐labelled setoperone. Arch Gen Psychiat 56: 705-711. Yau JLW, Noble J, Widdowson J, Seckl JR. 1997. Impact of adrenalectomy on 5‐HT6 and 5‐HT7 receptor gene expression in the rat hippocampus. Mol Brain Res 45: 182-186. Zhao S, Edwards J, Carroll J, Wuedholz L, Millstein RA, et al. 2006. Insertion mutation at the C‐terminus of the serotonin transporter disrupts brain serotonin function and emotion‐related behaviours in mice. Neuroscience 140: 321-334.

9

GABA Neurotransmission: An Overview

A. Schousboe . H. S. Waagepetersen

1

GABA—from Metabolite to Neurotransmitter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214

2 2.1 2.2 2.3

GABA Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214 GAD65 and GAD67 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214 GAD in Non‐GABAergic Neuronal Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 215 Glutamine as a GABA Precursor and Involvement of Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . 216

3 3.1 3.2

GABA Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 216 GABA‐T . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 216 Oxidative Degradation: Neurons versus Astrocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217

4 4.1 4.2

Inhibitors of GABA Synthesis and Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217 Carbonyl‐Trapping Agents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217 Active Site‐Directed Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 218

5 5.1 5.2

GABA Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 218 Vesicular Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 218 Nonvesicular Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219

6 6.1 6.2

GABA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219 Ionotropic Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219 Metabotropic Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 220

7 7.1 7.2 7.3 7.3.1

GABA Inactivation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 220 Receptor Desensitization and GABA Diffusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 220 High‐Affinity Plasma Membrane Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221 Inhibitors of GABA Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221 Functional Implications of GABA Transport Inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221

#

2008 Springer ScienceþBusiness Media, LLC.

214

9

GABA neurotransmission: An overview

Abstract: GABA neurotransmission involves biosynthesis and metabolic degradation of GABA, its stimulus‐ coupled release and receptor interaction, as well as inactivation by high‐affinity transport systems in neuronal and astrocytic plasma membranes. These entities are summarized to provide the reader with information about the fundamental properties of these processes. List of Abbreviations: AOAA, amino-oxyacetic acid; BGT-1, betaine-GABA transporter-1; CACA, cis-4aminocrotonic acid; CAMP, cis-2-(aminomethyl)cyclopropane-1-carboxylic acid; CIT, citrate; CNS, central nervous system; EF-1502, N-[4,4-bis(3-methyl-2-thienyl)-3-butenyl]-4-(methylamino)-4,5,6,7tetrahydrobenzo[d]isoxazol-3-ol; exo-THPO, 4-amino-4,5,6,7-tetrahydro-1,2-benzo[d]isoxazol-3-ol; GABA, gamma-aminobutyric acid; GABA-T, GABA-transaminase; GAD, glutamic acid decarboxylase; GAT 1–4, GABA transporters 1–4; Glu, glutamate; OAA, oxaloacetate; PAG, phosphate activated glutaminase; SSA, succinate semialdehyde; SUC, succinate; TCA, tricarboxylic acid; THIP, 4,5,6,7-tetrahydroisoxazolo[5,4-c]pyridin-3-ol; THPO, 4,5,6,7-tetrahydroisoxazolo[4,5-c]pyridin-3-ol; 2OG, 2-oxoglutarate; 3-APPA, 3-aminopropylphosphonic acid; 3-APMPA, 3-aminopropyl(methyl)phosphinic acid

1

GABA—from Metabolite to Neurotransmitter

Gamma‐aminobutyric acid (GABA), or at physiologic pH more appropriately referred to as the zwitterion gamma‐aminobutyrate, is a plant‐associated amino acid, which in 1950 by three groups of researchers was reported to be present in brain extracts (Awapara et al., 1950; Roberts and Frankel, 1950; Udenfriend, 1950). Among these, Eugene Roberts most energetically embarked on a long‐lasting research strategy aimed at elucidating the biochemistry of this intriguing amino acid and later its association with brain function and neurotransmission. Within the next ten years, the basic biochemical pathways involving GABA had been worked out in considerable detail and a putative function in neurotransmission had been envisaged. These aspects are outlined in the proceedings of the first international conference on GABA held in 1959 (Roberts et al., 1960). The present review will briefly describe the different functional entities involved in GABAergic neurotransmission, that is, metabolic pathways (> Figure 9‐1), release processes, receptor interaction, and inactivation processes. More detailed accounts of these topics can be found in other volumes.

2

GABA Synthesis

Already, at the time of its discovery in brain tissue it was clear that GABA was produced from glutamate (Roberts and Frankel, 1950), and the synthesizing enzyme L‐glutamate decarboxylase (GAD) was subsequently extensively characterized (Roberts and Simonsen, 1963; Wu and Roberts, 1974) and purified to homogeneity (Wu et al., 1973). Polyacrylamide gel electrophoresis of the purified enzyme treated with sodium dodecyl sulphate revealed multiple bands (Matsuda et al., 1973) hinting at a possible heterogeneity of the enzyme at the molecular level.

2.1 GAD65 and GAD67 Elaborate studies of GAD regulation and subsequent cloning have demonstrated the existence of two distinct molecular forms of GAD termed GAD65 and GAD67 referring to their molecular weights of 65 kDa and 67 kDa, respectively (see Martin and Rimvall, 1993; Soghomonian and Martin, 1998). These molecular forms of GAD are encoded for by two independent genes and they exhibit different properties with regard to binding of the coenzyme pyridoxal phosphate and regulation by phosphorylation. Thus, GAD65 exists to a large extent as dormant apoenzyme, which may be rapidly activated by binding of the coenzyme whereas GAD67 mainly is found as the catalytically active holoenzyme. The two isoforms of the enzyme also exhibit differences with regard to subcellular distribution, GAD67 being cytosolic and distributed throughout the GABAergic neurons, that is, both in the cell bodies and the processes. On the contrary, GAD65 is mainly

GABA neurotransmission: An overview

9

. Figure 9‐1 TCA cycle coupled to reactions involved in the GABA‐shunt, which constitutes an alternative pathway for the traditional TCA cycle. This involves the concerted action of the GABA‐metabolizing enzymes GAD and GABA‐T plus SSADH SSADH, succinate semialdehyde dehydrogenase; GAD, glutamic acid decarboxylase; GABA‐T, GABA‐transaminase; OAA, oxaloacetate; CIT, citrate; 2OG, 2‐oxoglutarate; SSA, succinate semialdehyde; SUC, succinate; Glu, glutamate; GABA, gamma‐aminobutyric acid

associated with the nerve endings, possibly in close association with vesicles storing GABA for neurotransmitter release. Both molecular forms are regulated by phosphorylation, GAD67 being inhibited by protein kinase A‐mediated phosphorylation and GAD65 being activated by phosphorylation. The difference in subcellular localization of the two GAD enzymes has led to speculations regarding distinctive functional roles in the biosynthesis of GABA in the cytoplasmic, metabolic pool and the neurotransmitter pool, respectively (Martin and Rimvall, 1993; Waagepetersen et al., 1999, 2001).

2.2 GAD in Non‐GABAergic Neuronal Systems Early immunohistochemical studies of the distribution of GAD in brain tissue (McLaughlin et al., 1974; Saito et al., 1974; Ribak et al., 1976) clearly associated GAD with GABAergic neuronal pathways, lending support to the general notion that GAD is a marker enzyme for GABAergic structures. Recent immunocytochemical studies have, however, provided evidence that GAD is likely to be present in certain glutamatergic neurons as well (Sloviter et al., 1996; Gutierrez, 2003). At present, the functional significance of this is poorly understood; however, it has been speculated that it may be related to the demonstration that GABA in addition to its neurotransmitter action can function as a neurodifferentiation molecule during early development of the CNS (Belhage et al., 1998; Waagepetersen et al., 1999; Fiszman and Schousboe, 2004). Interestingly, GABA mainly acts as an excitatory neurotransmitter at this early stage of development (Ben‐Ari et al., 1989; Cherubini et al., 1991; Ben‐Ari, 2002). It should also be noted that glutamatergic neurons may be able to accumulate GABA from surrounding GABAergic neurons, thereby maintaining a considerable intracellular concentration of GABA as recently demonstrated in cultures of dissociated cerebellum that consists mainly of glutamatergic neurons with a small population of GABAergic neurons (Sonnewald et al., 2004). Again, the functional significance of this can only be speculated upon.

215

216

9

GABA neurotransmission: An overview

2.3 Glutamine as a GABA Precursor and Involvement of Mitochondria Exogenous glutamine has repeatedly been shown in brain tissue preparations to be a more efficient substrate for GABA biosynthesis than its immediate precursor, glutamate (Reubi et al., 1978; Westergaard et al., 1995). This may actually reflect the function of the GABA–glutamine–glutamate cycle in vivo in which GABA is transferred from GABAergic neurons to surrounding astrocytes where it is transaminated to succinic semialdehyde using 2‐oxoglutarate (2-OG) to generate glutamate. This is subsequently used to generate glutamine catalyzed by glutamine synthetase, and glutamine is transferred back to the GABAergic neuron where it is hydrolyzed by phosphate activated glutaminase (PAG) to form glutamate, which is decarboxylated to form GABA (Waagepetersen et al., 2003). It should be noted that these reactions do not require any anaplerotic activity since no net usage of a tricarboxylic acid (TCA) cycle constituent is needed. Hence, GABA biosynthesis may be maintained by a stoichiometric operation of the GABA–glutamine cycle and usage of acetyl CoA derived by astroglial pyruvate dehydrogenase activity. The fact that only a small fraction of the GABA neurotransmitter pool is drained from the neurons by the astrocytic uptake of GABA (Schousboe et al., 2004a) highly increases the probability that this machinery may work. The fact that PAG and GAD have different subcellular localizations in mitochondria and cytosol, respectively (Balazs et al., 1966), may impose some complications regarding the synthesis of GABA from glutamine. Although PAG is a mitochondrial enzyme it is often assumed that the product of the reaction catalyzed by PAG, glutamate, is actually liberated through the outer mitochondrial membrane since PAG may be located at the outer surface of the inner membrane (Kvamme et al., 2001). However, other evidence suggest that glutamate may be accessible for mitochondrial metabolism (Palaiologos et al., 1988; Zieminska et al., 2004). Detailed studies of labeling patterns of glutamate and GABA using [U‐13C]glucose or [U‐13C]glutamine in GABAergic neurons under conditions where GABA release occurs selectively either from the vesicular pool or the cytoplasmic pool have provided evidence that GABA synthesis particularly in the vesicular pool involves mitochondrial TCA cycle activity (Waagepetersen et al., 1999; 2001). In fact, approximately 60% of newly synthesized vesicular GABA requires TCA cycle activity. Such a biosynthetic pathway may provide GABAergic neurons with a regulatory repertoire beyond regulation of GAD activity (Waagepetersen et al., 2003).

3

GABA Degradation

As illustrated in > Figure 9‐1, GABAergic neurons are provided with enzymes allowing the TCA cycle to operate by replacing 2‐OG dehydrogenase and succinyl‐CoA synthetase steps by the concerted action of GAD, GABA‐transaminase (GABA‐T), and succinate semialdehyde dehydrogenase (SSADH). These latter reactions are called the GABA‐shunt of the TCA cycle, resulting in a small reduction of the normal ATP production from TCA cycle activity. The flux through the GABA‐shunt may account for about 10% of the activity through the complete TCA cycle (Balazs et al., 1970; Machiyama et al., 1970). It should be noted that the reactions involved in the GABA‐shunt may operate by a concerted action of GABAergic neurons and astrocytes as well as in the former cells alone.

3.1 GABA‐T The key enzyme in GABA catabolism is GABA‐T, which is localized in the mitochondrial matrix (Schousboe et al., 1977) and present in essentially all tissues including the brain (Wu et al., 1978). It has almost identical activities in astrocytes, GABAergic neurons, and glutamatergic neurons (> Table 9‐1). The enzyme was first purified to homogeneity by Schousboe et al. (1973) and characterized with regard to substrate specificity, inhibitors, and Km values for GABA and 2-OG (Schousboe et al., 1973, 1974; Bloch‐ Tardy et al., 1974; Cash et al., 1974; Maitre et al., 1975; John and Fowler, 1976). The rather low Km values for both substrates would indicate that the enzyme is functionally active in vivo, which is compatible with the fact that GABA can be metabolized to CO2 in situ (Machiyama et al., 1970; Yu and Hertz, 1983).

GABA neurotransmission: An overview

9

. Table 9‐1 Activity of GABA‐T in mouse brain and astrocytes from mouse cerebral cortex and GABAergic and glutamatergic neurons Cell type/brain a

Mouse brain GABAergic neuronsa Glutamatergic neuronsb Astrocytes (cerebral cortex)c

GABA‐T activity (nmol
min1
mg1 protein) 3.2  1.2 (7) 1.0  0.2 (4) 1.4  0.3 (5) 1.6  0.2 (7)

Note: Enzyme activity was determined in homogenates of tissue or cells using the assay for enzyme activity described by Schousboe et al. (1973). Cultures of GABAergic neurons were prepared from embryonic cerebral cortex and glutamatergic neurons from 7‐day‐old cerebellum, as described by Hertz et al. (1989a) and Schousboe et al. (1989). Astrocytes cultures were prepared from cerebral cortex of newborn mice, as described by Hertz et al. (1989b). Neuronal cultures were grown for 8–10 days and astrocytic cultures for 3 weeks before the enzyme activity was determined. Values represent averages  SEM of n experiments a Values are from Larsson et al. (1985) b Values are from A. Schousboe, unpublished

3.2 Oxidative Degradation: Neurons versus Astrocytes The demonstration of CO2 formation from [14C]GABA in neuronal preparations may be interpreted as oxidative metabolism of GABA (Yu and Hertz, 1983). As can be seen from > Figure 9‐1, production of CO2 in the TCA cycle/GABA‐shunt does not represent a net degradation of GABA, but rather a replacement of carbon atoms by acetyl CoA. Net GABA degradation would require pyruvate recycling via the action of malic enzyme (see chapter in Volume 5.5 by Sonnewald et al., 2007). It is, however, the current notion that pyruvate recycling is much less prominent in neurons than in astrocytes (Waagepetersen et al., 2003), making it unlikely that GABA may function as an important energy source in neurons. However, in astrocytes where pyruvate recycling does take place (Waagepetersen et al., 2003) GABA can be oxidatively metabolized to CO2 and water. If this happens, it is obviously not available for the operation of the GABA– glutamine cycle. Therefore, GABA, which enters the oxidative pathway via pyruvate recycling and acetyl CoA, is lost permanently from the neurotransmitter pool. This may have functional consequences for the maintenance of optimal GABAergic activity in the CNS (Schousboe et al., 2004a, b, c).

4

Inhibitors of GABA Synthesis and Degradation

The two enzymes involved in GABA metabolism, GAD and GABA‐T, belong to the family of enzymes that require pyridoxal phosphate as coenzyme. Since pyridoxal phosphate is formed from vitamin B6 by phosphorylation, a reaction catalyzed by pyridoxal‐kinase, these enzymes are generally referred to as B6‐ requiring enzymes. This common property makes both enzymes sensitive to carbonyl‐trapping agents, which form a Schiff base with the carbonyl group in pyridoxal phosphate.

4.1 Carbonyl‐Trapping Agents Hydrazine and its derivatives are highly toxic compounds, a property partly related to their ability to form hydrazones. Thus, the effects of these compounds on the GABA metabolizing enzymes have been investigated in detail (Tapia, 1975). Along this line of research it was found that amino‐oxyacetic acid (AOAA) is a highly potent inhibitor of both enzymes with a tenfold higher affinity for GABA‐T compared with GAD (Schousboe et al., 1974; Wu and Roberts, 1974).

217

218

9

GABA neurotransmission: An overview

Hence, it is theoretically possible to use AOAA to preferentially inhibit GABA‐T but under most experimental conditions both enzymes will be affected. According to this, AOAA given to experimental animals will almost always lead to seizures, a condition associated with inhibition of GABA synthesis (Tapia and Meza‐Ruiz, 1975; Tapia et al., 1975). Another compound that was originally described as a pharmacological tool to induce seizures is mercaptopropionic acid (Sprince et al., 1969). A detailed spectroscopic and kinetic analysis of the action of 3‐mercaptopropionic acid on the purified GABA‐T from mouse brain showed that it reacts with the pyridoxal group of the coenzyme behaving as a competitive inhibitor with regard to GABA (Schousboe et al., 1974). Its Ki value for GABA‐T (13 mM) was as expected from the pharmacological action of the compound higher than that found for GAD (Wu and Roberts, 1974). Due to the pharmacological interest in compounds that might be able to effectively distinguish between GAD and GABA‐T, research programs were initiated investigating compounds that would act as active site‐ directed suicide inhibitors.

4.2 Active Site‐Directed Inhibitors From a drug development perspective, it would obviously be desirable to selectively inhibit GABA‐T since inhibition of GAD would inevitably lead to seizure activity (Tapia, 1975). Inhibition of GABA‐T could lead to increased GABA levels, thus offering protection against seizures (Tapia, 1975). Introducing alkylating entities into the GABA molecule, a substrate for GABA‐T, compounds were created, which on binding to the enzyme would form a Schiff base and simultaneously the active site would be covalently bound to the false substrate thereby irreversibly inhibiting the enzyme (Lippert et al., 1977). The most useful of this series of substrate mimetics turned out to be GABAculline and g‐vinyl GABA (Lippert et al., 1977). The latter GABA analog was subsequently developed into the first antiepileptic drug acting as a specific inhibitor of GABA‐T. These drugs lead to a significant increase in the synaptic pool of GABA (Iadarola and Gale, 1980; Wood et al., 1981; Gram et al., 1988). Interestingly, g‐vinyl GABA is more potent as an inhibitor of neuronal GABA‐T than of astroglial GABA‐T in intact cells (Larsson et al., 1986). This is not related to different affinities for GABA‐T in these cells, but may best be explained by the presence of a high‐affinity transporter for g‐vinyl GABA selectively in the neuronal plasma membrane (Schousboe et al., 1986). This transporter appears to be unrelated to the GABA transporters (Schousboe et al., 1986).

5

GABA Release

In accordance with its role as a neurotransmitter, GABA is stored in vesicles from which it can be released in a Ca2þ‐dependent manner on depolarization of the neuronal membrane (Curtis and Johnston, 1974; Schousboe et al., 1976; Otsuka, 1996). However, depending on the depolarizing conditions, GABA may also be released from the nonvesicular cytosolic pool, a mechanism that may play a physiological role (Bernath, 1991).

5.1 Vesicular Release Synaptic release of a neurotransmitter is thought to involve a vesicular pool of the transmitter and in keeping with this vesicles exist, which are equipped with transporters concentrating GABA 10‐ to 20‐fold over the cytosolic concentration (Otsuka, 1996). Vesicular, Ca2þ‐dependent GABA release has been extensively studied using a variety of brain tissue preparations such as slices, synaptosomes, and intact neurons in culture (Schousboe et al., 1976; Pin and Bockaert, 1989; Belhage et al., 1993). It may be interesting to note that in some of these preparations the relative magnitude of the vesicular release appears to be dependent on the nature of the depolarizing signal, that is, an excitatory amino acid or a high [Kþ] elicited response (Pin and Bockaert, 1989; Belhage et al., 1993).

GABA neurotransmission: An overview

9

5.2 Nonvesicular Release As discussed in detail by Bernath (1991), nonvesicular GABA release elicited by depolarization in a non‐ Ca2þ‐dependent manner can be of considerable magnitude. This was directly demonstrated to be the case in GABAergic neurons cultured from dissociated mouse cerebral cortex. Thus, the glutamate‐induced GABA release that could be inhibited by the nontransportable GABA transport blocker N‐diphenyl–butenyl– nipecotic acid was larger than the Kþ‐stimulated, Ca2þ‐dependent release, which was not blocked by diphenyl–butenyl–nipecotic acid (Belhage et al., 1993). In addition, in other neuronal preparations, this GABA release from the nonvesicular cytosolic pool of GABA can be of a considerable magnitude. Considering the increasing evidence that nonsynaptic GABA receptors may be of significant functional importance (Mody, 2001), this GABA release, which is likely to involve primarily nonsynaptic sites, could play a prominent role in tonic GABAergic inhibition (see Sonnewald et al., 2004).

6

GABA Receptors

Although electrophysiological experiments in the 1960s had clearly demonstrated hyperpolarizing actions of GABA in neuronal preparations (Curtis and Watkins, 1960; Krnjevic and Schwartz, 1967), it took many years until experimental evidence could be provided supporting the existence of specific GABA receptors, which might mediate this hyperpolarizing action of GABA. Thus, specific binding sites representing GABA receptors were demonstrated using [3H]GABA binding by Peck et al. (1973) and by the research group of Solomon Snyder (Zukin et al., 1974; Enna and Snyder, 1975; Enna et al., 1975). A discovery that a specific binding site of the benzodiazepine tranquilizing drugs was associated with the GABA receptor (Haefely et al., 1975; Mo¨hler and Okada, 1977; Squires and Bræstrup, 1977) facilitated research related to the characterization of GABA receptors enormously. This research was further advanced by the synthesis of 4,5,6,7‐tetrahydroisoxazolo[5,4‐c]pyridin‐3‐ol (THIP) and a large number of other GABA receptor agonists and antagonists (see Krogsgaard‐Larsen et al., 2002; Frølund et al., 2004). These studies ultimately led to the cloning of a large family of GABA‐receptor subunits, the combination of which into complexes of five subunits forms functional receptors, which on agonist (GABA) binding flux Cl- through the membrane (see Jensen et al., 2005). In addition to these receptors, a second class of GABA receptors was discovered by the aid of pharmacological tools by Bowery et al. (1980). Thus, it was found that certain GABA responses could be mimicked by the lipophilic GABA analog Baclophen, an effect that could not be blocked by the classical GABA receptor antagonist bicuculline (Bowery et al., 1980). This led to the nomenclature GABAA receptors for the Cl channel‐forming receptors and GABAB for this new class of receptor that did not gate an ion channel.

6.1 Ionotropic Receptors Cloning studies have led to the identification of a total of 19 subunits that can participate in the formation of five‐membered complexes, which form an ion channel that can gate Cl ions (Olsen and Macdonald, 2002; Schousboe and Waagepetersen, 2003; Jensen et al., 2005). > Table 9‐2 delineates the subunits of the ionotropic receptors, which based on pharmacological properties are referred to as GABAA and GABAC receptors (Johnston, 1997). In addition to the agonist‐ or antagonist‐binding sites, the GABAA receptors have multiple binding sites for a variety of modulators such as benzodiazepines, b‐carbolines, barbiturates, steroids, ethanol, and Zn2þ, making the receptor a multidrug target (Johnston, 1997). The subunit composition greatly influences the pharmacological profile of the receptor leading, for example, to receptors with different sensitivity of benzodiazepines (Wafford et al., 1993a, b; McKernan and Whiting, 1996). Moreover, the affinity and the efficacy of the agonist, GABA, and its structural analogs are influenced by the subunit composition of the receptor complex (Ebert et al., 1994, 1997; Krogsgaard‐Larsen et al., 1997).

219

220

9

GABA neurotransmission: An overview

. Table 9-2 Ionotropic GABA receptors: Subunit composition and basic pharmacology GABAA receptors

GABAC receptors Pharmacology

Pharmacology

Subunits

Agonists

Antagonist

Subunits

Agonists

Antagonist

a1–6 b1–3 g1–3 s e y p

GABA THIP Isoguvacine Isonipecotic acid Muscimol – –

Bicuculline Picrotoxinin – – – – –

r1–3

GABA CACA CAMP – – – –

3‐APMPA 3‐APPA – – – – –

– –

– – – –

Modified from Schousboe and Waagepetersen (2003) THIP, 4,5,6,7‐tetrahydroisoxazolo[5,4‐c]pyridin‐3‐ol; CACA, cis‐4‐aminocrotonic acid; CAMP, cis‐2‐(aminomethyl)cyclopropane‐1‐carboxylic acid; 3‐APMPA, 3‐aminopropyl(methyl)phosphinic acid; 3‐APPA, 3‐aminopropylphosphonic acid

This may be of functional importance for GABA‐mediated actions via synaptic and nonsynaptic receptors as these exhibit distinct differences with regard to subunit composition (Mody, 2001; Ebert et al., 2002).

