Morphological and Functional Characterization of the Neurotransmitter Gaba in Adult Rat Taste Buds

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MORPHOLOGICAL AND FUNCTIONAL CHARACTERIZATION OF THE NEUROTRANSMITTER GABA IN ADULT RAT TASTE BUDS

DISSERTATION

Presented in Partial Fulfillment of the Requirements for

the Degree Doctor of Philosophy in the Graduate

School of The Ohio State University

Yu Cao, B.M.

* * * * *

The Ohio State University

2006

Dissertation Committee: Approved by

Professor M. Scott Herness, Adviser ______

Professor Susan P. Travers Adviser Neuroscience Graduate Studies Program Professor Jackie D. Wood

ABSTRACT

γ-aminobutyric acid (GABA) is the major inhibitory neurotransmitter in the central nervous system (CNS). By activating the ionotropic GABAA receptor subtype or the metabotropic GABAB receptor subtype, GABA is thought to act as a neurotransmitter

or a neuromodulator to exert a wide variety of functions in the CNS including multiple

sensory systems. However, little is known regarding involvement of GABA in peripheral

gustatory signal processing in the taste buds.

GABA is among a broad array of neuroactive substances, which include several

neurotransmitters and neuropeptides, identified in mammalian taste buds. The present

study is the first to characterize the morphological features, distribution patterns and

functional consequences of GABAergic taste receptor cells (TRCs) in adult rat posterior

lingual epithelium. Subsets of TRCs were identified as the endogenous source of GABA

in taste buds of foliate or circumvallate papillae, as demonstrated by localization of

immunoreactivity to both GABA and the GABA synthetic enzyme, glutamate

decarboxylase (GAD). Morphologically, GABAergic TRCs resembled other TRCs with

known taste signaling functions. Double labeling immunofluorescent studies revealed

complex co-expression patterns of GAD with TRC population-specific protein markers

(i.e. taste-specific G-protein gustducin α subunit /Gα-gust, neural cell adhesion

ii molecule/NCAM and protein gene product 9.5/PGP 9.5), the presynaptic membrane-

specific protein marker synaptosomal-associated protein of 25 kDa (SNAP-25) and

neuropeptides (i.e. cholecystokinin/CCK and vasoactive intestinal polypeptide/VIP),

implying that GABAergic TRCs may have diverse functions in the taste buds.

GABA receptor subtypes, GABAA and GABAB, were also localized to subsets of

TRCs in rat foliate and circumvallate papillae, indicating that endogenously released

GABA may be able to modulate functions of TRCs by activating specific receptors. It appeared that the GABAA receptor α1 subunit, but not the α3 subunit, was present in a

group of cells with typical morphological features of TRCs expressing signal

transduction molecules, suggesting possible involvement of the GABAA receptor in

peripheral gustatory signal processing. The GABAB receptor R1 subunit- immunoreactivity was observed in a group of TRCs separate from the GAD-containing cells in the same bud, arguing a paracrine role for GABA by acting on the GABAB receptor. Interestingly, peptidergic TRCs seemed to receive differentially distributed modulatory input mediated by the GABAB receptor, as suggested by double labeling immunofluorescent studies showing co-localization of the GABAB receptor R1 subunit with the neuropeptide CCK, but not VIP.

Possible physiological effects of GABA in the taste buds were examined by patch clamp recordings of TRCs acutely dissociated from rat foliate and circumvallate papillae.

GABA and the GABAA receptor-specific agonists, muscimol and isoguvacine, all

enhanced isolated chloride currents recorded from TRCs in a dose-dependent manner,

consistent with GABAA receptor-mediated actions of GABA reported in the CNS. In addition, GABA and the GABAB receptor-specific agonist baclofen both elicited

iii increases of the inwardly rectifying potassium currents (Kir) recorded from TRCs. The

enhancing effect on Kir by baclofen could be blocked by the GABAB receptor antagonist

CGP 35348 and the G protein blocker GDP-βs, indicating that the G protein-coupled

GABAB receptor specifically mediated the effect. Therefore, GABA may modulate different electrical properties of TRC’s by acting on distinctive receptor subtypes.

In summary, GABA produced endogenously in the taste buds may modulate functions of various populations of TRCs by acting on specific receptor subtypes. The results of the present study add GABA to a growing list of recently characterized neuroactive substances in the taste buds that includes neurotransmitters and neuropeptides. These neuroactive substances mediating cell-to-cell communication within the taste buds are capable of shaping the final output of chemosensory TRCs and therefore may play important roles in peripheral gustatory signal processing.