6.2 Metabotropic Receptors The GABAB receptor, originally identified as a functional entity being activated by GABA and Baclofen in a bicuculline insensitive manner (Bowery et al., 1980), was shown to be coupled with G proteins and adenylate cyclase, the response of which leads to either an activation of Kþ channels with a subsequent increase in Kþ conductance and a hyperpolarization effect or a decrease in conductance of presynaptic Ca2þ channels resulting in a decreased transmitter release (Deisz, 1997). The cloning of this receptor (Klix and Bettler, 2002) has confirmed that it belongs to the 7TM superfamily of receptors and it has been shown to form a heteromeric complex of GABAB(1) and GABAB(2) subunits to be functionally active (Bettler and Bra¨uner‐Osborne, 2004). Recent knockout studies have shown that mice devoid of the GABAB(1) subunit exhibit epileptic seizures and such animals show lack of GABAB mediated responses (Prosser et al., 2001).

7

GABA Inactivation

All chemical neurotransmissions must induce a specific mechanism by which the receptor activation process can be terminated. In, by far, majority of transmitter systems this appears to be mediated by a combination of receptor desensitization and diffusion of the transmitter followed by high‐affinity transport into cellular elements lining the synaptic area.

7.1 Receptor Desensitization and GABA Diffusion GABAA receptors are characterized by a rapid desensitization, which is associated with a conformational change induced on agonist binding to the receptor complex (see Engblom et al., 2002). Since this is a rapid response it may well contribute significantly to the inactivation of the GABA‐induced hyperpolarization signal. On dissociation from the agonist binding site, GABA will move by diffusion in the narrow synaptic cleft before it reaches high‐affinity transport sites. Due to the short distances in the synaptic cleft, this process is also taking place at a rapid time scale.

GABA neurotransmission: An overview

9

7.2 High‐Affinity Plasma Membrane Transport High‐affinity GABA transport was originally described in spinal cord, brain slices, and homogenates (Iversen and Johnston, 1971; Iversen and Bloom, 1972; Balcar et al., 1973; Beart and Johnston, 1973), and in synaptosomes and bulk‐prepared glial cells (Henn and Hamberger, 1971; Levi and Raiteri, 1973). Moreover, autoradiographic analysis of [3H]GABA uptake in brain slices have demonstrated uptake into inhibitory nerve terminals (Bloom and Iversen, 1971; Iversen and Bloom, 1972) as well as glial elements (Ho¨kfelt and Ljungdahl, 1970). Subsequent kinetic studies of [3H]GABA transport in C‐6 glioma cells and primary cultures of astrocytes confirmed the ability of astroglial cells to perform high‐affinity GABA uptake (Hutchison et al., 1974; Schrier and Thompson, 1974; Schousboe et al., 1977). Hence, it was clear that such transport systems for GABA reside in both neuronal and glial elements, a notion confirmed by immunohistochemical analysis using specific antibodies to the cloned GABA transporters (see Schousboe and Kanner, 2002; Sarup et al., 2003b; Schousboe et al., 2004a). Studies on the uptake capacity in GABAergic neurons and in astrocytes have led to the assumption that the majority of GABA released as neurotransmitter is likely to be taken up into the nerve endings while a smaller fraction will be taken up into the surrounding glial elements (Schousboe et al., 2004a). Since the first high‐affinity GABA transporter was cloned (Guastella et al., 1990) three other transporters for GABA have been cloned (see Schousboe and Kanner, 2002). In the mouse these are referred to as GAT1, GAT2, GAT3, and GAT4 (Schousboe and Kanner, 2002), where GAT2 is identical to the betaine‐ GABA transporter 1 called BGT‐1 (Liu et al., 1993). It should be noted that since this was not called GAT2 in the rat, the current nomenclature for the GABA transporters in these two species differ in the way that GAT‐2 and GAT‐3 in the rat correspond to GAT3 and GAT4, respectively, in the mouse (Schousboe and Kanner, 2002).

7.3 Inhibitors of GABA Transporters Since inhibition of GABA transporters expressed in neurons and astrocytes provides a means of pharmacological manipulation with the GABA system, much interest has been focused on pharmacological characterization of these systems (see Volume 11 in Handbook of Neurochemistry for further details). The demonstration that the GABA analogs of restricted conformation, nipecotic acid, guvacine, and 4,5,6,7‐tetrahydroisoxazolo[4,5‐c]pyridin‐3‐ol (THPO) were specific inhibitors of GABA transport without any affinity for the GABA receptors (Krogsgaard‐Larsen and Johnston, 1975) has subsequently led to the development of a large number of GABA analogs reflecting these structures (Andersen et al., 1999, 2001; Falch et al., 1999; Knutsen et al., 1999; Sarup et al., 2003a, b; Clausen et al., 2005). Though a large number of these GABA analogs reflect the structure of nipecotic acid and guvacine, an alternative avenue was taken altering the structure of THPO leading to the development of a series of analogs based on the structure of 4‐amino‐4,5,6,7‐tetrahydro‐1,2‐benzo[d]isoxazol‐3‐ol (exo‐THPO). Such analogs have been shown to have interesting pharmacological properties suggesting astrocytic GABA transport to be an important drug target (Falch et al., 1999; Sarup et al., 2003a, b; White et al., 2002, 2005; Clausen et al., 2005).

7.3.1 Functional Implications of GABA Transport Inhibition Based on the consideration that GABAergic neurotransmission to a large extent is based on reuse of GABA taken up into the presynaptic nerve ending, it was speculated that selective inhibition of astrocytic GABA uptake might provide protection against seizures (Schousboe et al., 1983). Recent pharmacological studies using GABA transport inhibitors with different affinities for neuronal and astrocytic GABA transport have provided some evidence that anticonvulsant activity of such compounds correlates much better with the ability to inhibit astrocytic GABA uptake than the ability to inhibit neuronal GABA uptake (White et al., 2002).

221

222

9

GABA neurotransmission: An overview

It should, however, be kept in mind that Tiagabine, which is the only currently available clinically active antiepileptic drug (Kalviainen, 2002), is only marginally more potent as an inhibitor of astrocytic GABA uptake compared with neuronal uptake (Braestrup et al., 1990) and it inhibits GAT1 selectively (White et al., 2005). The recent finding that GABA analogs that inhibit GABA transport subtypes different from GAT1 (Dalby, 2003) has prompted renewed interest in this field of research leading to the discovery that inhibition of not only GAT1 but also GAT2 (BGT‐1) provides some rather interesting anticonvulsant properties. Hence, a newly synthesized exo‐THPO analog, N‐[4,4‐bis(3‐methyl‐2‐thienyl)‐3‐butenyl]‐4‐ (methylamino)‐4,5,6,7‐tetrahydrobenzo[d]isoxazol‐3‐ol (EF1502) has been shown to act synergistically with Tiagabine as an anticonvulsant, an action likely associated with inhibition of GAT2 (Clausen et al., 2005; White et al., 2005).

References Andersen KE, Lau J, Lundt BF, Petersen H, Huusfeldt PO, et al. 2001. Synthesis of novel GABA uptake inhibitors. Part 6: Preparation and evaluation of N‐Omega asymmetrically substituted nipecotic acid derivatives. Bioorg Med Chem 9: 2773-2785. Andersen KE, Sørensen JL, Huusfeldt PO, Knutsen LJ, Lau J, et al. 1999. Synthesis of novel GABA uptake inhibitors. 4. Bioisosteric transformation and successive optimization of known GABA uptake inhibitors leading to a series of potent anticonvulsant drug candidates. J Med Chem 42: 4281-4291. Awapara J, Landua AJ, Fuerst R, Seale B. 1950. Free g‐aminobutyric acid in brain. J Biol Chem 187: 35-39. Balazs R, Dahl D, Harwood JR. 1966. Subcellular distribution of enzymes of glutamate metabolism in rat brain. J Neurochem 13: 897-905. Balazs R, Machiyama Y, Hammond BJ, Julian T, Richter D. 1970. The operation of the gamma-aminobutyrate bypath of the tricarboxylic acid cycle in brain tissue in vitro. Biochem J 116: 445-461. Balcar VJ, Johnston GAR. 1973. High affinity uptake of transmitters: Studies on the uptake of L‐aspartate, GABA, L‐glutamate, and glycine in cat spinal cord. J Neurochem 20: 529-539. Beart PM, Johnston GAR. 1973. GABA uptake in rat brain slices: Inhibition by GABA analogues and by various drugs. J Neurochem 20: 319-324. Beart PM, Johnston GR. 1973. GABA uptake in rat brain slices: Inhibition by GABA analogues and by various drugs. J Neurochem 20: 319-324. Belhage B, Hansen GH, Schousboe A. 1993. Depolarization by K þ and glutamate activates different neurotransmitter release mechanisms in GABAergic neurons: Vesicular versus non‐vesicular release of GABA. Neuroscience 54: 1019-1034. Belhage B, Hansen GH, Elster E, Schousboe A. 1998. Effects of g‐aminobutyric acid (GABA) on synaptogenesis and synaptic function. Perspect Dev Neurobiol 5: 235-246.

Ben‐Ari Y. 2002. Excitatory actions of GABA during development: The nature of the nurture. Nat Rev Neurosci 3: 728-739 (Review). Ben‐Ari Y, Cherubini E, Corradetti R, Gaiarsa JL. 1989. Giant synaptic potentials in immature rat CA3 hippocampal neurons. J Physiol 416: 303-325. Bernath S. 1991. Calcium‐independent release of amino acid neurotransmitters: Fact or artifact? Prog Neurobiol 38: 57-91. Bettler B, Bra¨uner‐Osborne H. 2004. The GABAB receptor: from cloning to knockout mice. Molecular Neuropharmacology: Strategies and Methods. Schousboe A, Bra¨uner‐Osborne H, editors. Totowa, NJ: Humana Press; pp. 129-145. Bloch‐Tardy M, Rolland B, Gonnard P. 1974. Pig brain 4‐ aminobutyrate 2‐ketoglutarate transaminase. Purification, kinetics and physical properties. Biochimie 56: 823-832. Bloom FE, Iversen LL. 1971. Localizing 3H‐GABA in nerve terminals of rat cerebral cortex by electron microscopic autoradiography. Nature 229: 628-630. Bowery NG, Hill DR, Hudson AL, Doble A, Middlemiss DN, et al. 1980. (-)Baclofen decreases neurotransmitter release in the mammalian CNS by an action at a novel GABA receptor. Nature 283: 92-94. Braestrup C, Nielsen EB, Sonnewald U, Knutsen LJS, Andersen KE, et al. 1990. (R)‐N‐[4,4‐Bis(3‐methyl‐2‐ thienyl)but‐3‐en‐1‐yl]nipecotic acid binds with high affinity to the brain g‐aminobutyric acid uptake carrier. J Neurochem 54: 639-647. Cash C, Maitre M, Ciesielski L, Mandel P. 1974. Purification and partial characterisation of 4‐aminobutyrate 2‐ketoglutarae transaminase from human brain. FEBS Lett 47: 199-203. Cherubini E, Gaiarsa JL, Ben‐Ari Y. 1991. GABA: An excitatory transmitter in early postnatal life. TINS 14: 515-519. Clausen RP, Moltzen EK, Perregaard J, Lenz SM, Sanchez C, et al. 2005. Selective inhibitors of GABA uptake: Synthesis and molecular pharmacology of 3‐hydroxy‐4‐N‐methylamino‐4,5,6,7‐tetrahydro‐1,2‐benzo[d]isoxazole analogues. Bioorg Med Chem 13: 895-908.

GABA neurotransmission: An overview Curtis DR, Johnston GAR. 1974. Amino acid transmitters in the mammalian central nervous system. Ergeb Physiol 69: 97-188. Curtis DR, Watkins JC. 1960. The excitation and depression of spinal neurones by structurally related amino acids. J Neurochem 6: 117-141. Dalby NO. 2003. Inhibition of gamma‐aminobutyric acid uptake: Anatomy, physiology and effects against epileptic seizures. Eur J Pharmacol 479: 127-137. Deisz RA. 1997. Electrophysiology of GABAB receptors. The GABA Receptors, 2nd edn. Enna SJ, Bowery NG, editors. Totowa, NJ: Humana Press; pp. 157–208. Ebert B, Storustovu S, Mortensen M, Frølund B. 2002. Characterization of GABA(A) receptor ligands in the rat cortical wedge preparation: Evidence for action at extrasynaptic receptors? Br J Pharmacol 137: 1-8. Ebert B, Wafford KA, Whiting PJ, Krogsgaard‐Larsen P, Kemp JA. 1994. Molecular pharmacology of gamma‐ aminobutyric acid type A receptor agonists and partial agonists in oocytes injected with different alpha, beta, and gamma receptor subunit combinations. Mol Pharmacol 46: 957-963. Ebert B, Thompson SA, Saounatsou K, McKernan R, Krogsgaard‐Larsen P, et al. 1997. Differences in agonist/ antagonist binding affinity and receptor transduction using recombinant human gamma‐aminobutyric acid type A receptors. Mol Pharmacol 52: 1150-1156. Engblom AC, Carlson BX, Olsen RW, Schousboe A, Kristiansen U. 2002. Point mutation in the first transmembrane region of the beta2 subunit of the gamma‐aminobutyric acid type A receptor alters desensitization kinetics of gamma‐aminobutyric acid‐ and anesthetic‐induced channel gating. J Biol Chem 277: 17438-17447. Enna SJ, Kuhar MJ, Snyder SH. 1975. Regional distribution of postsynaptic receptor binding for gamma-aminobutyric acid (GABA) in monkey brain. Brain Res 93: 168-174. Enna SJ, Snyder SH. 1975. Properties of g‐aminobutyric acid (GABA) receptor binding in rat brain synaptic membrane fractions. Brain Res 100: 81-97. Falch E, Perregaard J, Frølund B, Søkilde B, Buur A, et al. 1999. Selective inhibitors of glial GABA uptake: Synthesis, absolute stereochemistry and pharmacology of the enantiomers of 3‐hydroxy‐4‐amino‐4,5,6,7‐tetrahydro‐1,2‐benzisoxazole (exo‐THPO) and analogues. J Med Chem 42: 5402-5414. Fiszman ML, Schousboe A. 2004. Role of calcium and kinases on the neurotrophic effect induced by g‐aminobutyric acid. J Neurosci Res 76: 435-441. Frølund B, Jørgensen AT, Liljefors T, Mortensen M, Krogsgaard‐Larsen P. 2004. Characterization of the GABAA receptor recognition site through ligand design

9

and pharmacophore modelling. Molecular Neuropharmacology: Strategies and Methods. Schousboe A, Bra¨uner‐Osborne H, editors. Totowa, NJ: Humana Press; pp. 113-127. Gram L, Larsson OM, Johnsen AH, Schousboe A. 1988. Effects of valproate, vigabatrin and aminooxyacetic acid on release of endogenous and exogenous GABA from cultured neurons. Epilepsy Res 2: 87-95. Guastella J, Nelson N, Nelson H, Czyzyk L, Keynan S, et al. 1990. Cloning and expression of a rat brain GABA transporter. Science 249: 1303-1306. Gutierrez R. 2003. The GABAergic phenotype of the glutamatergic granule cells of the dentate gyrus. Prog Neurobiol 71: 337-358. Haefely W, Kuskar A, Mo¨hler H, Pieri L, Polc P, et al. 1975. Possible involvement of GABA in the central actions of benzodiazepines. Adv Biochem Psychopharmacol 14: 131-152. Henn FA, Hamberger A. 1971. Glial cell function: Uptake of transmitter substances. Proc Natl Acad Sci USA 68: 2686-2690. Hertz L, Juurlink BHJ, Hertz E, Fosmark H, Schousboe A. 1989b. Preparation of primary cultures of mouse (rat) astrocytes. In: A Dissection and Tissue Culture Manual for the Nervous System. Shahar A, De Vellis J, Vernadakis A, Haber B, editors. New York: Alan R, Liss: pp. 105-108. Hertz E, Yu ACH, Hertz, L, Juurlink BHJ, Schousboe A. 1989a. Preparation of primary cultures of mouse cortical neurons. In: A Dissection and Tissue Culture Manual for the Nervous System. Shahar A, De Vellis J, Vernadakis A, Haber B, editors. New York: Alan R, Liss: 183-186. Ho¨kfelt T, Ljungdahl A˚. 1970. Cellular localization of labelled gamma‐aminobutyric acid (3H‐GABA) in rat cerebellar cortex: An autoradiographic study. Brain Res 22: 391-396. Hutchison HT, Werrbach K, Cance C, Haber B. 1974. Uptake of neurotransmitters by clonal lines of astrocytoma and neuroblastoma in culture. I. Transport of g‐aminobutyric acid. Brain Res 66: 265-274. Iadarola MJ, Gale K. 1980. Evaluation of increases in nerve terminal‐dependent vs nerve terminal‐independent compartments of GABA in vivo. Brain Res Bull 5(Suppl.)2: 13-19. Iversen LL, Bloom FE. 1972. Studies on the uptake of 3H‐ GABA and 3H‐glycine in slices and homogenates of rat brain and spinal cord by electron microscopic autoradiography. Brain Res 41: 131-143. Iversen LL, Johnston GAR. 1971. GABA uptake in rat central nervous system: Comparison of uptake in slices and homogenates and the effects of some inhibitors. J Neurochem 18: 1939-1950.

223

224

9

GABA neurotransmission: An overview

Jensen ML, Schousboe A, Ahring P. 2005. Charge selectivity of the Cys‐loop family of ligand gated ion channels. J Neurochem 92: 217-225. John RA, Fowler LJ. 1976. Kinetic and spectral properties of rabbit brain 4‐aminobutyrate aminotransferase. Biochem J 155: 645-651. Johnston GAR. 1997. Molecular biology, pharmacology, and physiology of GABAC receptors. The GABA Receptors, 2nd edn. Enna SJ, Bowery NG, editors. Totowa, NJ: Humana Press; pp. 297-323. Kalviainen R. 2002. Tiagabine. Clinical Efficacy and Use in Epilepsy. Levy RH, Mattson RH, Meldrum BS, Perucca E, editors. Philadelphia: Lippincott Williams & Wilkins; pp. 699-704. Klix N, Bettler B. 2002. Molecular structure of the GABAB receptors. Glutamate and GABA Receptors and Transporters. Egebjerg J, Schousboe A, Krogsgaard‐Larsen P, editors. London, UK: Taylor & Francis; pp. 277-286. Knutsen LJ, Andersen KE, Lau J, Lundt BF, HenryRF, et al. 1999. Synthesis of novel GABA uptake inhibitors. 3. Diaryloxime and diarylvinyl ether derivatives of nipecotic acid and guvacine as anticonvulsant agents. J Med Chem 42: 3447-3462. Krnjevic K, Schwartz S. 1967. The action of g‐aminobutyric acid on cortical neurons. Exp Brain Res 3: 320-336. Krogsgaard‐Larsen P, Johnston GAR. 1975. Inhibition of GABA uptake in rat brain slices by nipecotic acid, various isoxazoles and related compounds. J Neurochem 25: 797-802. Krogsgaard‐Larsen P, Frølund B, Ebert B. 1997. GABAA receptor agonists, partial agonists, and antagonists. The GABA Receptors, 2nd edn. Enna SJ, Bowery NG, editors. Totowa, NJ: Humana Press; pp. 37-81. Krogsgaard‐Larsen P, Frølund B, Kristiansen U, Ebert B. 2002. Ligands for the GABAA receptor complex. Glutamate and GABA Receptors and Transporters. Egebjerg J, Schousboe A, Krogsgaard‐Larsen P, editors. London, UK: Taylor & Francis; pp. 236-274. Kvamme E, Torgner IA, Roberg B. 2001. Kinetics and localization of brain phosphate activated glutaminase. J Neurosci Res 66: 951-958. Larsson OM, Drejer J, Kvamme E, Svenneby G, Hertz L, Schousboe A. 1985. Ontogenetic development of glutamate and GABA metabolizing enzymes in cultured cerebral cortex interneurons and in cerebral cortex in vivo. Int J Devl Neurosci 3: 177-185. Larsson OM, Gram L, Schousboe I, Schousboe A. 1986. Differential effect of gamma‐vinyl GABA and valproate on GABA‐transaminase from cultured neurons and astrocytes. Neuropharmacology 25: 617-625. Levi G, Raiteri M. 1973. Detectability of high and low affinity uptake systems for GABA and glutamate in rat brain slices and synaptosomes. Life Sci 12: 81-88.

Lippert B, Metcalf BW, Jung MJ, Casara P. 1977. 4‐Aminohex‐ 5‐enoic acid, a selective catalytic inhibitor of 4‐aminobutyric acid aminotransferase in mammalian brain. Eur J Biochem 74: 441-445. Liu QR, Lopez‐Corcuera B, Mandiyan S, Nelson H, Nelson N. 1993. Molecular characterization of four pharmacologically distinct g‐aminobutyric acid transporters in mouse brain. J Biol Chem 268: 2104-2112. Machiyama Y, Bala´zs R, Hammond BJ, Julian T, Richter D. 1970. The metabolism of g‐aminobutyrate and glucose in potassium ion‐stimulated brain tissue in vitro. Biochem J 116: 469-481. Maitre M, Ciesielski L, Cash C, Mandel P. 1975. Purification and studies on some properties of the 4‐aminobutyrate: 2‐ oxoglutarate transaminase from rat brain. Eur J Biochem 52: 157-169. Martin DL, Rimval K. 1993. Regulation of g‐aminobutyric acid synthesis in the brain. J Neurochem 60: 395-407. Matsuda T, Wu J‐Y, Roberts E. 1973. Electrophoresis of glutamic acid decarboxylase (EC 4.1.1.15) from mouse brain in sodium dodecyl sulphate polyacrylamide gels. J Neurochem 21: 167-172. McKernan RM, Whiting PJ. 1996. Which GABAA‐receptor subtypes really occur in the brain? Trends Neurosci 19: 139-143. McLaughlin BJ, Wood JG, Saito K, Barber R, Vaughn JE, et al. 1974. The fine structural localization of glutamate decarboxylase in synaptic terminals of rodent cerebellum. Brain Res 76: 377-391. Mo¨hler H, Okada T. 1977. Benzodiazepine receptors: Demonstration in the central nervous system. Science 198: 849-851. Mody I. 2001. Distinguishing between GABA(A) receptors responsible for tonic and phasic conductances. Neurochem Res 26: 907-913. Olsen RW, Macdonald RL. 2002. GABAA receptor complex: Structure and complex. Glutamate and GABA Receptors and Transporters. Egebjerg J, Schousboe A, Krogsgaard‐ Larsen P, editors. London, UK: Taylor & Francis; pp. 202-235. Otsuka M. 1996. Establishment of GABA as an inhibitory neurotransmitter at crustacean neuromuscular junction and in the mammalian central nervous system. GABA: Receptors, Transporters and Metabolism. Tanaka C, Bowery NG, editors. Basel: Birkha¨user Verlag; pp. 1-6. Palaiologos G, Hertz L, Schousboe A. 1988. Evidence that aspartate amino transferase activity and ketodicarboxylate carrier function are essential for biosynthesis of transmitter glutamate. J Neurochem 51: 317-320. Peck EJ, Jr, Schaeffer JM, Clark JH. 1973. g‐Aminobutyric acid, bicuculline, and post‐synaptic binding sites. Biochem Biophys Res Commun 52: 394-400.

GABA neurotransmission: An overview Pin JP, Bockaert J. 1989. Two distinct mechanisms, differentially affected by excitatory amino acids, trigger GABA release from fetal mouse striatal neurons in primary culture. J Neurosci 9: 648-656. Prosser HM, Gill CH, Hirst WD, Grau E, Robbins M, et al. 2001. Epileptogenesis and enhanced prepulse inhibition in GABAB1‐deficient mice. Mol Cell Neurosci 17: 1059-1070. Reubi JC, Van den Berg CJ, Cue´nod M. 1978. Glutamine as precursor for the GABA and glutamate transmitter pools. Neurosci Lett 10: 171-174. Ribak CE, Vaughn JE, Saito K, Barber R, Roberts E. 1976. Immunocytochemical localization of glutamate decarboxylase in rat substantia nigra. Brain Res 116: 287-298. Roberts E, Frankel S. 1950. g‐Aminobutyric acid in brain: Its formation from glutamic acid. J Biol Chem 187: 55-63. Roberts E, Simonsen DG. 1963. Some properties of L‐glutamic decarboxylase in mouse brain. Biochem Pharmacol 12: 113-134. Roberts E, Baxter CF, Van Harreveld A, Wiersma CAG, Adey WR, et al. (eds) 1960. Inhibition in the Nervous System and Gamma‐aminobutyric Acid. Oxford: Pergamon Press. Saito K, Barber R, Wu J‐Y, Matsuda T, Roberts E, et al. 1974. Immunohistochemical localization of glutamate decarboxylase in rat cerebellum. Proc Natl Acad Sci USA 71: 269-273. Sarup A, Larsson OM, Schousboe A. 2003b. GABA transporters and GABA‐transaminase as drug targets. Curr Drug Targ CNS Neurol Dis 2: 269-277. Sarup A, Larsson OM, Bolvig T, Frølund B, Krogsgaard‐ Larsen P, et al. 2003a. Effects of 3‐hydroxy‐4‐amino‐ 4,5,6,7‐tetrahydro‐1,2‐benzisoxazol (exo‐THPO) and its N‐substituted analogs on GABA transport in cultured neurons and astrocytes and by the four cloned mouse GABA transporters. Neurochem Int 43: 445-451. Schousboe A, Kanner B. 2002. GABA transporters: Functional and pharmacological properties. Glutamate and GABA Receptors and Transporters. Egebjerg J, Schousboe A, Krogsgaard‐Larsen P, editors. London, UK: Taylor & Francis Publ.; pp. 337-349. Schousboe A, Waagepetersen HS. 2003. GABA. Encyclopedia of Neuroscience. Adelman G, Smith BH, editors. New York, USA: Elsevier Science; pp. 166-176.www.neuroscion.com Schousboe A, Hertz L, Svenneby G. 1977. Uptake and metabolism of GABA in astrocytes cultured from dissociated mouse brain hemispheres. Neurochem Res 2: 217-229. Schousboe A, Larsson OM, Seiler N. 1986. Stereoselective uptake of the GABA‐transaminase inhibitors gamma‐vinyl GABA and gamma‐acetylenic GABA into neurons and astrocytes. Neurochem Res 11: 1497-1505. Schousboe A, Lisy V, Hertz L. 1976. Postnatal alterations in effects of potassium on uptake and release of glutamate and GABA in rat brain cortex slices. J Neurochem 26: 1023-1027.

9

Schousboe A, Wu J‐Y, Roberts E. 1973. Purification and characterization of the 4‐aminobutyrate‐2‐ketogluterate transaminase from mouse brain. Biochemistry 12: 2868-2873. Schousboe A, Wu J‐Y, Roberts E. 1974. Subunit structure and kinetic properties of 4‐aminobutyrate‐2‐ketoglutarate transaminase from mouse brain. J Neurochem 23: 11891195. Schousboe A, Larsson OM, Sarup A, White HS. 2004c. Role of the betaine/GABA transporter (BGT‐1/GAT2) for the control of epilepsy. Eur J Pharmacol 500: 281-287. Schousboe A, Larsson OM, Wood JD, Krogsgaard‐Larsen P. 1983. Transport and metabolism of GABA in neurons and glia: Implications for epilepsy. Epilepsia 24: 531-538. Schousboe A, Meier E, Drejer J, Hertz L. 1989. Preparation of primary cultures of mouse (rat) cerebellar granule cells. In: A Dissection and Tissue Culture Manual for the Nervous System. Shahar A, De Vellis J, Vernadakis A, Haber B, editors. New York: Alan R, Liss: pp. 203-206. Schousboe A, Sarup A, Larsson OM, White HS. 2004b. GABA transporters as drug targets for modulation of GABAergic activity. Biochem Pharmacol 68: 1557-1563. Schousboe A, Sarup A, Bak LK, Waagepetersen HS, Larsson OM. 2004a. Role of astrocytic transport processes in glutamatergic and GABAergic neurotransmission. Neurochem Int 45: 512-527. Schousboe I, Bro B, Schousboe A. 1977. Intramitochondrial localization of the 4-aminobutyrate-2-ketoglutarate transaminase from ox brain. Biochem J 162: 303-307. Schrier BK, Thompson EJ. 1974. On the role of glial cells in the mammalian nervous system. Uptake, excretion, metabolism of putative neurotransmitters by cultured glial tumor cells. J Biol Chem 249: 1769-1780. Sloviter M, Dichter MA, Rachinsky TL, Dean E, Goodman JH, et al. 1996. Basal expression and induction of glutamate decarboxylase and GABA in excitatory granule cells of the rat and monkey hippocampal dentate gyrus. J Comp Neurol 373: 593-618. Soghomonian JJ, Martin DL. 1998. Two isoforms of glutamate decarboxylase: Why? Trends Pharmacol Sci 19: 500-505. Sonnewald U, Syversen T, Schousboe A, Waagepetersen HS, Aschner M. 2007. Actions of toxins on cerebral metabolism at the cellular level. In: Handbook of Neurochemistry and Molecular Biology, Vol. 5. Lajtha A, editor. Springer-Verlag Berlin Heidelberg, pp. 569-585. Sonnewald U, Olstad E, Qu H, Babot Z, Cristo`fol R, et al. 2004. First direct demonstration of extensive GABA synthesis in mouse cerebellar neuronal cultures. J Neurochem 91: 796-803. Sprince H, Parker CM, Josephs JA, Jr, Magazino J. 1969. Convulsant activity of homocysteine and other short‐chain mercaptoacids: Protection therefrom. Ann N Y Acad Sci 166: 323-325.

225

226

9

GABA neurotransmission: An overview

Squires C, Bræstrup RF. 1977. Specific benzodiazepine receptors in rat brain characterized by high‐affinity (3H)diazepam binding. Proc Natl Acad Sci USA 74: 3805-3809. Tapia R. 1975. Biochemical pharmacology of GABA in CNS. Handbook of Psychopharmacology, Vol. 4. Iverson LL, Iverson SD, Snyder SH, editors. New York: Plunum Publ. Corp.; pp. 1-58. Tapia R, Meza‐Ruiz G. 1975. Differences in some properties of newborn and adult brain glutamate decarboxylase. J Neurobiol 6: 171-181. Tapia R, Pasantes‐Morales H, Taborda E, Pe´rez de la Mora M. 1975. Seizure susceptibility in the developing mouse and its relationship to glutamate decarboxylase in pyridoxal phosphate in brain. J Neurobiol 6: 159-170. Udenfriend S. 1950. Identification of g‐aminobutyric acid in brain by the isotope derivative method. J Biol Chem 187: 65-69. Waagepetersen HS, Sonnewald U, Schousboe A. 1999. The GABA paradox: Multiple roles as metabolite, neurotransmitter, and neurodifferentiative agent. J Neurochem 73: 1335-1342. Waagepetersen HS, Sonnewald U, Schousboe A. 2003. Compartmentation of glutamine, glutamate and GABA metabolism in neurons and astrocytes: Functional implications. Neuroscientist 9: 398-403. Waagepetersen HS, Sonnewald U, Gegelashvili G, Larsson OM, Schousboe A. 2001. Metabolic distinction between vesicular and cytosolic GABA in cultured GABAergic neurons using 13C MRS. J Neurosci Res 63: 347-355. Wafford KA, Whiting PJ, Kemp JA. 1993a. Differences in affinity and efficacy of benzodiazepine receptor ligands on recombinant GABAA receptor subtypes. Mol Pharmacol 43: 240-244. Wafford KA, Bain CJ, Whiting PJ, Kemp JA. 1993b. A functional comparison of the role of g subunits in recombinant g‐aminobutyric acid type A/benzodiazepine receptors. Mol Pharmacol 43: 437-442. Westergaard N, Sonnewald U, Petersen SB, Schousboe A. 1995. Glutamate and glutamine metabolism in cultured GABAergic neurons studied by 13C NMR spectroscopy:

Evidence for compartmentation and mitochondrial heterogeneity. Neurosci Lett 185: 24-28. White HS, Sarup A, Bolvig T, Kristensen AS, Petersen G, et al. 2002. Correlation between anticonvulsant activity and inhibitory action on glial GABA uptake of the highly selective mouse GAT1 inhibitor 3‐hydroxy‐4‐ amino‐4,5,6,7‐tetrahydro‐1,2‐benzisoxazole(exo‐THPO)and its N‐alkylated analogs. J Pharmacol Exp Ther 302: 636-644. White HS, Watson WP, Hansen S, Slough S, Sarup A, et al. 2005. First demonstration of a functional role for CNS betaine/GABA transporter (mGAT2) based on synergistic anticonvulsant action among inhibitors of mGAT1 and mGAT2. J Pharmacol Exp Ther 312: 866-874. Wood JD, Kurylo E, Tsui SK. 1981. Interactions of di‐n‐ propylacetate, gabaculine, and aminooxyacetic acid: Anticonvulsant activity and the gamma‐aminobutyrate system. J Neurochem 37: 1440-1447. Wu J‐Y, Roberts E. 1974. Properties of brain L‐glutamate decarboxylase: Inhibition studies. J Neurochem 23: 759-767. Wu J‐Y, Matsuda T, Roberts E. 1973. Purification and characterization of glutamate decarboxylase from mouse brain. J Biol Chem 248: 3039-3034. Wu J‐Y, Moss LG, Chude O. 1978. Distribution and tissue specificity of 4‐aminobutyrate‐2‐oxoglutarate aminotransferase. Neurochem Res 3: 207-219. Yu ACH, Hertz L. 1983. Metabolic sources of energy in astrocytes. Glutamine, Glutamate, and GABA in the Central Nervous System. Hertz L, Kvamme E, McGeer EG, Schousboe A, editors. New York: Alan R. Liss, Inc.; pp. 431-438. Zieminska E, Hilgier W, Waagepetersen HS, Hertz L, Sonnewald U, et al. 2004. Analysis of glutamine accumulation in rat brain mitochondria in the presence of a glutamine uptake inhibitor, histidine, reveals glutamine pools with a distinct access to deamidation. Neurochem Res 29: 2121-2123. Zukin SR, Young AB, Snyder SH. 1974. Gamma‐aminobutyric acid binding to receptor sites in the rat central nervous system. Proc Natl Acad Sci USA 71: 4802-4807.

10

ATP‐Mediated Signaling in the Nervous System

B. Sperla´gh

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 228

2

Synthesis, Utilization, and Storage of ATP in the Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229

3

The Release of ATP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231

4

The Extracellular Inactivation of ATP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233

5

ATP Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235

6

The Fast Transmitter Action of ATP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238

7

The Presynaptic Modulatory Role of ATP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238

8

The Role of ATP in Glia–Neuron and Glia–Glia Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241

9

The Role of ATP in Sensory Transmission and in the Generation of Pain . . . . . . . . . . . . . . . . . . . . . 243

10 The Role of P2 Receptors in Behavioral Paradigms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 244 11 ATP as a Neuroimmunomodulator . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 244 12 Involvement of ATP Receptors CNS Diseases and their Potential Therapeutic Exploitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 245 13 Conclusions and Future Avenues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 246

#

2008 Springer ScienceþBusiness Media, LLC.

228

10

ATP‐mediated signaling in the nervous system

Abstract: ATP functions as a ubiquitous signaling substance in neuronal and nonneuronal tissues. It is synthesized and stored in the nerve terminals, glial cells, and postsynaptic target cells, and is released in response to neuronal activity and a variety of other stimuli (activation of pre‐ and postsynaptic receptors, metabolic distress, inflammation, hypoosmotic stimuli, and cellular damage). ATP acts on various subtypes of ionotropic P2X and metabotropic P2Y receptors, which are widely distributed in the nervous system. P2X receptors mediate the fast transmitter action of ATP, which has been identified in a number of central and peripheral synapses. In addition, ATP modulates synaptic transmission pre‐ and postsynaptically, both in positive and negative directions via activation of P2X and P2Y receptors, respectively. Moreover, ATP acts as a transmitter not only in neuron–neuronal and neuro‐effector synapses, but also transmits signals between glial cells and neurons and within glial networks. Rapidly emerging data indicate that ATP plays an important role in sensory systems, that is, in the processing of the pain, in mechano‐ and chemosensory transduction and in the microglial response to inflammatory challenge. Finally, ATP might also act as a pathological mediator in acute and chronic neurodegeneration and in the following repair process. List of Abbreviations: Aβ, amyloid beta peptide; ABC, ATP binding cassette; ADP, adenosine 50 -diphosphate; 2-AG, 2-arachidonoylglycerol; AMP, adenosine 50 -monophosphate; AMPA, α-amino-5-hydroxy-3methyl-4-isoxazole propionic acid; Ap4A, diadenosine tetraphosphate; Ap5A, diadenosine pentaphosphate; ATP, adenosine 50 -triphosphate; CNS, central nervous system; CNT, concentrative nucleoside transporter; COX-2, cyclooxigenase-2; EEG, electroencephalography; E-NPPs, ecto-nucleotide pyrophosphatases; ENT, equilibrative nucleoside transporter; E-NTPDases, ectonucleoside triphosphate diphospho-hydrolases; ERK/JNK, extracellular signal regulated protein kinase; GABA, γ-amino-butyric acid; GPI, glycosylphosphatidyl inositol; GTP, guanosine 50 -triphosphate; HPLC, high performance liquid chromatography; IFNγ, interferon–γ; IL-1α, interleukin-1α; IL-1β, interleukin-1β; IL-6, interleukin-6; IL-18, interleukin-18; IMP, inosine monophosphate; iNOS, inducible nitric oxide synthase; LPS, bacterial lypopolisaccharide; mRNA, messenger RNA; NBMPR, nitrobenzylthioinosine; NMDA, N-methyl-D-aspartate; NO, nitric oxide; NTS, nucleus tractus solitarii; 6-OHDA, 6-hydroxidopamine; p38MAPK, p38 mitogen activated protein kinase; PNS, peripheral nervous system; PPADS, pyridoxal-phosphate-6-azophenyl-20 ,40 disulphonic acid; ROI, reactive oxygen intermediates; siRNA, small interfering RNA; TNFα, tumor necrosis factor α; UDP, uridine 50 -diphosphate; UTP, uridine 50 -triphosphate

1 Introduction ATP is well known as the universal energy currency of living cells. The first observation suggesting that ATP also plays an important signaling role, besides its central role in the cellular energy homeostasis, was reported by Drury and Szent‐Gyo¨rgyi in 1929. They observed that adenylyl compounds, including ATP, have a profound effect on heart rate and cardiovascular function (Drury and Szent-Gyo¨rgyi, 1929). This idea was rediscovered by Geoffrey Burnstock in the early 1970s, who proposed the purinergic nerve hypothesis, suggesting that ATP acts as a specific neurotransmitter in the nervous system (Burnstock, 1972). In the past three decades, this concept has gained solid experimental proof in a number of synapses of the PNS and the CNS. Moreover, it also turned out that extracellular ATP has a more versatile function in the neuronal information processing than a classical neurotransmitter, participating in pre‐ and postsynaptic neuromodulation, glia–neuron and glia–glia interactions, and neuroimmunomodulation. The purinergic signaling system also offers a number of intervention sites for therapeutic exploitation in diseases of the nervous system, which potentially could be used for drug development. There are a number of excellent review articles on different aspects of the purinergic signaling system. In this chapter, a summary is given on the current knowledge on the synthesis, storage, release, action, and inactivation of extracellular ATP in the nervous system and more detailed information can be found in specialized reviews (Sperla´gh and Vizi, 1996; Abbracchio and Burnstock, 1998; Ralevic and Burnstock, 1998; Illes et al., 2000; North and Surprenant, 2000; Sperlagh and Vizi, 2000; Zimmermann, 2000, 2001; Cunha and Ribeiro, 2000a; Cunha, 2001; Khakh, 2001; Khakh et al., 2001; Kennedy et al., 2003; Illes and Ribeiro, 2004; Kennedy, 2005; Koles et al., 2005; Franke and Illes, 2006).

ATP‐mediated signaling in the nervous system

10

It is also important to note that although the source, release, and extracellular fate of ATP and its extracellular breakdown product, adenosine, is tightly coupled, they form separate signaling systems at the level of receptors. Therefore, the present chapter focuses only on the signaling functions mediated by ATP and other nucleotides, and those mediated by adenosine and other nucleosides will be detailed in Chapter 15.

2 Synthesis, Utilization, and Storage of ATP in the Nervous System Since ATP is ubiquitous, all metabolically active cells, including neurons, are able to synthesize ATP. The majority of ATP under normal metabolic conditions are formed from ADP by oxidative phosphorylation, in the mitochondria. In addition, ATP is also generated in a minor amount in the glycolytic pathway and in the citric acid cycle (> Figures 10-1 and > 10-2). The direct precursor of ATP formation is ADP, from which ATP is generated depending on the metabolic demand. This is the so‐called respiratory control, as the mitochondrial ATP production is coupled to the respiratory chain and is driven by the actual ADP concentration of the cell. The adenine ring of the ATP molecule is synthesized during the multiple steps of de novo purine biosynthesis from phosphoribosyl pyrophosphate, resulting in inosine monophosphate (IMP) production. IMP is then transaminated to AMP and then directly phosphorylated to ADP, serving as a substrate for mitochondrial oxidative phosphorylation (> Figure 10-1). However, the de novo purine biosynthesis is an energy‐consuming process, therefore purine salvage mechanisms also exist, which enable nerve terminals to use purines from exogenous sources. Among them, the most important is that nerve terminals take up adenosine via the nucleoside transport systems (see later) and the adenosine kinase enzyme converts adenosine to AMP, which then enters the reactions detailed earlier. Taken together, the activity of all these

. Figure 10-1 The chemical formula of adenosine 50 ‐triphosphate (ATP). The majority of ATP under normal metabolic conditions is formed from ADP, in the mitochondria by oxidative phosphorylation. In addition, ATP is also generated in a minor amount in the glycolytic pathway and in the citric acid cycle. The adenine ring of the ATP molecule is synthesized during the multiple steps of de novo purine biosynthesis from phosphoribosyl pyrophosphate, resulting in IMP production. IMP is then transaminated to AMP and then directly phosphorylated to ADP, serving as a substrate for mitochondrial oxidative phosphorylation. The ribose moiety is synthesized in the pentose phosphate pathway

229

230

10

ATP‐mediated signaling in the nervous system

. Figure 10-2 Storage, release, and interconversion of ATP and adenosine in the synapse. Adenosine is taken up to the nerve terminal by the bidirectional, ENT transporter. Subsequently, it is rephosphorylated to AMP and ADP by adenosine kinase and adenylate kinase enzymes, respectively. The major pathway of ATP production is the mitochondrial oxidative phosphorylation, which converts ADP to ATP according to the energy demand of the cell, that is, to the actual amount of ADP. ATP is then transported out from the mitochondrion and is taken up by synaptic vesicles and released by vesicular exocytosis. In addition, ATP could be also released from the cytoplasm of pre‐ and postsynaptic cells. If ADP and AMP are accumulated in the cytoplasm, due to insufficient oxidative phosphorylation, cytosolic 50 ‐nucleotidase enzyme produces adenosine from AMP, which could leave the cell via the nucleoside carrier. ATP and ADP are hydrolyzed to AMP in the extracellular space by E‐NTPDase, E‐NPP, and by the alkaline phosphatase enzymes, and AMP is hydrolyzed to adenosine by the ecto‐50 ‐nucleotidase enzyme. In addition, there is also catalytic activity in the extracellular space for an ATP regenerating, reverse process, that is, to rephosporylate and interconvert nucleosides and nucleotides to ATP by the ecto‐ adenylate kinase enzyme. Finally, adenosine could be deaminated either intra‐ or extracellularly by adenosine deaminase enzyme giving rise to formation of inosine

reactions results in approximately 10 mM ATP concentration in the cytoplasm under normal metabolic conditions. The majority of ATP produced by the nerve terminal are used to fuel energy‐consuming cellular functions, and among them, the most important are (i) the maintenance of resting membrane potential by the Naþ/Kþ pump, (ii) the function of the other ion‐pumping mechanisms, such as the Ca2þ pumps of the plasma membrane and the mitochondria, (iii) the synthesis of neurotransmitters, receptors, ion channels, transporters, and other signaling proteins and molecules, like G proteins, protein kinases, GTP, etc., (iv) the build‐up of the vesicular proton gradient by the vacuolar Hþ ATPase, and (v) the steps of the

ATP‐mediated signaling in the nervous system

10

exocytosis itself. Nevertheless, cytoplasmic ATP is also available for intra‐ and extracellular signaling process, and ATP is also taken up and stored in synaptic vesicles (Sperla´gh, 1996) (> Figure 10-2). ATP is the known constituent of cholinergic, noradrenergic, and serotonergic vesicles and is probably present in other types of synaptic vesicles as well. In cholinergic and noradrenergic vesicles, the amount of ATP is usually outnumbered by its cotransmitter mate. Thus, storage ratios of 3:1 to 50:1 were established, depending upon the type of vesicles, corresponding to about 1–200 mM concentration of ATP inside the vesicle. However, ATP content may differ in individual vesicles and can change under various conditions, for example, upon different patterns of neuronal activity. Although direct proof on their presence is yet to be demonstrated, it is also possible that purely ATP‐containing vesicles exist. In addition to ATP, other nucleotides are also stored in synaptic vesicles, such as ADP, AMP, UTP, Ap4A, Ap5A, and guanine nucleotides, which are thought to play role as signaling substances (Zimmermann, 2001). Although their concentration is less than that of ATP, it is still relatively high, that is, in millimolar range, and enough to serve as a pool for their release.