Dedicated to my parents

ACKNOWLEDGMENTS

I cannot feel more grateful to my adviser, Dr. Scott Herness, for always being a

considerate mentor, for his incredible intellectual support, for his continuing passionate

encouragement, and for his great patience throughout the whole process that makes this

dissertation to become true.

I wish to thank Dr. Fangli Zhao, for never hesitating to share his academic and

technical expertise, for his invaluable input into this dissertation, and for his countless

helps both inside and outside the laboratory.

I also wish to thank Dr. Tiansheng Shen for her pioneering work that forms the

base of my dissertation project and for her long-lasting technical support.

I am grateful to all current and previous members of the laboratory, especially Dr.

Shaogang Lu and Tamara Kolli for their assistance at all times.

I appreciate all the kind helps from Dr. Georgia A. Bishop, Barbara Deiner and Dr.

James S. King.

I am greatly indebted to my husband, Dr. Jinbin Tian, who has been on my side

for almost eight happy years.

At last, I thank my parents and my twin sister from the bottom of my heart.

Without them supporting me unconditionally and helping me taking care of my daughter,

this dissertation would not be possible.

VITA

March 24, 1976 ………………………….. Born – Nei Mongol, P.R. China

1994 – 1999 ……………………………… Bachelor of Medicine, Peking University

Health Sciences Center.

1999 – present …………………………… Graduate Research Associate, the Ohio State

University

PUBLICATIONS

Research Publication

1. Wang Y, Huang C, Cao Y, Han JS. 1999. Synergistic analgesia between ketamine and clonidine in pain modulation. J Beijing Medical University 2. Wang Y, Huang C, Cao Y, Han JS. 1999. Repeated injection of low dose ketamine for the treatment of acute period of adjuvant arthritic inflammatory pain. Chinese J of Pain Medicine 3. Wang Y, Huang C, Cao Y, Han JS. 2000. Repeated administration of low dose ketamine for the treatment of monoarthritic pain in the rat. Life Sci. Jun 8;67(3):261-7 4. Cao Y, Herness MS. 2001. Localization and functional investigation of CREB in rat taste receptor cells. Soc. Neurosci. Abstr. 27. Program # 287.2 5. Cao Y, Shreffler C, Herness MS. 2002. Localization and functional investigation of the transcription factor CREB in rat taste receptor cells. NeuroReport 13:1321-25 6. Shen T, Kaya N, Zhao FL, Lu SG, Cao Y, and Herness S. 2005. Expression patterns of the neuropeptides VIP and CCK and the transduction molecules α-gustducin and T1R2 in rat taste receptor cells. Neuroscience 130(1): 229-38 7. Herness S, Zhao FL, Kaya N, Shen T, Lu SG, and Cao Y. 2005. Communication routes within the taste bud by neurotransmitters and neuropeptides. Chemical Senses. Jan 30; Suppl 1: i37-8

vii 8. Cao Y, Zhao FL, Herness MS. 2005. GABA as an inhibitory transmitter in the taste bud. Chemical Senses 30:A105-06 9. Zhao FL, Cao Y, Herness MS. 2005. Antagonistic actions of neuropeptides CCK and NPY on rat taste receptor cells. Chemical Senses 30:A101-02 10. Zhao FL, Shen T, Kaya N, Lu SG, Cao Y, Herness S. 2005. Expression, physiological action, and coexpression patterns of neuropeptide Y in rat taste-bud cells. Proc Natl Acad Sci U S A. 102(31):11100-5 11. Wang Y, Zhang Y, Wang W, Cao Y, Han JS. 2005. Effects of synchronous or asynchronous electroacupuncture stimulation with low versus high frequency on spinal opioid release and tail flick nociception. Exp Neurol. 192(1):156-62

12. Cao Y, Zhao FL, Herness MS. 2006. Further characterization of neuropeptides in rat taste receptor cells. Chemical Senses (in press)

FIELDS OF STUDY

Major Field: Neuroscience

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TABLE OF CONTENTS

Page Abstract ……………………………………………………………………………... ii Dedication …………………………………………………………………………... v Acknowledgments …………………………………………………………………... vi Vita ………………………………………………………………………………….. vii List of Tables ………………………………………………………………………... xi List of Figures ……………………………………………………………………….. xii Abbreviation ………………………………………………………………………….. xiv

Chapters:

1. Introduction ……………………………………………………………………… 1

1.1. Intrinsic neuromodulation in taste buds …………………………………….. 2 1.2. Communication between TRCs in taste bud ………………………………... 4 1.2.1. Neurotransmitters in TB: 5-HT, NA, glutamate and ATP ………... 6 1.2.2. Neuropeptides in TB: CCK and NPY …………………………….. 9 1.2.3. Other neuroactive substances in TB …………………………….. 10 1.3. GABA as a candidate for mediating neuromodulation in taste bud ………. 11 1.3.1. GABA synthesis ………………………………………………… 11 1.3.2. GABA transporters ……………………………………………… 12 1.3.3. GABA receptors ………………………………………………… 12 1.3.4. GABA in the gustatory system ………………………………….. 15

2. Methods ………………………………………………………………………… 19

2.1. Immunocytochemistry …………………………………………………….. 19 2.1.1. Tissue preparation ……………………………………………….. 19 2.1.2. Conventional immunocytochemistry protocol …………………... 19 2.1.3. TSA-amplified immunocytochemistry protocol ………………… 20 2.1.4. Double-labeling immunocytochemistry protocol ……………….. 21 2.1.5. Cell counting and data analysis ………………………………….. 24 2.2. Western blotting …………………………………………………………… 25 2.3. Patch clamp recording ……………………………………………………... 26 2.3.1. Dissociation of rat taste receptor cells …………………………... 26 2.3.2. Solutions for patch clamp recordings …………………………… 27

ix 2.3.3. Patch clamp recording procedures ………………………………. 28 3. Results ………………………………………………………………………….. 31

3.1. Distribution of GABAergic TRCs in rat posterior lingual epithelium ……. 32 3.1.1. Expression of GABA and the GABA synthetic enzyme GAD65/67 in TRCs …………………………………………………………. 32 3.1.2. Phenotypes of GABAergic TRCs with respects to TRC population- specific protein markers ………………………………………… 33 3.1.3. Co-localization of GAD65/67 with the presynaptic membrane- specific protein marker SNAP-25 ………………………………. 36 3.1.4. Co-localization of GAD65/67 with neuropeptides CCK and VIP.. 38 3.2. Distributions of GABA receptors in taste buds of rat posterior lingual epithelium …………………………………………………………………. 40 3.2.1. Expression of the GABAA receptor α1 subunit in TRCs …………... 40 3.2.2. Expression of the GABAB receptor R1 subunit in TRCs …………... 41 3.2.3. Distribution of GABAergic TRCs with respect to GABAB receptor subtype …………………………………………………………..….. 42 3.2.4. Co-localization patterns of the GABAB receptor R1 subunit with SNAP-25, Gα-gust as well as neuropeptides CCK and VIP ……….. 43 3.3. Physiological actions of GABA on TRCs in rat posterior lingual epithelium …………………………………………………………………. 45 3.3.1. Effects of GABA and the GABAA agonists on chloride currents recorded from TRCs ………………………………………………... 46 3.3.2. Effects of GABA and the GABAB agonist Baclofen on inwardly rectifying potassium currents recorded from TRCs …………..……. 48

4. Discussion ……………………………………………………………………… 72

4.1. Presence of GABAergic TRCs in the taste buds ……………………...... 73 4.1.1. GABA as a neurotransmitter in the taste buds ……………………... 73 4.1.2. Phenotypes of GABAergic TRCs …………………………………... 75 4.1.3. Co-transmission in GABAergic TRCs ……………………...……… 77 4.2. GABA receptors in the taste buds …………………………………………. 79 4.3. Physiological actions of GABA on TRCs …………………..………..…… 81 4.4. GABA’s role in peripheral gustatory signal processing ………………...… 82 4.5. Concluding remarks and future directions ...... ………...…… 84

List of references …………………………………………………………...…...... 87

Appendix ……………………………………………………………………………….. 98

LIST OF TABLES

Table Page

1.1. Examples of neurotransmitters/neuromodulators as well as their actions in TB.. 18

2.1. List of primary antibodies ……………………………………………………… 30

3.1. Cell counting results of single labeling immunocytochemistry ………………... 51

3.2. Cell counting results of GAD65/67 double labeling immunocytochemical experiments …………………………………………………………………..… 51

3.3. Cell counting results of GABAB receptor R1 subunit double labeling immunocytochemical experiments …………………………………………….. 52