3 The Release of ATP The participation of ATP in the intercellular communication presumes its release to the extracellular space upon physiological and/or pathological stimuli. Indeed, a wide variety of stimuli are known to release ATP such as (i) electrical or chemical depolarization of nerve terminals, (ii) activation of cell surface receptors, (iii) mechanical stimuli, (iv) hypoxia/hypoglycemia/ischemia and the consequent cellular energy deprivation, (v) hypoosmotic challenge, (vi) inflammatory stimuli, and (vii) cellular damage. The release of ATP upon neuronal activity was demonstrated for the first time by Holton (1959) using antidromic stimulation of sensory nerves. Since then, the stimulation‐dependent release of endogenous ATP has been reported from a wide variety of in vitro brain slice and nerve terminal preparations and from isolated tissues innervated by the peripheral and autonomic nervous system, using electrical field stimulation, direct stimulation of specific neuronal pathways or chemical depolarization (cf. Sperla´gh and Vizi, 1996; Sperlagh and Vizi, 2000). The most frequently used techniques capable of detecting endogenous ATP release are the luciferin–luciferase assay and high performance liquid chromatography (HPLC) coupled with ultraviolet or fluorescent detection. The former method is preferable for rapid determination of ATP levels and is advantageous because of its high sensitivity, specificity, and simplicity. On the other hand, the HPLC method offers the opportunity to separate and identify all purine compounds released to the extracellular space. In addition, preloading the tissues with [3H]adenosine or [3H]adenine and then measuring the tritium efflux is also often used to detect [3H]purine release; however, the released radioactive label in this case is a mixture of released purines and their metabolic degradation products, and therefore it is necessary to analyze the composition of radioactive label by HPLC subsequently to identify the released purine compounds. Recently, a technical breakthrough was achieved in extracellular purine analysis by the introduction of the enzyme‐based microelectrode biosensor technique, which is able to follow ATP and adenosine release in a real‐time scale, both in vitro and in vivo. Using this method, stimulation‐dependent physiological ATP release was demonstrated during locomotor activity from spinal networks (Llaudet et al., 2005), during the hypoxic ventilatory response from the carotid body and in response to the elevation of blood pCO2 from the chemosensitive region of the medulla oblongata (Spyer et al., 2004; Gourine et al., 2005a, b), and from the retinal pigment epithelium to regulate the proliferation of the neuronal retina (Pearson et al., 2005). Another recent achievement in this area is the combination of the luciferin–luciferase assay with fluorescent imaging, whereby ATP release could be followed from cell layers or even from individual cells, for example, from Muller glial cells of the retina (Newman, 2003a). Given its ubiquitous nature, ATP could be released not only from neurons, but from any kind of cells within and outside the nervous system also. Therefore, one of the most intriguing questions is to identify the source of ATP involved in the regulation of neuronal functions. Several neuronal pathways have been identified as a source of ATP release during neuronal activity, which include the septohabenular projection, innervating the rat medial habenula (Sperlagh et al., 1995, 1997, 1998a), the ventral noradrenergic bundle, identified as a source of ATP release in the hypothalamus (Sperlagh et al., 1998b), the Shaffer collateral

231

232

10

ATP‐mediated signaling in the nervous system

pathway providing the main excitatory input to the hippocampus (Wieraszko et al., 1989), the afferent nerve bundle projecting to the rat superior cervical ganglion (Vizi et al., 1997), and sympathetic nerves innervating the vas deferens (Kirkpatrick and Burnstock, 1987; Sperlagh and Vizi, 1992). ATP is shown to be released from these pathways in a vesicular fashion, that is, upon sodium channel activity and subsequent Ca2þ‐dependent exocytosis, alone or together with its cotransmitter mate. In other studies, no evidence was found for the neuronal origin of released ATP, which implicates that ATP may be also secreted via autocrine–paracrine pathways from nonneuronal cells (Hamann and Attwell, 1996; Juranyi et al., 1997; Sperlagh et al., 1999). Thus, besides neurons, one has to count on with glial cells, postsynaptic target cells, blood vessel endothelium, and the resident immune cells of the neural tissue, as the potential source of extracellular ATP. There are numerous stimuli that are able to release ATP from cultured glial cells, which include mechanical and inflammatory stimuli and the activation of ATP‐sensitive receptors themselves (Ballerini et al., 1996; Verderio and Matteoli, 2001; Coco et al., 2003). Nevertheless, the evidence that ATP is released upon similar stimuli from in situ glial cells is yet to be found. An additional mechanism, whereby ATP could be released in the nervous system is a receptor‐operated retrograde release from the postsynaptic target cells, which seems to be a significant mechanism in the periphery, at the neuromuscular junction (Smith, 1991; Vizi et al., 2000), and at the autonomic ganglia (Vizi et al., 1997) and neuro‐effector junctions (Vizi et al., 1992; Vizi and Sperlagh, 1999; Shinozuka et al., 2002). Postsynaptic ATP could contribute to neuronal ATP release and serves as an amplifying mechanism of the pre‐ and postsynaptic actions of extracellular ATP and its degradation product, adenosine. As ATP is a highly polarized molecule, which cannot pass freely through the cell membrane, it is also of interest to identify the mechanism, whereby it could enter the extracellular space. These include (i) vesicular exocytosis, (ii) carrier‐mediated release, (iii) release through channels and membrane pores, and (iv) cytolytic release. Vesicular exocytosis is a prototype mechanism for neurotransmitters to enter the extracellular space, which is expected to be a [Ca2þ]o‐dependent process. As ATP is the constituent of synaptic vesicles, it is reasonable to assume that exocytosis is accompanied by the release of ATP to the extracellular space. Indeed, [Ca2þ]o‐dependent ATP release in response to neuronal stimulation appears in almost all known neuro‐ neuronal and neuro‐effector connections of the nervous system (cf. Sperla´gh, 1996). Moreover, recent findings indicate that vesicular ATP release could be derived not only from nerve terminals but also from astrocytes (Coco et al., 2003). Although specific transporters, capable for the transmembrane movement of ATP, are yet to be molecularly identified in neurons, some data suggest that ATP could be also released in a carrier‐mediated manner. Especially, postsynaptic ATP release from smooth muscle cells appears to be driven by an ATP carrier, as this release, opposing from vesicular exocytosis, is strictly temperature dependent (> Figure 10-3) (Vizi and Sperlagh, 1999). In nonneuronal cells, ATP binding cassette (ABC) proteins have been implicated as an ATP transporter (al‐Awqati, 1995; Wang et al., 1996; Schwiebert, 1999), and these transporters are also expressed in glial cells (Ballerini et al., 2002) and mediate ATP release upon hypoosmotic challenge (Abdipranoto et al., 2003; Darby et al., 2003), but no evidence was found until now that these transporters are present and functional in central or peripheral nerves. Channels and pores are also potential candidates to drive the transmembrane movement of ATP. Connexin hemichannels are gap junction proteins, and their main function is to mediate electrical signaling, but they are also able to release neuroactive substances such as glutamate and ATP (Stout et al., 2004). Thus connexin hemichannels have been shown to mediate ATP release from astrocytes in response to mechanical stress (Stout et al., 2002) from xenopus oocytes (Bahima et al., 2006), from retina epithelium (Pearson et al., 2005), and from corneal endothelial cells (Gomes et al., 2005). Finally, a massive ATP release has been hypothesized to occur during any kind of cellular injury. This cytolytic ATP release may gain significance under a wide variety of pathological situations, for example, in hypoxia/ischemia, acute brain or spinal cord injury as well as in chronic neurodegenerative diseases, such as Alzheimer’s disease or sclerosis multiplex and in the generation of chronic pain, as demonstrated recently (Cook and McCleskey, 2002).

ATP‐mediated signaling in the nervous system

10

. Figure 10-3 The a1adrenoceptor‐mediated, postsynaptic ATP release and subsequent contraction is carried out by a temperature‐dependent, carrier‐mediated mechanism. Stimulation of the postsynaptic a1‐adrenoceptors by noradrenaline (NA, 300 mM) in the guinea‐pig vas deferens elicits ATP release, measured by the luciferin– luciferase assay (a, b) and a concomitant biphasic contraction (c, d), consisting of an initial twitch and a second, tonic response with superimposed oscillations. Cooling the bath temperature from 37
C to 12
C, which prevents carrier‐mediated mechanisms, inhibits ATP release and twitch contraction, but not the second phase of the contractile response, indicating that muscle contraction itself is not carrier mediated. ATP release is expressed in pico moles per gram, whereas twitch tension is expressed in millinewton. Diagrams show the mean  SEM of 5–8 experiments, **p < 0.01, calculated by the Student’s t‐test

4 The Extracellular Inactivation of ATP ATP, if it is released to the extracellular space, is no longer stable; its action is temporally and spatially terminated by the hydrolysis by the ecto‐nucleotidases, a widely expressed series of membrane bound enzymes, capable to dephosphorylate purine nucleotides and giving rise to the formation of purine nucleosides, including adenosine.

233

234

10

ATP‐mediated signaling in the nervous system

Several enzyme families are responsible for the extracellular degradation of ATP (> Figure 10-2). The first step of the inactivation of ATP is mediated by the family of ecto‐nucleoside triphosphate diphospho‐ hydrolases (E‐NTPDases, EC 3.6.1.5, also known as ectoATPase or apyrase) (cf. Zimmermann, 2000, 2001). E‐NTPDases have a molecular mass of 55–60 kDa in unglycosylated form, have one or two transmembrane domains, and highly conserved catalytic region faced to the extracellular space. Until now eight members of this enzyme family have been identified in molecular terms, numbered from E‐NTPDase 1 and to E‐NTPDase 8, and among them E‐NTPDase 1, 2, and 3 are the major ATP catabolizing enzyme of the brain (Kegel et al., 1997). Whereas E‐NTPDase 1 is able to hydrolyze ATP and ADP to AMP, E‐NTPDase 2 converts the nucleotide triphosphates to the respective diphosphates (cf. Zimmermann, 2000, 2001). These enzymes show widespread distribution in the brain (Wang et al., 1997; Wang and Guidotti, 1998) and have low micromolar Km for ATP and ADP giving rise to rapid and highly effective hydrolysis of ATP in almost all neuronal tissues, although Km values derived from biochemical determinations may vary between tissues and preparations. Thus, endogenous ATP is converted to adenosine to activate A1 adenosine receptors within a second in the hippocampus (Dunwiddie et al., 1997; Cunha et al., 1998) whereas the hydrolysis of ATP seems slower in other brain regions, such as the cerebral cortex (Cunha et al., 1994). Nevertheless, the short half‐life of ATP in the extracellular space could be still long enough for the activation of ionotropic ATP receptors, which act in a millisecond time scale, and probably also for metabotropic ATP receptors, which act on a slower, hundreds of millisecond–second time scale. E‐NTPDases have been shown to be upregulated after in vivo ischemia (Braun et al., 1998), indicating an increased endogenous ATP efflux to the extracellular space and the regulatory role of these enzymes between the nucleotidergic and adenosinergic signaling pathways under these conditions. In addition to the E‐NTPDase family, ATP could be also dephosphorylated by ecto‐nucleotide pyrophosphatases (E‐NPPs) and by alkaline phosphatases, both having broader substrate specificity, but also widespread tissue distribution (Zimmermann, 2000). The next step of extracellular inactivation is the hydrolysis of AMP by the ecto‐50 ‐nucleotidase (EC 3.1.3.5) enzyme, which is the rate‐limiting step giving rise to the formation of adenosine, which is a new extracellular signal, acting on its own receptors. The 50 ‐nucleotidase enzyme is a glycosylphosphatidyl inositol (GPI) anchored 62–74 kDa protein, which appears to exist mainly in homodimer form and linked to the plasma membrane with its active site exposed to the extracellular space (Zimmermann, 2001). Ecto‐ 50 ‐nucleotidase exhibits micromolar Km for AMP and is feed‐forwardly inhibited by ATP, and the synthetic analog a, b‐methylene adenosine diphosphate, which results in a delayed, burst‐like adenosine production (Cunha, 2001). It is also widely present in the brain and in the periphery, and it is predominantly associated to glial cells (Schoen et al., 1987; Grondal et al., 1988), although its expression has also been demonstrated in purified nerve terminals (Cunha et al., 1992; James and Richardson, 1993). Ecto‐50 ‐nucleotidase enzyme has been also implicated in disease conditions, for example, in epilepsy (Schoen et al., 1999; Bonan et al., 2000) and in vivo ischemia (Braun et al., 1997). In addition to enzymatic mechanisms, inactivating extracellular ATP, and giving rise to the formation of adenosine, there is also catalytic activity in the extracellular space for an ATP‐regenerating, reverse process, that is, to rephosporylate and interconvert nucleosides and nucleotides to ATP (> Figure 10-2). Although the ecto‐adenylate kinase (EC 2.7.4.3) and the ecto‐nucleoside diphosphokinase enzyme (EC 2.7.4.6), which can interconvert AMP, ADP, UDP, and UTP and ATP so far have been only demonstrated in nonneuronal cells (Harden et al., 1997; Yegutkin et al., 2002), one can assume that similar activity may also be present on the surface of neurons or glial cells. Moreover, the presence of an ectoATP:AMP phosphotransferase activity has already been verified on the surface of nerve terminals (Nagy et al., 1989; Terrian et al., 1989). Adenosine, generated by the ecto‐50 ‐nucleotidase or released on its own right, is then taken up to the nerve terminal and rapidly reincorporated to ATP stores, or deaminated extra‐ or intracellularly by the adenosine deaminase enzyme (> Figure 10-2). Specific nucleoside transporters, responsible for the uptake of adenosine, have two families: Equilibrative transporters (ENT) and concentrative transporters (CNT); the former is driven by the concentration gradient and the latter by the sodium gradient (Thorn and Jarvis, 1996; Cass et al., 1999). ENT transporters could carry different nucleosides including adenosine and inosine, but not nucleotides across the cell membrane in both directions, and are regarded as the dominant nucleoside transporters of the brain. ENT transporters seem to be widely expressed in both neuronal and

ATP‐mediated signaling in the nervous system

10

glial elements of the CNS (Anderson et al., 1999), whereas CNT transporters appear to exhibit a more restricted localization (Anderson et al., 1996). ENT transporters have two isoforms, ENT1 and ENT2; while the former is sensitive to inhibition by the adenosine uptake inhibitor nitrobenzylthioinosine (NBMPR), the latter is not (Griffiths et al., 1997; Yao et al., 1997). On the other hand, another transporter inhibitor, dipyridamole, appears to block both ENT1 and ENT2 transporters (Lee and Jarvis, 1988). Since the intracellular adenosine level under normal metabolic state is in the micromolar range (Latini and Pedata, 2001), if the ENT transporter is loaded from the extracellular space by excess adenosine it mediates its uptake into the nerve terminals. When adenosine is taken up intracellular adenosine‐eliminating mechanisms, that is the enzymes adenosine kinase and, in a lesser amount, adenosine deaminase convert it to AMP and inosine, respectively, thereby maintaining the driving force of the carrier. On the other hand, the ENT transporter could also act in a reverse direction under certain circumstances, mediating the release of adenosine into the extracellular space (> Figure 10-2). This could occur during energy deprivation or metabolic distress, when ATP stores are depleted and AMP is generated intracellularly. Cytosolic 50 ‐nucleotidase, which has a relatively high Km for AMP (1–14 mM), becomes active under these conditions and accumulates adenosine intracellularly. The resultant intracellular adenosine accumulation then flows out to the extracellular space in a transporter‐mediated manner. Since the intracellular concentration of ATP is about 50 times higher than that of AMP, even a small level of ATP depletion causes a relatively large increase in AMP concentration and leads to subsequent adenosine efflux; therefore, this mechanism is a very sensitive sensor of intracellular metabolic distress (Cunha, 2001; Latini and Pedata, 2001). ENT transporters could also mediate homo‐ or heteroexchange, if loaded from the extracellular space with relatively high concentration of nucleotides or nucleosides. Intracellular adenosine‐eliminating mechanisms in this case are not able to keep up with the increased uptake and could reverse the transporter. ATP and other nucleotides in this way may elicit adenosine release and indirectly influence adenosine receptor‐ mediated actions (Sperlagh et al., 2003). As for extracellular deamination of adenosine, it generates inosine, which could be also taken up by the ENT transporters. The proportion of the uptake and extracellular deamination may vary between tissues; in general, deamination seems to be more important in nonneuronal tissues, for example, in the cardiovascular system.

5 ATP Receptors ATP exerts its biological action through diverse families of P2 nucleotide receptors. P2 receptors could be subdivided into two families, ionotropic (P2X) and metabotropic (P2Y) receptors (Ralevic and Burnstock, 1998). Ionotropic P2X receptors are 379–595 amino acids long ligand‐gated cation channels, having two transmembrane domains (TM1 and TM2) and a large extracellular loop (> Figure 10-4) (Valera et al., 1994; Khakh et al., 2001). The ligand‐binding domain is located within a cysteine‐rich extracellular loop between the lysine residues of 69 and 71 positions, and binding sites for different antagonists have also been identified on the extracellular domain of the receptor protein. P2X receptors are nonselective cation channels having permeability to both monovalent (Naþ, Kþ) and divalent (Ca2þ) cations. Moreover, upon prolonged or repetitive agonist application they also display the property of pore dilation which makes the channel permeable to high molecular weight cations up to 800 Da. Until now, seven individual members of this receptor family have been identified molecularly, which are numbered from P2X1 to P2X7, having individual kinetics and pharmacological phenotype (> Table 10-1) (North and Surprenant, 2000). These receptor proteins, however, do not function as individual receptors, but coassemble into various homo‐ or heterooligomeric assemblies to form functional receptors. Among possible combinations, so far 16 variations have been proved to be functional (Torres et al., 1999). These are all of the homooligomeric receptors, except P2X6, which does not function in homooligomeric form, and the rest are heterooligomers, formed from P2X1–P2X6 subunits. However, recently it has been reported that by N‐glycosylation even the homomeric P2X6 receptor can be rendered in function (Jones et al., 2004). On the other hand, the P2X7 receptor functions only in homooligomeric form and does not coassemble with other known P2X receptor subunits. Basically, P2X receptors are sensitive to ATP and ADP but not to AMP and adenosine, and the ligand‐binding profile of homomeric P2X receptors are well established (for further information, see North

235

236

10

ATP‐mediated signaling in the nervous system

. Figure 10-4 Structure of P2X receptors. The receptor is 379–595 amino acids long, having two transmembrane domain (TM1 and TM2), intracellular N and C termini, and a large extracellular loop. The ligand‐binding domain is located within a cysteine‐rich extracellular loop between the lysine resides of 69 and 71 positions

and Surprenant, 2000). On the other hand, less is known about the pharmacology of heteromeric receptors; among them, the pharmacological profile of P2X2/3, P2X2/6, P2X1/2, P2X1/4, P2X1/5, and P2X4/6 are described (> Table 10-1) (Lewis et al., 1995; Le et al., 1998; Torres et al., 1998; King et al., 2000; Brown et al., 2002; Nicke et al., 2005). Briefly, the identification of receptor subtypes relies on their sensitivity to the agonist, a, b‐methylene ATP, as all the receptors except P2X2, P2X5, P2X7, and P2X2/6 are sensitive to this agonist, and on their sensitivity to the P2 receptor antagonists, PPADS and suramin. In addition, selective antagonists for certain subtypes of P2X receptors are also available, for example, NF449, which is a selective antagonist at the P2X1 receptor, or Brilliant Blue G, which is a potent and selective antagonist at the P2X7 receptor. As for the stoichiometry of the subunits, biochemical studies indicate that functional receptors are formed as trimers. In situ hybridization studies with specific riboprobes and immunocytochemical studies using antibodies raised against individual P2X receptor subunits revealed that all seven P2X receptors are widely expressed in the nervous system; however, the expression of individual receptor subunits are different and show region‐ and cell‐type‐specific distinct distribution (Collo et al., 1996). Among the P2X receptors, P2X2, P2X4, and P2X6 seem to be most abundantly expressed in the brain, whereas other subunits show more restricted localization (Collo et al., 1996; Vulchanova et al., 1996; Atkinson et al., 2000; Rubio and Soto, 2001). The typical localization of P2X2 receptor is on nerve terminals of the brain and the periphery (Vulchanova et al., 1997; Kanjhan et al., 1999; Atkinson et al., 2000), although it also appears postsynaptically (Rubio and Soto, 2001). P2X1 receptor has initially been suggested to be exclusively expressed on smooth muscle membrane, consistent with its role to mediate fast synaptic transmission at the autonomic neuro‐effector junction (Collo et al., 1996). However, more recent studies with more sensitive probes revealed that its expression is more widespread, that is, it is also present on the central and the peripheral neurons (Vulchanova et al., 1996; Calvert and Evans, 2004). The same holds true for P2X3 receptors, which are primarily associated to sensory pathways, but functional studies indicate that they are also expressed in other brain regions and autonomic pathways (Papp et al., 2004a; Knott et al., 2005; Rodrigues et al., 2005). P2X4 receptor shows heavy expression in several brain areas such as the cerebral cortex, the hippocampus,

ATP‐mediated signaling in the nervous system

10

. Table 10-1 Classification of identified purine/pyrimidine receptors Purine/pyrimidine receptors Nucleotide receptors

Adenosine receptors

Ionotropic

Metabotropic

Metabotropic

Homomeric

Heteromeric

Homomeric

Heteromeric

Homomeric

P2X1 P2X2 P2X3 P2X4 P2X5 P2X6 P2X7

P2X1/P2X2 P2X2/P2X3 P2X2/P2X6 P2X4/P2X6 P2X1/P2X4 P2X1/P2X5

P2Y1 P2Y2 P2Y4 P2Y6 P2Y11 P2Y12 P2Y13 P2Y14

P2Y1/A1

A1 A2A A2B A3

Purine and pyrimidine sensitive receptors are divided into nucleotide‐ and adenosine receptors. Nucleotide receptors, which include all ATP‐sensitive receptors, can be further subdivided into subfamilies of ionotropic P2X and metabotropic P2Y receptors, whereas adenosine receptors are all metabotropic receptors. The P2X receptor family has seven individual members, which are numbered from P2X1 to P2X7. These receptor proteins, however, do not function as individual receptors, but coassemble into various homo‐ or heterooligomeric assemblies to form functional receptors. Among possible combinations, 16 variations have been proved to be functional, these are all of the homooligomeric receptors, whereas among the functional heteromeric receptors, the pharmacological phenotype of P2X2/3, P2X2/6, P2X1/2, P2X1/4, P2X1/5, and P2X4/6 are described. P2Y receptor family has eight individual members, numbered as P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13, and P2Y14. P2Y1 receptors could also heteromerize with A1 adenosine receptors, which results in a hybrid receptor, activated by both A1‐adenosine and P2‐receptor agonists

the thalamus, and the brainstem (Le et al., 1997) and is associated with postsynaptic specialization of synaptic contacts (Rubio and Soto, 2001). P2X5 subunits have the most restricted localization in the brain, although it shows strong representation in certain areas, for example, nucleus tractus solitarii (NTS) (Yao et al., 2001). Finally, the expression of P2X7 receptor in the nervous system is subject of a current debate, in contrast to the initial in situ hybridization studies, which proposed that P2X7 receptors are not expressed in the adult brain, except in reactive microglia and astroglia (Collo et al., 1997), immunocytochemical studies revealed a widespread presynaptic expression of P2X7 receptor immunoreactivity in a number of different brain areas, including the brainstem, the hippocampus, the cortex, the spinal cord, and the skeletal neuromuscular junction (Deuchars et al., 2001; Sperlagh et al., 2002; Atkinson et al., 2004). Although two studies demonstrated a pseudo‐immunoreactivity in the brain of P2X7 receptor knockout animals (Kukley et al., 2004; Sim et al., 2004), a more recent study indicates that the pseudo‐immunoreactivity represents a brain analog of P2X7 receptor, which shares its antibody‐binding domain with the cloned P2X7 receptor and partially retains its functionality (Sanchez‐Nogueiro et al., 2005). Whether this molecular entity is a genuine new receptor, a splice variant of P2X7 receptor, or a developmental side product, which is present only in P2X7 null mice, warrants further investigation. P2Y receptors all belong to G protein‐coupled receptors, having seven hydrophobic transmembrane domains, and possess their ATP‐binding site on the external side of TM3 and TM7 domains (von Kugelgen and Wetter, 2000; Barnard and Simon, 2001; Boarder and Webb, 2001; Communi et al., 2001; Abbracchio et al., 2003). P2Y receptor family has eight individual members, numbered as P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13, and P2Y14 (> Table 10-1). P2Y receptors are basically activated by adenine and uridine nucleotides, such as ATP, ADP, UDP, and UTP, but not by nucleosides and they could be further subdivided into adenine nucleotide‐preferring and uridine nucleotide‐preferring subgroups. P2Y1, P2Y11, P2Y12, and P2Y13 receptors belong to the former subgroup, whereas P2Y2, P2Y4, P2Y6, and P2Y14 receptors belong to

237

238

10

ATP‐mediated signaling in the nervous system

the latter. Although the pharmacological phenotype of all cloned P2Y receptor subtypes are yet to be elaborated, some ligands are also available which display considerable selectivity to certain subtypes of the P2Y receptor family. Thus 2‐methylthio‐ADP is a selective agonist at the P2Y1 and P2Y12 receptors, whereas MRS2179 is a selective antagonist of P2Y1 receptors. Interestingly, P2Y1 receptors could also heteromerize with A1 adenosine receptors, which results in a hybrid receptor, activated by both A1‐adenosine and P2‐receptor agonists (> Table 10-1) (Yoshioka et al., 2001). In comparison with P2X receptors, our knowledge is less complete on the exact localization and cell type‐specific distribution of P2Y receptors. mRNA‐encoding P2Y1, P2Y12, P2Y13, and P2Y14 are undoubtedly present in the brain (Chambers et al., 2000; Communi et al., 2001; Hollopeter et al., 2001; Nicholas, 2001), and among them, P2Y1 receptor has widespread distribution also at different brain areas and associated with both neurons and astrocytes at the protein level (Moran‐Jimenez and Matute, 2000; Moore et al., 2001).

6 The Fast Transmitter Action of ATP The principal function attributed to extracellular ATP is that in activating postsynaptic P2X receptors, it acts as a fast excitatory neurotransmitter in neuro‐neuronal and neuro‐effector synapses. Thus, P2X receptor‐ gated synaptic currents were identified in numerous parts of the CNS and the PNS, and the first in this row in the brain was the identification of a P2 receptor‐mediated excitatory synaptic current in the medial habenula (Edwards et al., 1992), followed by the demonstration of similar currents in other neuro‐neuronal synapses such as the submucosal and celiac neurons (Evans et al., 1992; Silinsky et al., 1992), enteric neurons (Bardoni et al., 1997), locus coeruleus nucleus of the brainstem (Nieber et al., 1997; Poelchen et al., 2001), lateral hypothalamus (Jo and Role, 2002), rat trigeminal mesencephalic neurons (Patel et al., 2001), dorsal horn of the spinal cord (Galligan and Bertrand, 1994), CA1 and CA3 regions of the hippocampus (Pankratov et al., 1998), and the somato‐sensory cortex (Pankratov et al., 2002). Nevertheless, it should be noted that even after a decade of the first demonstration of a purinergic synapse in the brain, the number of identified purinergic currents is still relatively limited. Moreover, purinergic currents could be recorded usually only in a proportion of cells and only during evoked but not in spontaneous synaptic transmission. In general, relatively strong stimulation paradigm and simultaneous blockade of the action of other excitatory and inhibitory neurotransmitters, that is, GABA, acetylcholine, and serotonin, are also necessary conditions to observe purinergic currents (Khakh, 2001; Illes and Ribeiro, 2004). On the other hand, identification of purinergic pathways is hindered by the lack of potent and selective antagonists and the lack of specific morphological markers, which otherwise are routinely used for the identification of transmitters, for example, in case of classical transmitters or peptides. It is also unclear, whether ATP acts in these synapses as a genuine cotransmitter, released from common vesicles with its cotransmitter mate or an individual transmitter. Finally, future studies should prove the in vivo relevance of purinergic synapses. In neuro‐effector synapses, the cotransmitter role of ATP is well established, as it acts as a cotransmitter with noradrenaline at the sympathetic neuro‐effector junction, which has been demonstrated in a number of tissue preparations, including the vas deferens, several blood vessels, cutaneous microcirculation, etc. (Sneddon et al., 2000). It also acts as a cotransmitter with acetylcholine in certain parts of the parasympathetic neuro‐effector transmission sites, such as at the smooth muscle of the urinary bladder (Kennedy, 2001).

7 The Presynaptic Modulatory Role of ATP The presynaptic nerve terminal is an important checkpoint, whereby the efficacy of synaptic transmission could be locally and efficiently controlled (McGehee and Role, 1996; MacDermott et al., 1999; Vizi, 2000; Boehm and Kubista, 2002). Although originally it was suggested that ATP‐sensitive P2 receptors are located

ATP‐mediated signaling in the nervous system

10

exclusively on postsynaptic sites, it was already recognized in the early 1990s that they are involved in the regulation of transmitter release. Thus it has been reported that a, b‐methylene ATP, an analog of ATP which is resistant to degradation, facilitates electrically evoked acetylcholine release from myenteric plexus of guinea pig, an effect which was not blocked by the antagonists of adenosine receptors, and therefore mediated by P2 receptors (Sperlagh and Vizi, 1991). Later on, this suggestion has been confirmed by electrophysiological tools (Sun and Stanley, 1996), and ATP‐activated ligand‐gated ion channels have been identified to be responsible for the facilitation acetylcholine release. Moreover, ATP is able not only to facilitate, but also directly trigger transmitter release, without preceding action potential and subsequent activation of voltage‐sensitive Ca2þ channels. Since P2X receptors have relatively high Ca2þ permeability (Rogers et al., 1997; Egan and Khakh, 2004), this property makes them capable to initiate neurotransmitter release by the Ca2þ influx through the receptor ion channel complex, provided that they are located near the release sites. Activation of P2X receptors elicits noradrenaline release from sympathetic nerve terminals of the rat (Boehm, 1999) and guinea‐pig (Sperlagh et al., 2000), as well as from central noradrenergic varicosities innervating the hippocampus (Papp et al., 2004a). As for the P2X receptor subunits involved, there is a considerable species and region heterogeneity; for example, P2X2 receptors have been suggested to be responsible for the enhancement of noradrenaline release from sympathetic nerve terminals of the rat (Boehm, 1999), whereas in the guinea pig P2X3 or the heteromeric P2X2/3 receptors have been shown to be involved (Sperlagh et al., 2000; Queiroz et al., 2003); moreover, P2X1 receptors also contribute to the response in central noradrenergic pathways (Papp et al., 2004a). In addition, P2 receptors enhance the release of serotonin from the hippocampus (Okada et al., 1999) and that of dopamine from the striatum (Zhang et al., 1995, 1996) and the nucleus accumbens (Krugel et al., 1999, 2001) in vivo, although the latter effects are thought to be mediated by P2Y receptors. The release of other transmitters, including the main excitatory and inhibitory transmitters of the brain, are also subject to facilitation by presynaptic P2X receptors both in the CNS and in the periphery, as confirmed partly by neurochemical and partly by electrophysiological methods. Activation of P2X receptors elicits glutamate release in the spinal cord (Gu and MacDermott, 1997; Nakatsuka and Gu, 2001), brainstem (Khakh and Henderson, 1998; Kato and Shigetomi, 2001; Shigetomi and Kato, 2004), and the hippocampus (Sperlagh et al., 2002; Khakh et al., 2003; Rodrigues et al., 2005; Fellin et al., 2006). As for the underlying receptor subunits involved in these effects, P2X2, P2X7 (Sperlagh et al., 2002; Khakh et al., 2003; Papp et al., 2004b; Fellin et al., 2006) (also confirmed by the use of subunit specific knockout mice), as well as P2X1, P2X3, and P2X2/3 receptors (Rodrigues et al., 2005) are all identified in the hippocampus, whereas in the spinal cord, P2X3 and an unknown receptor (Nakatsuka et al., 2003) are implicated. On the other hand, the regulation of GABA release by P2X receptors seems to be more restricted, with the exception of spinal cord (Hugel and Schlichter, 2000), cultured cortical (> Figure 10-5) (Wirkner et al., 2005) and hippocampal cells (Inoue et al., 1999), and the brainstem, where the excitatory and the inhibitory synaptic transmissions are subtype specifically facilitated, via P2X3 and P2X1 receptors, respectively (Watano et al., 2004). On the other hand, no evidence was found for a direct facilitation of GABA release by P2 receptor in the hippocampal nerve terminal preparation (Cunha and Ribeiro, 2000b). Nevertheless, the release of another inhibitory transmitter, glycine, is augmented by P2X receptor activation in the dorsal horn (Rhee et al., 2000) and in the brainstem trigeminal nucleus (Wang et al., 2001). In addition to facilitatory modulation, P2 receptors are also involved in the inhibitory modulation of the release of various transmitters and the metabotropic P2Y receptors, which are engineered to act on a longer time scale, play a major role in these actions. Hence, in the CNS, ATP inhibits the release of acetylcholine (Cunha et al., 1994), noradrenaline (von Kugelgen et al., 1994; Koch et al., 1997), serotonin (von Kugelgen et al., 1997), dopamine (Trendelenburg and Bultmann, 2000), and glutamate (Koizumi and Inoue, 1997; Inoue et al., 1999; Bennett and Boarder, 2000; Mendoza‐Fernandez et al., 2000), whereas the release of GABA seems to be not subject to inhibitory neuromodulation by P2 receptors. Convincing evidence is also available on similar modulation of acetylcholine and noradrenaline release in the periphery (for further references, see Cunha and Ribeiro, 2000a).

239

240

10

ATP‐mediated signaling in the nervous system

. Figure 10-5 The P2X receptor agonist BzATP enhances the frequency, but not the amplitude of GABAA receptor‐mediated mIPSCs in neurons of rat cortical cell cultures (10–15 DIV) with a presynaptic site of action (from Wirkner et al., 2005, with permission from Blackwell Publishing). (a, b) Consecutive traces showing typical experiments in two individual cells. The P2X7 receptor‐selective antagonist Brilliant Blue G (BBG) alone has no effect (a), whereas it prevents the effect of BzATP on the frequency of mIPSCs (b). (c) Increase of the mean frequency (empty columns) but not amplitude (filled columns) of mIPSCs by BzATP; no increase of mIPSC amplitude in the presence of BBG is detected (n ¼ 7). The changes were expressed as percentage potentiation of the time‐ matching controls recorded in drug‐free ACSF (for control traces see a; n ¼ 5). Average values of amplitude and frequency of mIPSCs were calculated during a control period of 3 min, during the last 3 min of the subsequent application of BBG (0.3 mM) for 10 min in total, as well as during the last 3 min of further superfusion with BBG alone or in combination with BzATP (300 mM) for 10 min. *p < 0.05; statistically significant difference from zero. **p < 0.05; statistically significant difference from the effect of BzATP in the absence of BBG. (d) Effect of BzATP (300 mM) on the inward current induced by the GABAA agonist muscimol (10 mM) locally superfused for 2 s with 3‐min intervals. BzATP was superfused for 6 min during two consecutive muscimol applications. Typical recording out of five similar ones. mIPSCs were recorded at a holding potential of 60 mV. CNQX (10 mM), AP‐5 (50 mM), and TTX (0.5 mM) were all present in the medium

ATP‐mediated signaling in the nervous system

10

8 The Role of ATP in Glia–Neuron and Glia–Glia Signaling In addition to its role as a fast transmitter and as a presynaptic modulator, rapidly emerging data indicate that ATP is an important signaling molecule in the communication between glia and neurons and within glial networks. Although glial cells are traditionally regarded as a simple support for the neuronal networks, now it has become clear that they are more active players in synaptic transmission (Araque et al., 2001; Newman, 2003b). Thus, there is a bidirectional communication between neurons and glial cells, and not only does the glia respond to signals originating from neurons, but it can also release transmitters, which then act on cell‐surface receptors present on the neuronal membrane and modulate synaptic activity pre‐ and postsynaptically. Among these gliotransmitters, ATP seems to be one of the most important, in addition to glutamate and other amino acids (> Figure 10-6) (Fields and Stevens, 2000). Hence, mechanical stimulation of astrocytes in hippocampal cell culture leads to the generation of Ca2þ waves in astrocytes, which spread by the release of ATP and subsequent activation of P2 receptors and lead to the depression of excitatory synaptic transmission between neurons (Koizumi et al., 2003). This glia‐driven synaptic depression is partly mediated by ATP itself acting on P2Y receptors and partly by adenosine acting on A1 adenosine receptors (Koizumi et al., 2003), and a similar mechanism has been demonstrated in the retina, where glial‐derived ATP depress the firing of ganglion cells via activation of A1 adenosine receptors (Newman, 2003a). Moreover, endogenous ATP, released activity dependently from hippocampal astrocytes in response to the action of glutamate on non‐NMDA receptors, seems also to act similarly causing homo‐ and heterosynaptic suppression of excitatory transmission, presumably again via its degradation product adenosine (Zhang et al., 2003). On the other hand, astrocytic ATP is also involved in the modulation of inhibitory transmission in the hippocampus, by the excitation of inhibitory interneurons via P2Y1 receptors, which leads to an increased synaptic inhibition within intact hippocampal synaptic networks (Bowser and Khakh, 2004) and in long‐term synaptic plasticity events (Pascual et al., 2005). Finally, in addition to presynaptic modulation, glial‐derived ATP could also cause enduring changes in postsynaptic efficacy; in the hypothalamic paraventricular nucleus, noradrenaline, acting on a1‐adrenoceptors, releases ATP from astrocytes, which then acting on P2X7 receptors enhance excitatory transmission postsynaptically by the activation of phosphatidylinositol 3‐kinase and subsequent insertion of AMPA receptors to the cell membrane (Gordon et al., 2005). In addition to its mediator role between glial cells and neurons, ATP is also the primary mediator of the extracellular communication between astrocytes (Guthrie et al., 1999). Astrocyte populations coordinate their functions via Ca2þ waves, and the spread of the Ca2þ signal is implemented by two ways: An intercellular pathway mediated by gap junctions and an extracellular pathway mediated by ATP and P2

. Figure 10-6 ATP is a gliotransmitter. Mechanical stimulation of astrocytes leads to the generation of Ca2þ waves in astrocytes, which spread by the release of ATP and subsequent activation of P2 receptors and lead to the depression of excitatory synaptic transmission between neurons. This glia‐driven synaptic depression is partly mediated by ATP itself acting on P2Y receptors and partly by adenosine, acting on A1 adenosine receptors. Moreover, endogenous ATP, released activity dependently from hippocampal astrocytes in response to the action of glutamate on non‐NMDA receptors, seems also to act similarly causing homo‐ and heterosynaptic suppression of excitatory transmission. On the other hand, astrocytic ATP is also involved in the modulation of inhibitory transmission, by the excitation of inhibitory interneurons via P2Y1 receptors, which leads to an increased synaptic inhibition within intact hippocampal synaptic networks. Finally, in addition to presynaptic modulation, glial‐derived ATP could also cause enduring changes in postsynaptic efficacy. In the hypothalamic paraventricular nucleus, noradrenaline, acting on a1‐adrenoceptors, releases ATP from astrocytes, which then acting on P2X7 receptors enhances excitatory transmission postsynaptically by the activation of phosphatidylinositol 3‐kinase and subsequent insertion of AMPA receptors to the cell membrane. Astrocyte‐derived ATP may also activate P2X7 receptors on microglial cells and elicit Ca2þ signals in the microglia, which eventually leads to cytolysis of this cell type. For corresponding references, see text

241

242

10

ATP‐mediated signaling in the nervous system

. Figure 10-6 (continued)

receptors (Guthrie et al., 1999). Astrocytes communicate by calcium‐mediated signaling not only with each other but also with neighboring cells including neurons (see earlier, Koizumi et al., 2003) and microglia. Thus, astrocyte‐derived ATP activates P2X7 receptors on microglial cells and elicits Ca2þ signals in the microglia, which eventually leads to cytolysis of this cell type (Verderio and Matteoli, 2001).

ATP‐mediated signaling in the nervous system

10

9 The Role of ATP in Sensory Transmission and in the Generation of Pain ATP and its action on ligand‐gated P2X and metabotropic P2Y receptors appear also to strongly participate in the sensory transmission, and in particular in the generation of chronic inflammatory and neuropathic pain (Chizh and Illes, 2001; Kennedy et al., 2003; Kennedy, 2005) and in the mechano‐ and chemosensory transduction (Burnstock and Wood, 1996; Burnstock, 2001, 2006). Although the algogenic action of ATP has been early recognized (Collier et al., 1966), the molecular mechanism underlying this action has only recently been understood, and several different P2 receptors seem to be involved. P2X receptors are expressed along the nociceptive pathways; mRNA encoding all subunits of the P2X receptor is expressed in sensory ganglia, including dorsal root, trigeminal, and nodose ganglia (Dunn et al., 2001; Ruan et al., 2005), and among them, the expression of mRNA‐encoding P2X3 receptor is in particular high. At the protein level, primarily, the small‐diameter IB4 expressing, capsaicin‐sensitive C neurons are those that express P2X3 receptor (Vulchanova et al., 1997; Bradbury et al., 1998), whereas functional studies indicate that P2X2, P2X2/3 (Li et al., 1999; Petruska et al., 2000), P2X1/5, and P2X4/6 receptors are expressed on the medium‐diameter capsaicin‐insensitive Ad fibers (Nakatsuka et al., 2003; Tsuzuki et al., 2003). All these P2X receptor subunits are also expressed at the central terminals of sensory afferents, and their activation releases glutamate and thereby facilitate excitatory transmission (Gu and MacDermott, 1997; Nakatsuka and Gu, 2001). As evidenced by the use of the P2X3‐selective antagonist A317491, antisense (Barclay et al., 2002; Honore et al., 2002), siRNA (Dorn et al., 2004), and knockout (Cockayne et al., 2000; Souslova et al., 2000) strategies, the activation of P2X3 receptor is involved in the generation of pain of different chronic inflammatory and neuropathic pain models, while they seem to be silent during acute pain and nociceptive reactions, which makes P2X3 receptor an attractive therapeutic target in the management of these disease states. As for the mechanism underlying the P2X3 receptor‐mediated pain, the constant activation of P2X3 receptors by ATP, released from the sensory neurons themselves (Holton, 1959) or by cytolysis (Cook and McCleskey, 2002), has been postulated, which could activate P2X3 receptors present on the nerve terminals in a self‐regenerative way and leading to the sensitization of the pathway and causing hyperalgesia and allodynia, characterized by chronic pain states (Kennedy, 2005). P2X2/3 and P2X3 receptors are also involved in the mechanosensory transduction of the bladder, where ATP is released from epithelial cells in response to bladder distension and participates in the induction of visceral pain and subsequent local reflexes (Vlaskovska et al., 2001; Burnstock, 2006; Cockayne et al., 2005). A similar role of purinergic mechanosensory transduction has also been implicated in case of a variety of other viscera, including the ureter, gut, and tooth pulp (see Burnstock, 2006). In addition to P2X3 and P2X2/3 receptors, other types of P2 receptors are also involved in sensory mechanisms and in the initiation and the regulation of pain. Hence, pharmacological blockade of P2X4 receptor reverse tactile allodynia (Tsuda et al., 2003) and genetic deletion of P2X7 receptors completely abolish chronic inflammatory and neuropathic pain in the Freund adjuvant‐induced and partial nerve ligation‐induced models, respectively (Chessell et al., 2005); however, in these latter studies, the action of ATP is attributed to P2X receptors expressed on microglial cells, which are activity dependently expressed during microglial activation and involved in the production of inflammatory mediators (see later). Finally, P2Y1 and P2Y4 receptor mRNA and protein are also heavily expressed on primary afferent nerve terminals and colocalize with P2X3 receptor (Ruan and Burnstock, 2003). However, in contrast to P2X receptors, the activation of P2Y1 receptors decreases the release of sensory transmitters, and thereby may counterbalance the algogenic action of ATP on P2X receptors (Gerevich et al., 2004). In addition to mechanosensory transduction, rapidly emerging data support a key role of extracellular ATP in central and peripheral chemosensory transduction, controlling respiration (Gourine, 2005). ATP, released from the glomus caroticum and activating P2X2 and/or P2X2/3 receptors located on the dendrites of carotid sinus nerve, controls the ventilatory response to hypoxia at the peripheral chemoreceptor area (Rong et al., 2003). Moreover, ATP is also released from the ventral surface of the central chemosensitive areas of the medulla oblongata and participates in the central chemosensory transduction evoked by the change of blood Hþ/CO2, that is, hypercapnia (Thomas et al., 1999, 2001; Thomas and Spyer, 2000;

243

244

10

ATP‐mediated signaling in the nervous system

Gourine et al., 2005a, b), and the involvement of P2 receptors in the Hþ/CO2‐dependent regulation in the cortical excitability has also been observed (Dulla et al., 2005). Nevertheless, the identity of P2 receptors involved in the latter actions is still elusive.

10 The Role of P2 Receptors in Behavioral Paradigms Apart from data detailed earlier on sensory information processing, there is still a paucity of information on the function of ATP‐sensitive P2 receptors in complex neuronal functions such as learning, memory, and behavior. Although studies using knockout mice for one or more P2 receptor subunits have not revealed gross abnormalities in behavior (Cockayne et al., 2005), one cannot rule out the potential compensatory developmental upregulation in these studies, and this area clearly deserves further investigations, including more sensitive test batteries. Nevertheless, some hints of data indicate that P2 receptors indeed regulate various aspects of behavior. Intraaccumbal injection of the P2 receptor agonist 2‐methyl‐thio‐ATP enhances locomotor activity and elicits correspondent changes in EEG pattern, an effect abolished by 6‐OHDA pretreatment, and therefore is mediated by the mesolimbic dopaminergic pathway (Kittner et al., 2000). The P2 receptor antagonist, PPADS, prevents both the acute locomotor effects of amphetamine and the behavioral sensitization caused by repeated amphetamine injections in rats, indicating the participation of an endogenous purinergic signaling mechanism in the behavioral effect of psychostimulants (Kittner et al., 2001). Moreover, the blockade of P2 receptors by PPADS in the nucleus accumbens also suppresses feeding‐evoked dopamine release and feeding behavior (Kittner et al., 2004). On the other hand, it is also revealed that exogenous and endogenous activation of P2Y1 receptors elicits anxiety‐like behavior in the elevated plus maze test in rats through the activation of NO signaling (Kittner et al., 2003).

11 ATP as a Neuroimmunomodulator Microglial cells originate from monocyte or macrophage precursors and are regarded as the major immunocompetent cell type of the nervous system. Thus, they are rapidly activated in response to pathological signals such as ischemia and inflammation and respond with morphological changes transforming the resting ramified microglia to an amoeboid form with phagocytic activity, proliferation, and the production of a wide array of inflammatory mediators. Therefore, microglial activation is heavily implicated in the pathogenesis in CNS diseases and the following repair process. It has been known for a considerable time that microglial cells respond with both ionotropic and metabotropic response to ATP application (Walz et al., 1993; Norenberg et al., 1994). Later it was confirmed that all members of the P2 receptor family are expressed on resting and activated microglial cells kept in culture at the mRNA and/or protein level (Bianco et al., 2005; Xiang and Burnstock, 2005). Among various subtypes of the P2 receptors, the role of P2X7 receptors in microglial response is especially well delineated. Microglial cell lines respond to P2X7 receptor activation (Haas et al., 1996; Chessell et al., 1997; Visentin et al., 1999), with membrane depolarization, a sustained increase in intracellular free Ca2þ (Ferrari et al., 1996), the uptake of high molecular weight fluorescent dyes (Ferrari et al., 1996; Chessell et al., 1997), and the secretion of IL‐1b upon LPS stimulus (Ferrari et al., 1997b, c; Sanz and Di Virgilio, 2000). The central role of P2X7 receptors, as costimulators of the posttranslational processing of IL‐1b in microglial cells upon LPS challenge, has been repeatedly proven (Ferrari et al., 1997b, c; Sanz and Di Virgilio, 2000; Brough et al., 2002). The mechanism underlying ATP‐dependent IL‐1b maturation and release involves an outwardly directed Kþ conductance and the activation of the interleukin‐1‐converting enzyme (ICE, also known as caspase 1) responsible for the cleavage of pro‐IL1b to the mature, 17 kDa form (Sanz and Di Virgilio, 2000). This mechanism appears to participate not only in the exogenous but also in the endogenous activation of P2X7 receptors upon LPS challenge (Ferrari et al., 1997c). Moreover,

ATP‐mediated signaling in the nervous system

10

after priming of the cells by ATP challenge, ADP and AMP also act as promoters of membrane currents and LPS‐induced IL‐1b secretion in these cells (Chakfe et al., 2002). In addition to IL‐1b, the synthesis and release of other cytokines and inflammatory mediators are also stimulated by P2X7 receptor activation in the microglia. Hence, ATP is a full stimulus (i.e., without the requirement of priming by LPS) to induce TNFa production via a Ca2þ‐dependent, ERK/JNK/p38 signaling pathway (Hide et al., 2000; Suzuki et al., 2004), induce cyclooxigenase‐2 (COX‐2) expression and seems to participate in the regulation of IL‐6 production (Chessell et al., 2005), although other subtypes of P2 receptors may be also involved in this latter effect (Inoue, 2002; Shigetomi and Kato, 2004). Moreover, Rampe et al. (2004) revealed that P2X7 receptors play a role in the distinct modulation of cytokine secretory pathways not only after LPS, but also upon amyloid beta peptide (Ab) preactivation. Whereas the production of IL‐1b, IL‐1a, TNFa, and IL‐18 was increased, that of IL‐6, the antiinflammatory cytokine, was attenuated under these conditions, implicating the involvement of P2X7 receptors in the pathogenesis of Alzheimer’s disease (Rampe et al., 2004). P2X7 receptor activation also induces iNOS mRNA expression and increases NO production from rat microglia (Ohtani et al., 2000), enhances IFNg‐induced iNOS expression and subsequent NO production in the murine BV‐2 microglial cell line (Gendron et al., 2003), and promotes the generation of reactive oxygen intermediates (ROI) by the p38MAPK pathway (Parvathenani et al., 2003). Finally, the activation of P2X7 receptors elicits a pronounced increase in 2‐AG secretion in the astroglial cells (Walter et al., 2004) and in the microglia (Witting et al., 2004), and low concentration of agonists stimulates the release of the neuroprotective mediator plasminogen from cultured microglia (Inoue et al., 1998). Therefore, regulation of the production of putatively protective (plasminogen, TNFa, 2‐AG) and harmful (IL‐1b, NO) inflammatory mediators by P2X7 receptors appears to follow a highly time‐ and concentration‐dependent pattern (Inoue, 2002). According to its pore‐forming property, the activation of P2X7 receptors also leads to cytolysis in an apoptotic fashion in the microglia (Ferrari et al., 1997a), which involves the proteolytic pathway of the caspase activation, although this is not an absolute requirement for the membrane damage and cytolysis (Ferrari et al., 1999; Brough et al., 2002). In addition to P2X7 receptors, other subtypes of the P2 receptor family are also involved in different aspects of neuroimmunomodulation, which responds to lower concentration of ATP (cf. McLarnon, 2005). Thus microglial Ca2þ influx could be also initiated by lower agonist concentrations, effects, which could be attributed to a non‐P2X7 ionotropic, probably P2X4 receptor activation, and there is also a metabotropic, P2Y receptor‐mediated response (Visentin et al., 1999; McLarnon, 2005). The activation of these receptors also induces COX‐2 in the human microglia (Choi et al., 2003), and P2Y1 receptors mediate the inhibition of the LPS‐induced IL‐1b, IL‐6, and TNF‐a production (Ogata et al., 2003). The level of proinflammatory cytokines, therefore seems to be differentially and oppositely regulated by ionotropic and metabotropic P2 receptors, similar to that observed in the case of the regulation of transmitter release. Finally, the microglial expression of P2X4 receptors and subsequent activation of the p38 MAP kinase pathway have been shown to be involved in the generation of inflammatory and neuropathic pain (see also Inoue et al., 2004; Guo et al., 2005; Schwab et al., 2005).