3.4. Enhancing effects of GABA and the GABAA receptor agonists (muscimol and isoguvacine) on isolated chloride currents recorded from dissociated TRCs ….. 52

3.5. Enhancing effects of GABA and the GABAB receptor agonist baclofen on inwardly rectifying potassium currents recorded from dissociated TRCs ……... 53

3.6. Effects of the GABAB receptor antagonist CGP35348 and the G protein inhibitor GDP-βs on blocking baclofen’s enhancement of inwardly rectifying potassium currents recorded from dissociated TRCs ……………………………………… 53

4.1 Summary of protein co-localization in taste buds ……………………………… 84

LIST OF FIGURES

Figure Page

3.1. Localization of GABA-immunoreactivity in rat posterior TRCs …………..… 54

3.2. Localization of GABA synthetic enzyme GAD65/67-immunoreactivity in rat posterior TRCs ………………………………………………………………... 55

3.3. Non-overlapping of GABA synthetic enzyme GAD65/67-immunoreactivity with immunoreactivity of NCAM (neural cell adhesion molecule) or PGP 9.5 (protein gene product 9.5) in rat posterior TRCs ………………………………………. 56

3.4. Overlapping of GABA synthetic enzyme GAD65/67-immunoreactivity with immunoreactivity of Gα-gust (G protein gustducin α subunit) in rat posterior TRCs …………………………………………………………………………... 57

3.5. Overlapping of GABA synthetic enzyme GAD65/67-immunoreactivity with immunoreactivity of the presynaptic membrane-specific protein SNAP-25 in rat posterior TRCs ………………………………………………………………… 58

3.6. Overlapping of GABA synthetic enzyme GAD65/67-immunoreactivity with immunoreactivity of the neuropeptides CCK (cholecystokinin) or VIP (vasoactive intestinal peptide) in rat posterior TRCs ………………………………………. 59

3.7. Localization of the GABAA receptor α1 & α3 subunits in rat posterior TRCs ... 60

3.8. Localization of the GABAB receptor R1 subunit-immunoreactivity in rat posterior TRCs …………………………………………………………………………… 61

3.9. Non-overlapping of GABA synthetic enzyme GAD65/67-immunoreactivity with immunoreactivity of the GABAB receptor R1 subunit in rat posterior TRCs …. 62

3.10. Overlapping of GABAB receptor R1 subunit-immunoreactivity with immunoreactivity of the presynaptic membrane-specific protein SNAP-25 in rat posterior TRCs …………………………………………………………………. 63

3.11. Non-overlapping of GABAB receptor R1 subunit-immunoreactivity with immunoreactivity of Gα-gust and the neuropeptide VIP in rat posterior TRCs ..64

xii

3.12. Overlapping of GABAB receptor R1 subunit-immunoreactivity with immunoreactivity of the neuropeptide CCK in rat posterior TRCs …………..... 65

3.13. GABA’s enhancing effects on chloride currents recorded from TRCs ………... 66

3.14. Effects of the GABAA receptor agonists muscimol on chloride currents recorded from TRCs ...…….……………………………………………………………... 67

3.15. Effects of the GABAA receptor agonists isoguvacine on chloride currents recorded from TRCs …………………………………………………………… 68

3.16. Effects of GABA on KIR recorded from TRCs .…………..………………….… 69

3.17. Effects of the GABAB receptor agonist baclofen on KIR recorded from TRCs ... 70

3.18. The GABAB receptor antagonist CGP 35348 and the G protein blocker GDP-βs abolished effects of the GABAB receptor agonist baclofen on KIR recorded from TRCs ………………………………………………………………………..….. 71

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ABBREVIATIONS

5-HT Serotonin ACh Acetylcholine ATP Adenosine Triphosphate CB Carotid Body CCK Cholecystokinin CNS Central Nervous System ECF Extracellular Fluid Gα-gust Gustducin α Subunit GABA γ-aminobutyric acid GAD Glutamate Decarboxylase GAT GABA Transporter gNST Gustatory Portion of Nucleus of the Solitary Tract GPCR Guanine Nucleotide-Binding Protein-Coupled Receptor ICF Intracellular Fluid