12 Involvement of ATP Receptors CNS Diseases and their Potential Therapeutic Exploitation The widespread involvement of ATP and its receptors in different neuronal functions implicate their role in the pathology of the nervous system as well. Moreover, ATP seems to play a more active pathogenetic role in a variety of CNS diseases, where the upregulation or the distinct activation of the ATP‐mediated signaling system provides a basis for disease‐selective therapeutic intervention. The disease conditions, where P2 receptors seem to play a role include ischemia or hypoxia, neurotrauma, epilepsy, Alzheimer’s and Parkinson’s disease, pain, drug addiction, and brain tumors (see for more details Koles et al., 2005; Franke and Illes, 2006).

245

246

10

ATP‐mediated signaling in the nervous system

13 Conclusions and Future Avenues It is now clear that ATP is one of the important signaling molecules in the nervous system, which is involved in a wide variety of neuronal functions, including synaptic transmission, neuromodulation, glia–neuron interactions, and neuroimmunomodulation. Therefore, the initial idea of purinergic nervous system developed into a more diverse concept of purinergic signaling system. Nevertheless, there are a number of aspects which need further investigation. Despite the wealth of data on ATP‐mediated signaling at the molecular and the cellular levels, the present knowledge is still limited at the systems level. This holds true to any aspect of purinergic mechanism, including the release and the inactivation mechanisms and to ATP‐receptor‐mediated responses as well, which are well characterized in recombinant systems, but poorly extrapolated to in vivo conditions. The progress along this line might lead to the therapeutic utilization of purinergic signaling system, which offers a number of potential target sites for pharmacological intervention.

Acknowledgments This study was supported by grants of the Hungarian Research Foundation (OTKA T037457), Hungarian Medical Research Council (472/2003), Hungarian Research and Development Fund (NKFP1A/002/2004), and the Volkswagen Foundation (I777 854).

References Abbracchio MP, Burnstock G. 1998. Purinergic signalling: Pathophysiological roles. Jpn J Pharmacol 78: 113-145. Abbracchio MP, Boeynaems JM, Barnard EA, Boyer JL, Kennedy C, et al. 2003. Characterization of the UDP‐glucose receptor (re‐named here the P2Y14 receptor) adds diversity to the P2Y receptor family. Trends Pharmacol Sci 24: 52-55. Abdipranoto A, Liu GJ, Werry EL, Bennett MR. 2003. Mechanisms of secretion of ATP from cortical astrocytes triggered by uridine triphosphate. Neuroreport 14: 2177-2181. al‐Awqati Q. 1995. Regulation of ion channels by ABC transporters that secrete ATP [comment]. Science 269: 805-806. Anderson CM, Xiong W, Young JD, Cass CE, Parkinson FE. 1996. Demonstration of the existence of mRNAs encoding N1/cif and N2/cit sodium/nucleoside cotransporters in rat brain. Brain Res Mol Brain Res 42: 358-361. Anderson CM, Xiong W, Geiger JD, Young JD, Cass CE, et al. 1999. Distribution of equilibrative, nitrobenzylthioinosine‐ sensitive nucleoside transporters (ENT1) in brain. J Neurochem 73: 867-873. Araque A, Carmignoto G, Haydon PG. 2001. Dynamic signaling between astrocytes and neurons. Annu Rev Physiol 63: 795-813. Atkinson L, Batten TF, Deuchars J. 2000. P2X(2) receptor immunoreactivity in the dorsal vagal complex and area postrema of the rat. Neuroscience 99: 683-696. Atkinson L, Batten TF, Moores TS, Varoqui H, Erickson JD, et al. 2004. Differential co‐localisation of the P2X7 receptor

subunit with vesicular glutamate transporters VGLUT1 and VGLUT2 in rat CNS. Neuroscience 123: 761-768. Bahima L, Aleu J, Elias M, Martin‐Satue M, Muhaisen A, et al. 2006. Endogenous hemichannels play a role in the release of ATP from Xenopus oocytes. J Cell Physiol 206: 95-102. Ballerini P, Di Iorio P, Ciccarelli R, Nargi E, D’Alimonte I, et al. 2002. Glial cells express multiple ATP binding cassette proteins which are involved in ATP release. Neuroreport 13: 1789-1792. Ballerini P, Rathbone MP, Di Iorio P, Renzetti A, Giuliani P, et al. 1996. Rat astroglial P2Z (P2X7) receptors regulate intracellular calcium and purine release. Neuroreport 7: 2533-2537. Barclay J, Patel S, Dorn G, Wotherspoon G, Moffatt S, et al. 2002. Functional downregulation of P2X3 receptor subunit in rat sensory neurons reveals a significant role in chronic neuropathic and inflammatory pain. J Neurosci 22: 8139-8147. Bardoni R, Goldstein PA, Lee CJ, Gu JG, MacDermott AB. 1997. ATP P2X receptors mediate fast synaptic transmission in the dorsal horn of the rat spinal cord. J Neurosci 17: 5297-5304. Barnard EA, Simon J. 2001. An elusive receptor is finally caught: P2Y(120 ), an important drug target in platelets. Trends Pharmacol Sci 22: 388-391. Bennett GC, Boarder MR. 2000. The effect of nucleotides and adenosine on stimulus‐evoked glutamate release from rat brain cortical slices. Br J Pharmacol 131: 617-623.

ATP‐mediated signaling in the nervous system Bianco F, Fumagalli M, Pravettoni E, D’Ambrosi N, Volonte C, et al. 2005. Pathophysiological roles of extracellular nucleotides in glial cells: Differential expression of purinergic receptors in resting and activated microglia. Brain Res Brain Res Rev 48: 144-156. Boarder MR, Webb TE. 2001. P2Y receptors: Structure and function. In: Handbook of experimental pharmacology: Purinergic and pyrimidiniergic signalling Vol. 151/I, Abbracchio MP, Williams M, editors. Berlin: Springer; pp. 65-89. Boehm S. 1999. ATP stimulates sympathetic transmitter release via presynaptic P2X purinoceptors. J Neurosci 19: 737-746. Boehm S, Kubista H. 2002. Fine tuning of sympathetic transmitter release via ionotropic and metabotropic presynaptic receptors. Pharmacol Rev 54: 43-99. Bonan CD, Walz R, Pereira GS, Worm PV, Battastini AM, et al. 2000. Changes in synaptosomal ectonucleotidase activities in two rat models of temporal lobe epilepsy. Epilepsy Res 39: 229-238. Bowser DN, Khakh BS. 2004. ATP excites interneurons and astrocytes to increase synaptic inhibition in neuronal networks. J Neurosci 24: 8606-8620. Bradbury EJ, Burnstock G, McMahon SB. 1998. The expression of P2X3 purinoreceptors in sensory neurons: Effects of axotomy and glial‐derived neurotrophic factor. Mol Cell Neurosci 12: 256-268. Braun N, Lenz C, Gillardon F, Zimmermann M, Zimmermann H. 1997. Focal cerebral ischemia enhances glial expression of ecto‐50 ‐nucleotidase. Brain Res 766: 213-226. Braun N, Zhu Y, Krieglstein J, Culmsee C, Zimmermann H. 1998. Upregulation of the enzyme chain hydrolyzing extracellular ATP after transient forebrain ischemia in the rat. J Neurosci 18: 4891-4900. Brough D, Le Feuvre RA, Iwakura Y, Rothwell NJ. 2002. Purinergic (P2X7) receptor activation of microglia induces cell death via an interleukin‐1‐independent mechanism. Mol Cell Neurosci 19: 272-280. Brown SG, Townsend‐Nicholson A, Jacobson KA, Burnstock G, King BF. 2002. Heteromultimeric P2X(1/2) receptors show a novel sensitivity to extracellular pH. J Pharmacol Exp Ther 300: 673-680. Burnstock G. 1972. Purinergic nerves. Pharmacol Rev 24: 509-581. Burnstock G. 2001. Purine‐mediated signalling in pain and visceral perception. Trends Pharmacol Sci 22: 182-188. Burnstock G. 2006. Purinergic P2 receptors as targets for novel analgesics. Pharmacol Ther 110(3): 433-454. Burnstock G, Wood JN. 1996. Purinergic receptors: Their role in nociception and primary afferent neurotransmission. Curr Opin Neurobiol 6: 526-532.

10

Calvert JA, Evans RJ. 2004. Heterogeneity of P2X receptors in sympathetic neurons: Contribution of neuronal P2X1 receptors revealed using knockout mice. Mol Pharmacol 65: 139-148. Cass CE, Young JD, Baldwin SA, Cabrita MA, Graham KA, et al. 1999. Nucleoside transporters of mammalian cells. Pharm Biotechnol 12: 313-352. Chakfe Y, Seguin R, Antel JP, Morissette C, Malo D, et al. 2002. ADP and AMP induce interleukin‐1beta release from microglial cells through activation of ATP‐primed P2X7 receptor channels. J Neurosci 22: 3061-3069. Chambers JK, Macdonald LE, Sarau HM, Ames RS, Freeman K, et al. 2000. A G protein‐coupled receptor for UDP‐ glucose. J Biol Chem 275: 10767-10771. Chessell IP, Michel AD, Humphrey PP. 1997. Properties of the pore‐forming P2X7 purinoceptor in mouse NTW8 microglial cells. Br J Pharmacol 121: 1429-1437. Chessell IP, Hatcher JP, Bountra C, Michel AD, Hughes JP, et al. 2005. Disruption of the P2X7 purinoceptor gene abolishes chronic inflammatory and neuropathic pain. Pain 114: 386-396. Chizh BA, Illes P. 2001. P2X receptors and nociception. Pharmacol Rev 53: 553-568. Choi HB, Hong SH, Ryu JK, Kim SU, McLarnon JG. 2003. Differential activation of subtype purinergic receptors modulates Ca(2þ) mobilization and COX‐2 in human microglia. Glia 43: 95-103. Cockayne DA, Dunn PM, Zhong Y, Rong W, Hamilton SG, et al. 2005. P2X2 knockout mice and P2X2/P2X3 double knockout mice reveal a role for the P2X2 receptor subunit in mediating multiple sensory effects of ATP. J Physiol 567: 621-639. Cockayne DA, Hamilton SG, Zhu QM, Dunn PM, Zhong Y, et al. 2000. Urinary bladder hyporeflexia and reduced pain‐ related behaviour in P2X3‐deficient mice. Nature 407: 1011-1015. Coco S, Calegari F, Pravettoni E, Pozzi D, Taverna E, et al. 2003. Storage and release of ATP from astrocytes in culture. J Biol Chem 278: 1354-1362. Collier HO, James GW, Schneider C. 1966. Antagonism by aspirin and fenamates of bronchoconstriction and nociception induced by adenosine‐50 ‐triphosphate. Nature 212: 411-412. Collo G, Neidhart S, Kawashima E, Kosco‐Vilbois M, North RA, et al. 1997. Tissue distribution of the P2X7 receptor. Neuropharmacology 36: 1277-1283. Collo G, North RA, Kawashima E, Merlo‐Pich E, Neidhart S, et al. 1996. Cloning of P2X5 and P2X6 receptors and the distribution and properties of an extended family of ATP‐gated ion channels. J Neurosci 16: 24952507.

247

248

10

ATP‐mediated signaling in the nervous system

Communi D, Gonzalez NS, Detheux M, Brezillon S, Lannoy V, et al. 2001. Identification of a novel human ADP receptor coupled to G(i). J Biol Chem 276: 41479-41485. Cook SP, McCleskey EW. 2002. Cell damage excites nociceptors through release of cytosolic ATP. Pain 95: 41-47. Cunha RA. 2001. Adenosine as a neuromodulator and as a homeostatic regulator in the nervous system: Different roles, different sources and different receptors. Neurochem Int 38: 107-125. Cunha RA, Ribeiro JA. 2000a. ATP as a presynaptic modulator. Life Sci 68: 119-137. Cunha RA, Ribeiro JA. 2000b. Purinergic modulation of [(3)H]GABA release from rat hippocampal nerve terminals. Neuropharmacology 39: 1156-1167. Cunha RA, Ribeiro JA, Sebastiao AM. 1994. Purinergic modulation of the evoked release of [3H]acetylcholine from the hippocampus and cerebral cortex of the rat: Role of the ectonucleotidases. Eur J Neurosci 6: 33-42. Cunha RA, Sebastiao AM, Ribeiro JA. 1992. Ecto‐50 ‐nucleotidase is associated with cholinergic nerve terminals in the hippocampus but not in the cerebral cortex of the rat. J Neurochem 59: 657-666. Cunha RA, Sebastiao AM, Ribeiro JA. 1998. Inhibition by ATP of hippocampal synaptic transmission requires localized extracellular catabolism by ecto‐nucleotidases into adenosine and channeling to adenosine A1 receptors. J Neurosci 18: 1987-1995. Darby M, Kuzmiski JB, Panenka W, Feighan D, MacVicar BA. 2003. ATP released from astrocytes during swelling activates chloride channels. J Neurophysiol 89: 1870-1877. Deuchars SA, Atkinson L, Brooke RE, Musa H, Milligan CJ, et al. 2001. Neuronal P2X7 receptors are targeted to presynaptic terminals in the central and peripheral nervous systems. J Neurosci 21: 7143-7152. Dorn G, Patel S, Wotherspoon G, Hemmings‐Mieszczak M, Barclay J, et al. 2004. siRNA relieves chronic neuropathic pain. Nucleic Acids Res 32: e49. Drury AN, Szent‐Gyo¨rgyi A. 1929. The physiological action of adenine compounds with especial reference to their action on the mammalian heart. J Physiol (Lond) 68: 214-237. Dulla CG, Dobelis P, Pearson T, Frenguelli BG, Staley KJ, et al. 2005. Adenosine and ATP link PCO2 to cortical excitability via pH. Neuron 48: 1011-1023. Dunn PM, Zhong Y, Burnstock G. 2001. P2X receptors in peripheral neurons. Prog Neurobiol 65: 107-134. Dunwiddie TV, Diao L, Proctor WR. 1997. Adenine nucleotides undergo rapid, quantitative conversion to adenosine in the extracellular space in rat hippocampus. J Neurosci 17: 7673-7682. Edwards FA, Gibb AJ, Colquhoun D. 1992. ATP receptor‐ mediated synaptic currents in the central nervous system. Nature 359: 144-147.

Egan TM, Khakh BS. 2004. Contribution of calcium ions to P2X channel responses. J Neurosci 24: 3413-3420. Evans RJ, Derkach V, Surprenant A. 1992. ATP mediates fast synaptic transmission in mammalian neurons. Nature 357: 503-505. Fellin T, Pozzan T, Carmignoto G. 2006. Purinergic receptors mediate two distinct glutamate release pathways in hippocampal astrocytes. J Biol Chem. 281: 4274-4284. Ferrari D, Chiozzi P, Falzoni S, Hanau S, Di Virgilio F. 1997c. Purinergic modulation of interleukin‐1 beta release from microglial cells stimulated with bacterial endotoxin. J Exp Med 185: 579-582. Ferrari D, Chiozzi P, Falzoni S, Dal Susino M, Collo G, et al. 1997a. ATP‐mediated cytotoxicity in microglial cells. Neuropharmacology 36: 1295-1301. Ferrari D, Chiozzi P, Falzoni S, Dal Susino M, Melchiorri L, et al. 1997b. Extracellular ATP triggers IL‐1 beta release by activating the purinergic P2Z receptor of human macrophages. J Immunol 159: 1451-1458. Ferrari D, Los M, Bauer MK, Vandenabeele P, Wesselborg S, et al. 1999. P2Z purinoreceptor ligation induces activation of caspases with distinct roles in apoptotic and necrotic alterations of cell death. FEBS Lett 447: 71-75. Ferrari D, Villalba M, Chiozzi P, Falzoni S, Ricciardi‐Castagnoli P, et al. 1996. Mouse microglial cells express a plasma membrane pore gated by extracellular ATP. J Immunol 156: 1531-1539. Fields RD, Stevens B. 2000. ATP: An extracellular signaling molecule between neurons and glia. Trends Neurosci 23: 625-633. Franke H, Illes P. 2006. Involvement of P2 receptors in the growth and survival of neurons in the CNS. Pharmacol Ther 109(3): 297-324. Galligan JJ, Bertrand PP. 1994. ATP mediates fast synaptic potentials in enteric neurons. J Neurosci 14: 7563-7571. Gendron FP, Chalimoniuk M, Strosznajder J, Shen S, Gonzalez FA, et al. 2003. P2X7 nucleotide receptor activation enhances IFN gamma‐induced type II nitric oxide synthase activity in BV‐2 microglial cells. J Neurochem 87: 344-352. Gerevich Z, Borvendeg SJ, Schroder W, Franke H, Wirkner K, et al. 2004. Inhibition of N‐type voltage‐activated calcium channels in rat dorsal root ganglion neurons by P2Y receptors is a possible mechanism of ADP‐induced analgesia. J Neurosci 24: 797-807. Gomes P, Srinivas SP, Van Driessche W, Vereecke J, Himpens B. 2005. ATP release through connexin hemichannels in corneal endothelial cells. Invest Ophthalmol Vis Sci 46: 1208-1218. Gordon GR, Baimoukhametova DV, Hewitt SA, Rajapaksha WR, Fisher TE, et al. 2005. Norepinephrine triggers release of glial ATP to increase postsynaptic efficacy. Nat Neurosci 8: 1078-1086.

ATP‐mediated signaling in the nervous system Gourine AV. 2005. On the peripheral and central chemoreception and control of breathing: An emerging role of ATP. J Physiol 568: 715-724. Gourine AV, Llaudet E, Dale N, Spyer KM. 2005a. ATP is a mediator of chemosensory transduction in the central nervous system. Nature 436: 108-111. Gourine AV, Llaudet E, Dale N, Spyer KM. 2005b. Release of ATP in the ventral medulla during hypoxia in rats: Role in hypoxic ventilatory response. J Neurosci 25: 1211-1218. Griffiths M, Yao SY, Abidi F, Phillips SE, Cass CE, et al. 1997. Molecular cloning and characterization of a nitrobenzylthioinosine‐insensitive (ei) equilibrative nucleoside transporter from human placenta. Biochem J 328(Pt 3): 739-743. Grondal EJ, Janetzko A, Zimmermann H. 1988. Monospecific antiserum against 50 ‐nucleotidase from Torpedo electric organ: Immunocytochemical distribution of the enzyme and its association with Schwann cell membranes. Neuroscience 24: 351-363. Gu JG, MacDermott AB. 1997. Activation of ATP P2X receptors elicits glutamate release from sensory neuron synapses. Nature 389: 749-753. Guo LH, Trautmann K, Schluesener HJ. 2005. Expression of P2X4 receptor by lesional activated microglia during formalin‐induced inflammatory pain. J Neuroimmunol 163: 120-127. Guthrie PB, Knappenberger J, Segal M, Bennett MV, Charles AC, et al. 1999. ATP released from astrocytes mediates glial calcium waves. J Neurosci 19: 520-528. Haas S, Brockhaus J, Verkhratsky A, Kettenmann H. 1996. ATP‐induced membrane currents in ameboid microglia acutely isolated from mouse brain slices. Neuroscience 75: 257-261. Hamann M, Attwell D. 1996. Non‐synaptic release of ATP by electrical stimulation in slices of rat hippocampus, cerebellum and habenula. Eur J Neurosci 8: 1510-1515. Harden TK, Lazarowski ER, Boucher RC. 1997. Release, metabolism and interconversion of adenine and uridine nucleotides: Implications for G protein‐coupled P2 receptor agonist selectivity. Trends Pharmacol Sci 18: 43-46. Hide I, Tanaka M, Inoue A, Nakajima K, Kohsaka S, et al. 2000. Extracellular ATP triggers tumor necrosis factor‐ alpha release from rat microglia. J Neurochem 75: 965-972. Hollopeter G, Jantzen HM, Vincent D, Li G, England L, et al. 2001. Identification of the platelet ADP receptor targeted by antithrombotic drugs. Nature 409: 202-207. Holton P. 1959. The liberation of adenosine triphosphate on antidromic stimulation of sensory nerves. J Physiol 145: 494-504. Honore P, Kage K, Mikusa J, Watt AT, Johnston JF, et al. 2002. Analgesic profile of intrathecal P2X(3) antisense oligonucleotide treatment in chronic inflammatory and neuropathic pain states in rats. Pain 99: 11-19.

10

Hugel S, Schlichter R. 2000. Presynaptic P2X receptors facilitate inhibitory GABAergic transmission between cultured rat spinal cord dorsal horn neurons. J Neurosci 20: 2121-2130. Illes P, Alexandre Ribeiro J. 2004. Molecular physiology of P2 receptors in the central nervous system. Eur J Pharmacol 483: 5-17. Illes P, Klotz K‐N, Lohse MJ. 2000. Signaling by extracellular nucleotides and nucleosides. Naunyn Schmiedeberg’s Arch Pharmacol 362: 295-299. Inoue K. 2002. Microglial activation by purines and pyrimidines. Glia 40: 156-163. Inoue K, Tsuda M, Koizumi S. 2004. ATP‐ and adenosine‐ mediated signaling in the central nervous system: Chronic pain and microglia: Involvement of the ATP receptor P2X4. J Pharmacol Sci 94: 112-114. Inoue K, Koizumi S, Ueno S, Kita A, Tsuda M. 1999. The functions of ATP receptors in the synaptic transmission in the hippocampus. Prog Brain Res 120: 193-206. Inoue K, Nakajima K, Morimoto T, Kikuchi Y, Koizumi S, et al. 1998. ATP stimulation of Ca2þ‐dependent plasminogen release from cultured microglia. Br J Pharmacol 123: 1304-1310. James S, Richardson PJ. 1993. Production of adenosine from extracellular ATP at the striatal cholinergic synapse. J Neurochem 60: 219-227. Jo YH, Role LW. 2002. Coordinate release of ATP and GABA at in vitro synapses of lateral hypothalamic neurons. J Neurosci 22: 4794-4804. Jones CA, Vial C, Sellers LA, Humphrey PP, Evans RJ, et al. 2004. Functional regulation of P2X6 receptors by N‐linked glycosylation: Identification of a novel alpha beta‐methylene ATP‐sensitive phenotype. Mol Pharmacol 65: 979-985. Juranyi Z, Orso E, Janossy A, Szalay KS, Sperlagh B, et al. 1997. ATP and [3H]noradrenaline release and the presence of ecto‐Ca(2þ)‐ATPases in the capsule‐glomerulosa fraction of the rat adrenal gland. J Endocrinol 153: 105114. Kanjhan R, Housley GD, Burton LD, Christie DL, Kippenberger A, et al. 1999. Distribution of the P2X2 receptor subunit of the ATP‐gated ion channels in the rat central nervous system. J Comp Neurol 407: 11-32. Kato F, Shigetomi E. 2001. Distinct modulation of evoked and spontaneous EPSCs by purinoceptors in the nucleus tractus solitarii of the rat. J Physiol 530: 469-486. Kegel B, Braun N, Heine P, Maliszewski CR, Zimmermann H. 1997. An ecto‐ATPase and an ecto‐ATP diphosphohydrolase are expressed in rat brain. Neuropharmacology 36: 1189-1200. Kennedy C. 2001. The role of purines in the peripheral nervous system. In: Purinergic and pyrimidinergic signalling I. Molecular, Nervous and Urogenitary sytem function,

249

250

10

ATP‐mediated signaling in the nervous system

Vol. 151. Abbracchio MP, Williams M, editors. Berlin: Springer; pp. 289-305. Kennedy C. 2005. P2X receptors: Targets for novel analgesics? Neuroscientist 11: 345-356. Kennedy C, Assis TS, Currie AJ, Rowan EG. 2003. Crossing the pain barrier: P2 receptors as targets for novel analgesics. J Physiol 553: 683-694. Khakh BS. 2001. Molecular physiology of P2X receptors and ATP signalling at synapses. Nat Rev Neurosci 2: 165-174. Khakh BS, Henderson G. 1998. ATP receptor‐mediated enhancement of fast excitatory neurotransmitter release in the brain. Mol Pharmacol 54: 372-378. Khakh BS, Gittermann D, Cockayne DA, Jones A. 2003. ATP modulation of excitatory synapses onto interneurons. J Neurosci 23: 7426-7437. Khakh BS, Burnstock G, Kennedy C, King BF, North A, Se´guela P, et al. 2001. International union of pharmacology. XXIV. Current status of the nomenclature and properties of P2x receptors and their subunits. Pharmacol Rev 53: 107118. King BF, Townsend‐Nicholson A, Wildman SS, Thomas T, Spyer KM, et al. 2000. Coexpression of rat P2X2 and P2X6 subunits in Xenopus oocytes. J Neurosci 20: 4871-4877. Kirkpatrick K, Burnstock G. 1987. Sympathetic nerve‐ mediated release of ATP from the guinea‐pig vas deferens is unaffected by reserpine. Eur J Pharmacol 138: 207-214. Kittner H, Krugel U, Illes P. 2001. The purinergic P2 receptor antagonist pyridoxalphosphate‐6‐azophenyl‐20 40 ‐ disulphonic acid prevents both the acute locomotor effects of amphetamine and the behavioural sensitization caused by repeated amphetamine injections in rats. Neuroscience 102: 241-243. Kittner H, Krugel U, Hoffmann E, Illes P. 2000. Effects of intra‐accumbens injection of 2‐methylthio ATP: A combined open field and electroencephalographic study in rats. Psychopharmacology (Berl) 150: 123-131. Kittner H, Krugel U, Hoffmann E, Illes P. 2004. Modulation of feeding behaviour by blocking purinergic receptors in the rat nucleus accumbens: A combined microdialysis, electroencephalographic and behavioural study. Eur J Neurosci 19: 396-404. Kittner H, Franke H, Fischer W, Schultheis N, Krugel U, et al. 2003. Stimulation of P2Y1 receptors causes anxiolytic‐like effects in the rat elevated plus‐maze: Implications for the involvement of P2Y1 receptor‐mediated nitric oxide production. Neuropsychopharmacology 28: 435-444. Knott TK, Velazquez‐Marrero C, Lemos JR. 2005. ATP elicits inward currents in isolated vasopressinergic neurohypophysial terminals via P2X2 and P2X3 receptors. Pflugers Arch 450: 381-389.

Koch H, Kugelgen I, Starke K. 1997. P2‐receptor‐mediated inhibition of noradrenaline release in the rat hippocampus. Naunyn Schmiedebergs Arch Pharmacol 355: 707-715. Koizumi S, Inoue K. 1997. Inhibition by ATP of calcium oscillations in rat cultured hippocampal neurones. Br J Pharmacol 122: 51-58. Koizumi S, Fujishita K, Tsuda M, Shigemoto‐Mogami Y, Inoue K. 2003. Dynamic inhibition of excitatory synaptic transmission by astrocyte‐derived ATP in hippocampal cultures. Proc Natl Acad Sci USA 100: 11023-11028. Koles L, Furst S, Illes P. 2005. P2X and P2Y receptors as possible targets of therapeutic manipulations in CNS illnesses. Drug News Perspect 18: 85-101. Krugel U, Kittner H, Illes P. 1999. Adenosine 50 ‐triphosphate‐ induced dopamine release in the rat nucleus accumbens in vivo. Neurosci Lett 265: 49-52. Krugel U, Kittner H, Illes P. 2001. Mechanisms of adenosine 50 ‐triphosphate‐induced dopamine release in the rat nucleus accumbens in vivo. Synapse 39: 222-232. Kukley M, Stausberg P, Adelmann G, Chessell IP, Dietrich D. 2004. Ecto‐nucleotidases and nucleoside transporters mediate activation of adenosine receptors on hippocampal mossy fibers by P2X7 receptor agonist 20 ‐30 ‐O‐(4‐benzoylbenzoyl)‐ATP. J Neurosci 24: 7128-7139. Latini S, Pedata F. 2001. Adenosine in the central nervous system: Release mechanisms and extracellular concentrations. J Neurochem 79: 463-484. Le KT, Babinski K, Seguela P. 1998. Central P2X4 and P2X6 channel subunits coassemble into a novel heteromeric ATP receptor. J Neurosci 18: 7152-7159. Le KT, Paquet M, Nouel D, Babinski K, Seguela P. 1997. Primary structure and expression of a naturally truncated human P2X ATP receptor subunit from brain and immune system. FEBS Lett 418: 195-199. Lee CW, Jarvis SM. 1988. Nucleoside transport in rat cerebral‐ cortical synaptosomes. Evidence for two types of nucleoside transporters. Biochem J 249: 557-564. Lewis C, Neidhart S, Holy C, North RA, Buell G, et al. 1995. Coexpression of P2X2 and P2X3 receptor subunits can account for ATP‐gated currents in sensory neurons [see comments]. Nature 377: 432-435. Li C, Peoples RW, Lanthorn TH, Li ZW, Weight FF. 1999. Distinct ATP‐activated currents in different types of neurons dissociated from rat dorsal root ganglion. Neurosci Lett 263: 57-60. Llaudet E, Hatz S, Droniou M, Dale N. 2005. Microelectrode biosensor for real‐time measurement of ATP in biological tissue. Anal Chem 77: 3267-3273. MacDermott AB, Role LW, Siegelbaum SA. 1999. Presynaptic ionotropic receptors and the control of transmitter release. Annu Rev Neurosci 22: 443-485.

ATP‐mediated signaling in the nervous system McGehee DS, Role LW. 1996. Presynaptic ionotropic receptors. Curr Opin Neurobiol 6: 342-349. McLarnon JG. 2005. Purinergic mediated changes in Ca2þ mobilization and functional responses in microglia: Effects of low levels of ATP. J Neurosci Res 81: 349-356. Mendoza‐Fernandez V, Andrew RD, Barajas‐Lopez C. 2000. ATP inhibits glutamate synaptic release by acting at P2Y receptors in pyramidal neurons of hippocampal slices. J Pharmacol Exp Ther 293: 172-179. Moore DJ, Chambers JK, Wahlin JP, Tan KB, Moore GB, et al. 2001. Expression pattern of human P2Y receptor subtypes: A quantitative reverse transcription‐polymerase chain reaction study. Biochim Biophys Acta 1521: 107-119. Moran‐Jimenez MJ, Matute C. 2000. Immunohistochemical localization of the P2Y(1) purinergic receptor in neurons and glial cells of the central nervous system. Brain Res Mol Brain Res 78: 50-58. Nagy AK, Shuster TA, Delgado‐Escueta AV. 1989. Rat brain synaptosomal ATP:AMP‐phosphotransferase activity. J Neurochem 53: 1166-1172. Nakatsuka T, Gu JG. 2001. ATP P2X receptor‐mediated enhancement of glutamate release and evoked EPSCs in dorsal horn neurons of the rat spinal cord. J Neurosci 21: 6522-6531. Nakatsuka T, Tsuzuki K, Ling JX, Sonobe H, Gu JG. 2003. Distinct roles of P2X receptors in modulating glutamate release at different primary sensory synapses in rat spinal cord. J Neurophysiol 89: 3243-3252. Newman EA. 2003a. Glial cell inhibition of neurons by release of ATP. J Neurosci 23: 1659-1666. Newman EA. 2003b. New roles for astrocytes: Regulation of synaptic transmission. Trends Neurosci 26: 536-542. Nicholas RA. 2001. Identification of the P2Y(12) receptor: A novel member of the P2Y family of receptors activated by extracellular nucleotides. Mol Pharmacol 60: 416-420. Nicke A, Kerschensteiner D, Soto F. 2005. Biochemical and functional evidence for heteromeric assembly of P2X1 and P2X4 subunits. J Neurochem 92: 925-933. Nieber K, Poelchen W, Illes P. 1997. Role of ATP in fast excitatory synaptic potentials in locus coeruleus neurones of the rat. Br J Pharmacol 122: 423-430. Norenberg W, Langosch JM, Gebicke‐Haerter PJ, Illes P. 1994. Characterization and possible function of adenosine 50 ‐triphosphate receptors in activated rat microglia. Br J Pharmacol 111: 942-950. North RA, Surprenant A. 2000. Pharmacology of cloned P2X receptors. Annu Rev Pharmacol Toxicol 40: 563-580. Ogata T, Chuai M, Morino T, Yamamoto H, Nakamura Y, et al. 2003. Adenosine triphosphate inhibits cytokine release from lipopolysaccharide‐activated microglia via P2y receptors. Brain Res 981: 174-183.

10

Ohtani Y, Minami M, Satoh M. 2000. Expression of inducible nitric oxide synthase mRNA and production of nitric oxide are induced by adenosine triphosphate in cultured rat microglia. Neurosci Lett 293: 72-74. Okada M, Kawata Y, Murakami T, Wada K, Mizuno K, et al. 1999. Interaction between purinoceptor subtypes on hippocampal serotonergic transmission using in vivo microdialysis. Neuropharmacology 38: 707-715. Pankratov Y, Castro E, Miras‐Portugal MT, Krishtal O. 1998. A purinergic component of the excitatory postsynaptic current mediated by P2X receptors in the CA1 neurons of the rat hippocampus. Eur J Neurosci 10: 3898-3902. Pankratov Y, Lalo U, Krishtal O, Verkhratsky A. 2002. Ionotropic P2X purinoreceptors mediate synaptic transmission in rat pyramidal neurones of layer II/III of somato‐sensory cortex. J Physiol 542: 529-536. Pascual O, Casper KB, Kubera C, Zhang J, Revilla‐Sanchez R, et al. 2005. Astrocytic purinergic signaling coordinates synaptic networks. Science 310: 113-116. Papp L, Vizi ES, Sperlagh B. 2004b. Lack of ATP‐evoked GABA and glutamate release in the hippocampus of P2X7 receptor/ mice. Neuroreport 15: 2387-2391. Papp L, Balazsa T, Kofalvi A, Erdelyi F, Szabo G, et al. 2004a. P2X receptor activation elicits transporter‐mediated noradrenaline release from rat hippocampal slices. J Pharmacol Exp Ther 310: 973-980. Parvathenani LK, Tertyshnikova S, Greco CR, Roberts SB, Robertson B, et al. (2003) P2X7 mediates superoxide production in primary microglia and is up‐regulated in a transgenic mouse model of Alzheimer’s disease. J Biol Chem 278: 13309-13317. Patel MK, Khakh BS, Henderson G. 2001. Properties of native P2X receptors in rat trigeminal mesencephalic nucleus neurones: Lack of correlation with known, heterologously expressed P2X receptors. Neuropharmacology 40: 96-105. Pearson RA, Dale N, Llaudet E, Mobbs P. 2005. ATP released via gap junction hemichannels from the pigment epithelium regulates neural retinal progenitor proliferation. Neuron 46: 731-744. Petruska JC, Cooper BY, Gu JG, Rau KK, Johnson RD. 2000. Distribution of P2X1, P2X2, and P2X3 receptor subunits in rat primary afferents: Relation to population markers and specific cell types. J Chem Neuroanat 20: 141-162. Poelchen W, Sieler D, Wirkner K, Illes P. 2001. Co‐transmitter function of ATP in central catecholaminergic neurons of the rat. Neuroscience 102: 593-602. Queiroz G, Talaia C, Goncalves J. 2003. ATP modulates noradrenaline release by activation of inhibitory P2Y receptors and facilitatory P2X receptors in the rat vas deferens. J Pharmacol Exp Ther 307: 809-815.

251

252

10

ATP‐mediated signaling in the nervous system

Ralevic V, Burnstock G. 1998. Receptors for purines and pyrimidines. Pharmacol Rev 50: 413-492. Rampe D, Wang L, Ringheim GE. 2004. P2X7 receptor modulation of beta‐amyloid‐ and LPS‐induced cytokine secretion from human macrophages and microglia. J Neuroimmunol 147: 56-61. Rhee JS, Wang ZM, Nabekura J, Inoue K, Akaike N. 2000. ATP facilitates spontaneous glycinergic IPSC frequency at dissociated rat dorsal horn interneuron synapses. J Physiol 524 Pt 2: 471-483. Rodrigues RJ, Almeida T, Richardson PJ, Oliveira CR, Cunha RA. 2005. Dual presynaptic control by ATP of glutamate release via facilitatory P2X1, P2X2/3, and P2X3 and inhibitory P2Y1, P2Y2, and/or P2Y4 receptors in the rat hippocampus. J Neurosci 25: 6286-6295. Rogers M, Colquhoun LM, Patrick JW, Dani JA. 1997. Calcium flux through predominantly independent purinergic ATP and nicotinic acetylcholine receptors. J Neurophysiol 77: 1407-1417. Rong W, Gourine AV, Cockayne DA, Xiang Z, Ford AP, et al. 2003. Pivotal role of nucleotide P2X2 receptor subunit of the ATP‐gated ion channel mediating ventilatory responses to hypoxia. J Neurosci 23: 11315-11321. Ruan HZ, Burnstock G. 2003. Localisation of P2Y1 and P2Y4 receptors in dorsal root, nodose and trigeminal ganglia of the rat. Histochem Cell Biol 120: 415-426. Ruan HZ, Birder LA, de Groat WC, Tai C, Roppolo J, et al. 2005. Localization of P2X and P2Y receptors in dorsal root ganglia of the cat. J Histochem Cytochem 53: 1273-1282. Rubio M, Soto F. 2001. Distinct Localization of P2X receptors at excitatory postsynaptic specializations. J Neurosci 21: 641-653. Sanchez‐Nogueiro J, Marin‐Garcia P, Miras‐Portugal MT. 2005. Characterization of a functional P2X(7)‐like receptor in cerebellar granule neurons from P2X(7) knockout mice. FEBS Lett 579: 3783-3788. Sanz JM, Di Virgilio F. 2000. Kinetics and mechanism of ATP‐ dependent IL‐1 beta release from microglial cells. J Immunol 164: 4893-4898. Schoen SW, Ebert U, Loscher W. 1999. 50 ‐Nucleotidase activity of mossy fibers in the dentate gyrus of normal and epileptic rats. Neuroscience 93: 519-526. SchoenSW, Graeber MB, Reddington M, Kreutzberg GW. 1987. Light and electron microscopical immunocytochemistry of 50 ‐nucleotidase in rat cerebellum. Histochemistry 87: 107-113. Schwab JM, Guo L, Schluesener HJ. 2005. Spinal cord injury induces early and persistent lesional P2X4 receptor expression. J Neuroimmunol 163: 185-189. Schwiebert EM. 1999. ABC transporter‐facilitated ATP conductive transport. Am J Physiol 276: C1-8.

Shigetomi E, Kato F. 2004. Action potential‐independent release of glutamate by Ca2þ entry through presynaptic P2X receptors elicits postsynaptic firing in the brainstem autonomic network. J Neurosci 24: 3125-3135. Shinozuka K, Mizuno H, Nakamura K, Kunitomo M. 2002. Purinergic modulation of vascular sympathetic neurotransmission. Jpn J Pharmacol 88: 19-25. Silinsky EM, Gerzanich V, Vanner SM. 1992. ATP mediates excitatory synaptic transmission in mammalian neurones. Br J Pharmacol 106: 762-763. Sim JA, Young MT, Sung HY, North RA, Surprenant A. 2004. Reanalysis of P2X7 receptor expression in rodent brain. J Neurosci 24: 6307-6314. Smith DO. 1991. Sources of adenosine released during neuromuscular transmission in the rat. J Physiol 432: 343-354. Sneddon P, Westfall TD, Todorov LD, Todorova SM, Westfall DP, et al. 2000. The effect of P2 receptor antagonists and ATPase inhibition on sympathetic purinergic neurotransmission in the guinea‐pig isolated vas deferens. Br J Pharmacol 129: 1089-1094. Souslova V, Cesare P, Ding Y, Akopian AN, Stanfa L, et al. 2000. Warm‐coding deficits and aberrant inflammatory pain in mice lacking P2X3 receptors. Nature 407: 1015-1017. Sperla´gh B. Vizi ES. 1996. Neuronal synthesis, storage and release of ATP. Seminars in the Neurosciences 8: 175-186. Sperlagh B, Vizi ES. 1991. Effect of presynaptic P2 receptor stimulation on transmitter release. J Neurochem 56: 1466-1470. Sperlagh B, Vizi ES. 1992. Is the neuronal ATP release from guinea‐pig vas deferens subject to alpha 2‐adrenoceptor‐ mediated modulation? Neuroscience 51: 203-209. Sperlagh B, Vizi ES. 2000. Regulation of purine release. In: Handbook of experimental pharmacology, Vol. 151. Abbracchio MP, Williams M, editors. Springer; pp. 179-209. Sperlagh B, Andras I, Vizi S. 1997. Effect of subtype‐ specific Ca(2þ)‐antagonists and Ca(2þ)‐free media on the field stimulation‐evoked release of ATP and [3H]acetylcholine from rat habenula slices. Neurochem Res 22: 967-975. Sperlagh B, Erdelyi F, Szabo G, Vizi ES. 2000. Local regulation of [(3)H]‐noradrenaline release from the isolated guinea‐ pig right atrium by P(2X)‐receptors located on axon terminals. Br J Pharmacol 131: 1775-1783. Sperlagh B, Kittel A, Lajtha A, Vizi ES. 1995. ATP acts as fast neurotransmitter in rat habenula: Neurochemical and enzymecytochemical evidence. Neuroscience 66: 915-920. Sperlagh B, Magloczky Z, Vizi ES, Freund TF. 1998a. The triangular septal nucleus as the major source of ATP release in the rat habenula: A combined neurochemical and morphological study. Neuroscience 86: 1195-1207.

ATP‐mediated signaling in the nervous system Sperlagh B, Sershen H, Lajtha A, Vizi ES. 1998b. Co‐release of endogenous ATP and [3H]noradrenaline from rat hypothalamic slices: Origin and modulation by alpha2‐adrenoceptors. Neuroscience 82: 511-520. Sperlagh B, Mergl Z, Juranyi Z, Vizi ES, Makara GB. 1999. Local regulation of vasopressin and oxytocin secretion by extracellular ATP in the isolated posterior lobe of the rat hypophysis. J Endocrinol 160: 343-350. Sperlagh B, Szabo G, Erdelyi F, Baranyi M, Vizi ES. 2003. Homo‐ and heteroexchange of adenine nucleotides and nucleosides in rat hippocampal slices by the nucleoside transport system. Br J Pharmacol 139: 623-633. Sperlagh B, Kofalvi A, Deuchars J, Atkinson L, Milligan CJ, et al. 2002. Involvement of P2X7 receptors in the regulation of neurotransmitter release in the rat hippocampus. J Neurochem 81: 1196-1211. Spyer KM, Dale N, Gourine AV. 2004. ATP is a key mediator of central and peripheral chemosensory transduction. Exp Physiol 89: 53-59. Stout C, Goodenough DA, Paul DL. 2004. Connexins: Functions without junctions. Curr Opin Cell Biol 16: 507-512. Stout CE, Costantin JL, Naus CC, Charles AC. 2002. Intercellular calcium signaling in astrocytes via ATP release through connexin hemichannels. J Biol Chem 277: 10482-10488. Sun XP, Stanley EF. 1996. An ATP‐activated, ligand‐gated ion channel on a cholinergic presynaptic nerve terminal. Proc Natl Acad Sci USA 93: 1859-1863. Suzuki T, Hide I, Ido K, Kohsaka S, Inoue K, et al. 2004. Production and release of neuroprotective tumor necrosis factor by P2X7 receptor‐activated microglia. J Neurosci 24: 1-7. Terrian DM, Hernandez PG, Rea MA, Peters RI. 1989. ATP release, adenosine formation, and modulation of dynorphin and glutamic acid release by adenosine analogues in rat hippocampal mossy fiber synaptosomes. J Neurochem 53: 1390-1399. Thomas T, Spyer KM. 2000. ATP as a mediator of mammalian central CO2 chemoreception. J Physiol 523 Pt 2: 441-447. Thomas T, Ralevic V, Gadd CA, Spyer KM. 1999. Central CO2 chemoreception: A mechanism involving P2 purinoceptors localized in the ventrolateral medulla of the anaesthetized rat. J Physiol 517(Pt 3): 899-905. Thomas T, Ralevic V, Bardini M, Burnstock G, Spyer KM. 2001. Evidence for the involvement of purinergic signalling in the control of respiration. Neuroscience 107: 481-490. Thorn JA, Jarvis SM. 1996. Adenosine transporters. Gen Pharmacol 27: 613-620. Torres GE, Egan TM, Voigt MM. 1999. Hetero‐oligomeric assembly of P2X receptor subunits. Specificities exist with regard to possible partners. J Biol Chem 274: 6653-6659.

10

Torres GE, Haines WR, Egan TM, Voigt MM. 1998. Co‐ expression of P2X1 and P2X5 receptor subunits reveals a novel ATP‐gated ion channel. Mol Pharmacol 54: 989-993. Trendelenburg AU, Bultmann R. 2000. P2 receptor‐mediated inhibition of dopamine release in rat neostriatum. Neuroscience 96: 249-252. Tsuda M, Shigemoto‐Mogami Y, Koizumi S, Mizokoshi A, Kohsaka S, et al. 2003. P2X4 receptors induced in spinal microglia gate tactile allodynia after nerve injury. Nature 424: 778-783. Tsuzuki K, Ase A, Seguela P, Nakatsuka T, Wang CY, et al. 2003. TNP‐ATP‐resistant P2X ionic current on the central terminals and somata of rat primary sensory neurons. J Neurophysiol 89: 3235-3242. Valera S, Hussy N, Evans RJ, Adami N, North RA, et al. 1994. A new class of ligand‐gated ion channel defined by P2x receptor for extracellular ATP. Nature 371: 516-519. Verderio C, Matteoli M. 2001. ATP mediates calcium signaling between astrocytes and microglial cells: Modulation by IFN‐gamma. J Immunol 166: 6383-6391. Visentin S, Renzi M, Frank C, Greco A, Levi G. 1999. Two different ionotropic receptors are activated by ATP in rat microglia. J Physiol 519 Pt 3: 723-736. Vizi ES. 2000. Role of high‐affinity receptors and membrane transporters in nonsynaptic communication and drug action in the central nervous system. Pharmacol Rev 52: 63-89. Vizi ES, Sperlagh B. 1999. Receptor‐ and carrier‐mediated release of ATP of postsynaptic origin: Cascade transmission. Prog Brain Res 120: 159-169. Vizi ES, Sperlagh B, Baranyi M. 1992. Evidence that ATP released from the postsynaptic site by noradrenaline, is involved in mechanical responses of guinea‐pig vas deferens: Cascade transmission. Neuroscience 50: 455-465. Vizi ES, Nitahara K, Sato K, Sperlagh B. 2000. Stimulation‐ dependent release, breakdown, and action of endogenous ATP in mouse hemidiaphragm preparation: The possible role of ATP in neuromuscular transmission. J Auton Nerv Syst 81: 278-284. Vizi ES, Liang SD, Sperlagh B, Kittel A, Juranyi Z. 1997. Studies on the release and extracellular metabolism of endogenous ATP in rat superior cervical ganglion: Support for neurotransmitter role of ATP. Neuroscience 79: 893-903. Vlaskovska M, Kasakov L, Rong W, Bodin P, Bardini M, et al. 2001. P2X3 knock‐out mice reveal a major sensory role for urothelially released ATP. J Neurosci 21: 5670-5677. von Kugelgen I, Wetter A. 2000. Molecular pharmacology of P2Y‐receptors. Naunyn Schmiedebergs Arch Pharmacol 362: 310-323. von Kugelgen I, Koch H, Starke K. 1997. P2‐receptor‐ mediated inhibition of serotonin release in the rat brain cortex. Neuropharmacology 36: 1221-1227.

253

254

10

ATP‐mediated signaling in the nervous system

von Kugelgen I, Spath L, Starke K. 1994. Evidence for P2‐ purinoceptor‐mediated inhibition of noradrenaline release in rat brain cortex. Br J Pharmacol 113: 815-822. Vulchanova L, Arvidsson U, Riedl M, Wang J, Buell G, et al. 1996. Differential distribution of two ATP‐gated channels (P2X receptors) determined by immunocytochemistry. Proc Natl Acad Sci USA 93: 8063-8067. Vulchanova L, Riedl MS, Shuster SJ, Buell G, Surprenant A, et al. 1997. Immunohistochemical study of the P2X2 and P2X3 receptor subunits in rat and monkey sensory neurons and their central terminals. Neuropharmacology 36: 1229-1242. Walter L, Dinh T, Stella N. 2004. ATP induces a rapid and pronounced increase in 2‐arachidonoylglycerol production by astrocytes, a response limited by monoacylglycerol lipase. J Neurosci 24: 8068-8074. Walz W, Ilschner S, Ohlemeyer C, Banati R, Kettenmann H. 1993. Extracellular ATP activates a cation conductance and a Kþ conductance in cultured microglial cells from mouse brain. J Neurosci 13: 4403-4411. Wang TF, Guidotti G. 1998. Widespread expression of ecto‐ apyrase (CD39) in the central nervous system. Brain Res 790: 318-322. Wang TF, Rosenberg PA, Guidotti G. 1997. Characterization of brain ecto‐apyrase: Evidence for only one ecto‐apyrase (CD39) gene. Brain Res Mol Brain Res 47: 295-302. Wang Y, Roman R, Lidofsky SD, Fitz JG. 1996. Autocrine signaling through ATP release represents a novel mechanism for cell volume regulation. Proc Natl Acad Sci USA 93: 12020-12025. Wang ZM, Katsurabayashi S, Rhee JS, Brodwick M, Akaike N. 2001. Substance P abolishes the facilitatory effect of ATP on spontaneous glycine release in neurons of the trigeminal nucleus pars caudalis. J Neurosci 21: 2983-2991. Watano T, Calvert JA, Vial C, Forsythe ID, Evans RJ. 2004. P2x receptor subtype-specific modulation of excitatory and inhibitory synaptic inputs in the brainstem. J Physiol 558: 745-757. Wieraszko A, Goldsmith G, Seyfried TN. 1989. Stimulation‐ dependent release of adenosine triphosphate from hippocampal slices. Brain Res 485: 244-250. Wirkner K, Ko¨falvi A, Fischer W, Gu¨nther A, Heike F, et al. 2005. Supersensitivity of P2X7 receptors in cerebrocortical

cell cultures after in vitro ischemia. J Neurochem 95: 14211437. Witting A, Walter L, Wacker J, Moller T, Stella N. 2004. P2X7 receptors control 2‐arachidonoylglycerol production by microglial cells. Proc Natl Acad Sci USA 101: 3214-3219. Xiang Z, Burnstock G. 2005. Expression of P2X receptors on rat microglial cells during early development. Glia 52: 119-126. Yao ST, Barden JA, Lawrence AJ. 2001. On the immunohistochemical distribution of ionotropic P2X receptors in the nucleus tractus solitarius of the rat. Neuroscience 108: 673-685. Yao SY, Ng AM, Muzyka WR, Griffiths M, Cass CE, et al. 1997. Molecular cloning and functional characterization of nitrobenzylthioinosine (NBMPR)‐sensitive (es) and NBMPR‐ insensitive (ei) equilibrative nucleoside transporter proteins (rENT1 and rENT2) from rat tissues. J Biol Chem 272: 28423-28430. Yegutkin GG, Henttinen T, Samburski SS, Spychala J, Jalkanen S. 2002. The evidence for two opposite, ATP‐generating and ATP‐consuming, extracellular pathways on endothelial and lymphoid cells. Biochem J 367: 121-128. Yoshioka K, Saitoh O, Nakata H. 2001. Heteromeric association creates a P2Y‐like adenosine receptor. Proc Natl Acad Sci USA 98: 7617-7622. Zhang JM, Wang HK, Ye CQ, Ge W, Chen Y, et al. 2003. ATP released by astrocytes mediates glutamatergic activity‐ dependent heterosynaptic suppression. Neuron 40: 971982. Zhang YX, Yamashita H, Ohshita T, Sawamoto N, Nakamura S. 1995. ATP increases extracellular dopamine level through stimulation of P2Y purinoceptors in the rat striatum. Brain Res 691: 205-212. Zhang YX, Yamashita H, Ohshita T, Sawamoto N, Nakamura S. 1996. ATP induces release of newly synthesized dopamine in the rat striatum. Neurochem Int 28: 395-400. Zimmermann H. 2000. Extracellular metabolism of ATP and other nucleotides. Naunyn Schmiedebergs Arch Pharmacol 362: 299-309. Zimmermann H. 2001. Ecto‐nucleotidases. In: Handbook of experimental pharmacology: Purinergic and pyrimidinergic signalling, Vol. 151/1. Abbracchio MP, Williams M editors. Berlin: Springer; pp. 209-251.

11

Adenosine Neuromodulation and Neuroprotection

R. A. Cunha

1 1.1 1.2 1.3 1.4 1.5 1.6

Physiological Roles of Adenosine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 256 Adenosine as a Homeostatic Modulator . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 256 Pharmacology and Localization of Adenosine Receptors in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . 258 Neurotransmission and Neuromodulation – Adenosine as a Neuromodulator . . . . . . . . . . . . . . . . . . 260 Source of Endogenous Extracellular Adenosine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 263 Role of A1 Receptors in the Control of Synaptic Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 264 A2A Receptors and Modulation of Synaptic Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 265

2 Roles of Adenosine in Neurodegenerative Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 266 2.1 Therapeutic Opportunities to Manage Neurodegenerative Diseases Targeting the Adenosine Modulation System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 266 2.2 A1 Receptors as Hurls for the Development of Neuronal Dysfunction . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267 2.3 Role of A2A Receptors in the Control of Neurodegeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268 2.4 A2A Receptor Antagonists as Novel Anti‐Parkinsonian Drugs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 2.5 Role of A2A Receptors in Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 3

#

Final Comments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271

2008 Springer ScienceþBusiness Media, LLC.

256

11

Adenosine neuromodulation and neuroprotection

Abstract: The adenosine moiety fulfils an important intracellular homeostatic role since it is part of molecules that play key roles in defining the status of all cells, namely energy charge (ATP), redox status (NADH), and cell division (SAH/SAM). But the signaling role of the adenosine molecule itself is restricted to a paracrine role, signaling metabolic imbalance of cells within a tissue. Apart from this general role common to most tissues in mammals, adenosine fulfils a particular role as a neuromodulator in the nervous system. This involves a predominant inhibitory effect operated by adenosine A1 receptors, which results from a combined presynaptic inhibition of the release of excitatory neurotransmitters together with a postsynaptic action leading to neuronal hyperpolarization and an ability to depress plasticity by inhibition of NMDA receptors and voltage‐sensitive calcium channels. Overall, this A1 receptor‐mediated inhibition is aimed at decreasing the noise of excitatory transmission in brain circuits. Adenosine can also activate facilitatory A2A receptors, which only come into play at higher frequency of nerve stimulation and are directed at selectively shutting down A1 receptor inhibition in stimulated synapses to aid implementing changes in synaptic efficiency. Therefore, the combined role of A1 and A2A receptors is designed to increase salience of information in brain circuits. Apart from this main physiological role, adenosine also plays a relevant role in controlling the demise of damage in noxious brain conditions. In fact, the inhibitory A1 receptors are able to curtail brain damage. They play a role at the onset of brain damage and function as a hurl that needs to be overcome to allow the development of brain damage. In parallel, in chronic noxious brain conditions, A2A receptors contribute for brain damage, especially when insidious damage to synapses initiates neurodegeneration. Hence, A2A receptor antagonists are now being explored as novel neuroprotective strategies to interfere with the initial processes of neurodegenerative conditions, such as Parkinson’s and Alzheimer’s diseases. List of Abbreviations: CCPA, 2‐Chloro‐N6‐cyclopentyladenosine; CGS21680, 2‐[p‐(2‐carbonyl‐ethyl)‐ phenylethylamino]‐50 ‐N‐ethylcarboxamidoadenosine; Cl‐IB‐MECA, 1‐[2‐Chloro‐6‐[[(3‐iodophenyl)methyl] amino]‐9H‐purin‐9‐yl]‐1‐deoxy‐N‐methyl‐b‐D‐ribofuranuronamide; CP‐68,247, 8‐chloro‐4‐cyclohexyl‐ amino‐1‐(trifluoromethyl)[1,2,4]triazolo[4,3‐a] quinoxaline; CSC, 8‐(3‐chlorostyryl)caffeine; DPCPX, 1,3‐ dipropyl‐8‐cyclopentylxanthine; IB‐MECA, N6‐(3‐iodobenzyl)adenosine‐50 ‐N‐methyluronamide; KF17837, 1,3‐dipropyl‐8‐(3,4‐dimethoxystyryl)‐7‐methylxanthine; MRE3008F20, 5‐[[(4‐methoxyphenyl)amino] carbonyl]amino‐8‐ethyl‐2‐(2‐furyl)‐pyrazolo[4,3‐e]1,2,4‐triazolo[1,5‐c]pyrimidine; MRS1220, 9‐chloro‐ 2‐(2‐furanyl)‐5‐[(phenylacetyl)amino][1,2,4]‐triazolo[1,5‐c]quinazoline; MRS1334, 1,4‐Dihydro‐2‐methyl‐ 6‐phenyl‐4‐(phenylethynyl)‐3,5‐pyridinedicarboxylic acid 3‐ethyl‐5‐[(3‐nitrophenyl)methyl] ester; MRS1754, N‐(4‐cyano‐phenyl)‐2‐[4‐(2,6‐dioxo‐1,3‐dipropyl‐2,3,4,5,6,7‐hexahydro‐1H‐purin‐8‐yl)‐phenoxy]acetamide; R‐PIA, R‐N6‐(phenylisopropyl)‐adenosine; SCH58261, 5‐amino‐2‐(2‐furyl)‐7‐phenylethyl‐pyrazolo[4,3‐e]‐ 1,2,4‐triazolo[1,5‐c]pyrimidine; ZM241385, 4‐(2‐[7‐amino‐2‐[2‐furyl]‐[1,2,4]triazolo[2,3‐a]{1,3,5}triazin‐ 5‐yl‐amino]ethyl)phenol

1

Physiological Roles of Adenosine

1.1 Adenosine as a Homeostatic Modulator The viability of an organism ultimately depends on the ability of its cells to maintain a ‘‘constant’’ metabolism adapted to the needs of the organism (regulation) and in harmony with its environment. On the other hand, each cell should be able to change its metabolism according to modifications of the environment (control). This implies the need of devising signaling mechanisms (modulators) and devices (receptors and transducing systems), which allow sensing changes in the environment and implementing these changes inside cells to adequate their functioning. Since mammals are composed by cells organized in tissues, there is also the need to coordinate the response of each cell within a tissue (paracrine signaling) and of each tissue within the organism (endocrine signaling, which is aided by global control systems, such as the nervous or immune-inflammatory systems). One main paracrine signaling system is operated by adenosine (reviewed in Cunha, 2001). Adenosine is a purine nucleoside which normally occurs in concentrations of tenths of nanomolar within virtually all

Adenosine neuromodulation and neuroprotection

11

eukaryotic cells. However, considerably higher concentrations of chemical derivatives of this nucleoside are present within cells (> Figure 11‐1): for instance, it is present in the form of S‐adenosylhomocysteine (SAH) and S‐adenosylmethionine (SAM), nicotanimide and adenine nucleotides (NADþ and NADPþ) and adenine nucleotides (AMP, ADP, and ATP). In this sense, the adenosine moiety is directly involved in the three fundamental systems that regulate the functioning of eukaryotic cells: cell cycle (SAH/SAM), redox potential (NADH/NADþ), and energy charge (ATP, ADP, and AMP). However, the signaling role of the adenosine molecule as such is reserved to a particular situation of paracrine signaling of energetic modification with a tissue. . Figure 11‐1 Adenosine derivatives are directly involved in the three fundamental systems that regulate the functioning of eukaryotic cells. In fact, S‐adenosylhomocysteine (SAH) and S‐adenosylmethionine (SAM) control cell cycle, nicotanimide and adenine nucleotides (NADþ and NADPþ) define the redox state and adenine nucleotides (AMP, ADP, and ATP) define the energy status of cells. However, the signaling role of the adenosine molecule as such is reserved to a particular situation of paracrine signaling of energetic modification with a tissue

In a situation of energetic (dynamic) ‘‘equilibrium’’ between the generation and requirement of energy (ATP for the sake of simplicity), the concentration of this nucleotide is kept constant at a value of 3–10mM in different eukaryotic cells. In a situation of metabolic imbalance in a particular cell within a tissue, either because there is a shortage of oxidative potential or because there is an increased workload, the cell becomes metabolically stressed. This condition needs to be signaled to all neighboring cells in the tissue to enable them to rapidly adapt their metabolism to new environmental, potentially stressful conditions. Given the importance of the maintenance of tissue homeostasis, this signaling ought to be rapid, while the particular cell that first sensed the stressful change of environment is still beginning to become energetically imbalanced. This is possible, thanks to the difference in the extracellular concentration of ATP (circa 5mM) and of adenosine (circa 50nM). Hence, the concentration of adenosine will rise 100,000‐fold with a change of 1% of the concentration of ATP. This change in intracellular adenosine is converted to a gradient of extracellular adenosine around this cell since all eukaryotic cells are equipped with bidirectional nonconcentrative nucleoside (adenosine) transporters. These increased levels of intracellular adenosine can then activate the most abundant and ubiquitous adenosine receptor subtype, the A1 receptor, which has the ability of decreasing metabolic flow in eukaryotic cells (reviewed in Cunha, 2001) (> Figure 11‐2).

257

258

11

Adenosine neuromodulation and neuroprotection

. Figure 11‐2 Adenosine fulfils a paracrine role signalling situations of metabolic imbalance within tissues. When a stressful event occurs in a given cell (left part of the figure), ATP is used to attempt restoring the functioning of this cell. As a minor fraction of ATP is used, there is a massive formation of adenosine because of the near 250,000‐fold difference in their concentrations. This adenosine outflows from this cell through bidirectional and nonconcentrative nucleoside transporters (T). This adenosine gradient acts through adenosine A1 receptors (A1R) in neighboring cells where it decreases the metabolic rate allowing these cells to cope better with potential stressful stimuli that may reach them

This decrease of metabolic rate allows cells to cope better with stressful conditions and also increase their sensitivity to recruitment signals (by decreasing noise). This generic system of tissue paracrine signaling of metabolic imbalance has been exploited in several tissues as a signaling mechanism to coordinate tissue activity. This can be exemplified by the role of adenosine in the control of pacemaking in the heart (Shryock and Belardinelli, 1997), in the control of tubuloglomerular filtration rate in the kidney (Osswald et al., 1996), in the control of postprandial vasodilatation in the liver (Lautt, 1996), in the control of immune‐inflammatory reactivity to limit collateral damage (Sitkovsky et al., 2004), or in the control of the activity of neuronal circuits in the brain (reviewed in Dunwiddie and Masino, 2001). This homeostatic inhibitory effect of adenosine in the central nervous system is expected to be most relevant since the brain is equipped with a density of adenosine A1 receptors considerably greater than that of other tissues (reviewed in Fredholm et al., 2005). This is best illustrated by the fact that the consumption of caffeine, which only known mechanism of action at nontoxic concentrations is the antagonism of adenosine receptors (Fredholm et al., 1999), causes effects involving the modification of brain functioning (reviewed in Fredholm et al., 1999).

1.2 Pharmacology and Localization of Adenosine Receptors in the Brain Adenosine can act through the activation of four different adenosine receptors (A1, A2A, A2B, and A3). These receptors constitute the subfamily of P1 receptors to distinguish them from P2 receptors which are activated by adenosine‐50 ‐ and uridine‐50 ‐di‐ and tri‐phosphate (Ralevic and Burnstock, 1998). Adenosine receptors are G‐protein coupled receptors (see > Table 11‐1), which have limited sequence homology between each other in the different species where they were cloned (reviewed in Fredholm et al., 2005). The A1 receptor is the most abundantly expressed and its primary sequence is most conserved between species in the family. A1 Receptors couple to Gi/o proteins but their ability to inhibit neuronal activity may not only depend on their ability to inhibit cAMP accumulation and activate phospholipase C activity, but also to signal through b,g subunits of G proteins (Fredholm et al., 2005). A1 Receptors are widespread and abundantly expressed in the brain with a pre‐ and postsynaptic localization and with high levels in the cerebral cortex,

11

Adenosine neuromodulation and neuroprotection

. Table 11‐1 Represents the pharmacological properties of the most important adenosinergic ligands at the rat (r) and the human (h) receptors. Values are bold in columns of those receptors for which the ligand has the selectivity Agonist Adenosine

rA1

hA1 54

73 Inosine CCPA

8.1 0.4

R‐PIA CGS21680 IB‐MECA Cl‐IB‐MECA Antagonist DPCPX

3,100 54

rA1 0.73 0.3 0.6

SCH58261 caffeine ZM241385 CSC KF17387 CP‐68,247 MRS1334 MRS1220 MRE3008F20 MRS1754

rA2A 150

6,700 6.4 0.8 2.0 290 3.7 120 hA1 1.6 3.9

12,000 540

138 1,493 438 87 >100,000 305

3,900

22 56

hA2B 11,300

5,100

rA3

hA3 56

6,500

2,300 860 27

42 16 67

11,200a 88,800a

2,500 2,100

1.1 0.3

hA2A

340 682

130

50

4000

0.6

5,000b

10,000

1.2 8,100

2,400 1.7

rA2B

17,000

hA2B

13,000 31

0.67 61 17 41,561 >100,000 52 140 500 610

2,100 2.0

rA3

1.2 11

rA2A

1,100 400 17

rA2B

81

290 740 20,000

hA2A 960

hA3

190,000

80,000 270

3,850 >1,000

2.7 0.7 0.29 570

Ref. 1 2 1 3 4 4 4 11 4 4 Ref. 3 5 7 5 7 6 6 7 7 7 7 6 6 8 9 10

References: (1) Rank order of potency (EC50) values, in Chinese hamster ovary (CHO) cells transfected with human adenosine receptors (Fredholm et al., 2001b); (2) Rank order of potency (EC50) values, in Chinese hamster ovary (CHO) cells transfected with rat adenosine receptors (Yaar et al., 2004); (3) Binding affinities (Ki values) in rat brain cortical membranes and membranes of CHO cells expressing the human A1 receptor (Wittendorp et al., 2004); (4) Comparison of affinity of agonists at rat and human adenosine receptor subtypes (Ki values in nM). aEC50‐values (nM) for the agonist‐mediated stimulation of adenylyl cyclase activity in a membrane preparation (Klotz, 2000); (5) Binding affinity (Ki values) of antagonists at adenosine receptor subtypes. bInhibition of cAMP accummulation (Klotz, 2000); (6) Potency (KD values) of caffeine at rat and human adenosine receptor subtypes (Fredholm et al., 1999); (7) Potency of some adenosine receptor antagonists to displace CGS21680 from adenosine A2A receptors CHA from adenosine A1 receptors in rat striatum. Results are given as Ki values in nM (Fredholm and Lindstro¨m, 1999); (8) Binding affinity at human adenosine receptor subtypes in CHO cells (Ki values), (Varani et al., 2000). Not a useful ligand at rat receptors; (9) Binding affinity at human adenosine receptor subtypes in transfected HEK 293 cells (Ki values), (Kim et al., 2000); (10) Binding affinity at rat adenosine receptor subtypes in transfected HEK 293 cells (Ki values), (Kim et al., 2000); (11) Given are IC50 values, radioligands [3H]CHA (A1), [3H]NECA (A2A) (Hutchison

hippocampus, cerebellum, thalamus, brain stem, and spinal cord of rodents and Humans (see Fredholm et al., 2005). Adenosine A2A receptors are expressed at comparatively lower levels in the nervous system, with the exception of the basal ganglia, where they are expressed at exceptionally high levels in GABAergic medium spiny neurons (Svenningsson et al., 1999). Activation of A2A receptors has effects largely opposite

259

260

11

Adenosine neuromodulation and neuroprotection

to these of A1 receptors. Although they are mostly viewed as signaling through GS proteins, several neuronal effects of A2A receptors apparently involve other transducing systems (Fredholm et al., 2005), which still await to be clarified. Two other adenosine receptors have been cloned and pharmacologically characterized, namely A2B and A3 receptors. However, they have an expression lower than that of A1 and A2A receptors in the brain and a limited number of studies have so far explored their eventual role in controlling brain function (Fredholm et al., 2005). Adenosine is the primary full agonist at all four adenosine receptor subtypes. There are several adenosine receptor agonists which are mainly derivatives of adenosine with N6 and C2 substitutions of the adenine base and C5 substitutions of the ribose moiety (Fredholm et al., 2001). As illustrated in > Table 11‐1, these agonists effectively distinguish between A1, A2A, and A3 receptor mediated effects in in vitro experiments, whereas selective A2B receptor agonists await widespread acceptance, especially because this receptor has lower affinity for adenosine and for most agonists. The situation is less fortunate in vivo as the pharmacokinetics of these compounds have not been studied extensively. To be considered selective, the potency of a considered ligand between different adenosine receptors should ideally differ by at least two orders of magnitude. However, this is not always the case with adenosine receptor agonists, thus limiting their use for studying specific receptor function in vivo (Fredholm et al., 2001). There are no known endogenously produced adenosine receptor antagonists in mammals. However, there are several selective antagonists for the different adenosine receptors (see > Table 11‐1). The initial generation was constituted by derivatives of caffeine, the mostly widely consumed psychoactive drug (Fredholm et al., 1999). However, several different classes of compounds have been developed as pharmacological tools with appreciable selectivity for the different adenosine receptors (see > Table 11‐1). As for agonists, there are better and widespread used antagonists to manipulate A1 and A2A receptors. The A3 receptors display considerable difference in their profile of antagonist sensitivity between different species. Finally, the novel A2B receptor antagonists still await widespread validation. Hence, the combined density of the different adenosine receptors together with the availability of adequate phar