IP3R3 Type III Inositol 1,4,5-triphosphate Receptor Type 3

KIR Inwardly Rectifying Potassium Channel M1 Muscarinic Acetylcholine Receptor Type I mGluR Metabotropic Glutamate Receptor NA Noradrenaline NCAM Neural Cell Adhesion Molecule NMDA N-methyl-D-asparate Receptor for Glutamate NPY Neuropeptide Y PBS Phosphate Buffered Saline PGP 9.5 Protein Gene Product 9.5 PLCβ2 Phospholipase C β2

xiv PVDF Polyvinylidene Fluoride RT Room Temperature SE Standard Error SNAP-25 Synaptosomal Associated Protein of 25 Kilodaltons SS Somatostatin TB Taste Bud TBS Tris Buffered Saline TRC Taste Receptor Cell TRP Transient Receptor Potential TSA Tyramide Signal Amplification VIP Vasoactive Intestinal Polypeptide Y1 Neuropeptide Y Receptor Type I

CHAPTER 1

INTRODUCTION

Gustatory perception is initiated by stimulation of taste receptor cells (TRCs) that

are closely packed in taste buds (TBs) located throughout the oral cavity. In response to

taste stimuli, TRCs are thought to fire action potentials and release neurotransmitter(s) to

excite sensory afferent nerve fibers, which, in turn, transmit signals to the central nervous system (25). Increasing evidence now suggests that signal processing occurs within the taste buds before sensory afferent nerve activation and multiple neurotransmitters and neuropeptides may participate in modulating TRC’s output signals (e.g. 33).

Understanding this modulatory event in peripheral gustatory coding is fundamental and essential in order to completely decipher how the brain percepts the sense of taste.

For animals, the sense of taste serves to detect and distinguish palatable nutritious

food from toxins therefore is vital for animals’ survival. The ultimate behavior (i.e., food

acceptance or avoidance) elicited by the gustatory system is surprisingly simple

compared to the remarkably wide range of chemicals (in terms of the nature and the

concentration) that animals may encounter (71). Considerable amount of signal

integration therefore must exist between food recognition and the evoked behavioral

1 outputs. The emerging concept is that modulation of gustatory signaling begins within the

peripheral end organs, the taste buds.

1.1. Intrinsic neuromodulation in taste buds

It has been widely acknowledged that most, if not all, neuronal circuits are subject to neuromodulation from a variety of sources (e.g. 40). In some cases, neuromodulatory substances are released from cells intrinsic to a circuit and can affect other cells and synapses in the same circuit. This is classified as intrinsic neuromodulation and has been particularly well-documented in the sensory system. Since intrinsic neuromodulation is an integral part of the circuit, it operates whenever the circuit is active and enables local neuronal activity to regulate the sensitivity of these circuits to their respective stimuli

(39). Among the established models of intrinsic neuromodulation, an excellent analogy to the taste bud is the mammalian carotid body (CB), a polymodal chemical sensor with the ability to detect and maintain the chemical composition of arterial blood (58). Similarities between the taste bud and the carotid body lie in several levels from anatomical structures to physiological functions, strongly supporting the presence of intrinsic neuromodulation in taste buds.

First, both chemosensory organs manifest discrete functional units consisting of

various types of cells that are tightly clustered together in an organized manner.

Chemoreceptive cells (i.e., TRCs in TB; type I cells in CB) are intimately associated with

each other as well as with interneuron-like cells (e.g. Merkel-like basal cells in amphibian

TB, ref. 23) or glial-like cells (type II cells in CB) by reciprocal chemical and electrical

2 synapses (2, 8, 23, 25, 58, 66, 68, 69, 92, 93). This arrangement is ideally suited for both

autocrine and paracrine regulation of chemoreceptor function.

Second, both TRCs in TB and type I cells in CB can be excited by a broad range

of chemical stimuli. Tastants are currently categorized into five modalities (sweet, bitter,

sour, salty, and umami). Even tastants within the same modality may greatly vary from

each other not only in terms of molecular structures but also in terms of detection

threshold. Several populations of TRCs in a TB are thought to be segregated by their sensitivities to different taste modalities (71). In the CB, type I cells are recognized as polymodal sensors endowed with the ability to detect broadly the chemical composition of arterial blood, including low PO2 (hypoxia), elevated PCO2 (hypercapnia), low pH

(acidity), osmolality and low glucose (58). On the other hand, functional results of chemoreceptor activation are relatively much less complex: control of feeding behavior by TRCs or control of arterial ventilation by type I cells. Extensive intrinsic neuromodulation necessary for shaping the final output signals has been uncovered in CB.

Undoubtedly, intrinsic neuromodulation is greatly desirable in TB as well.

Finally, a remarkable feature is shared by TB and CB: the presence of a wide

diversity of endogenous neuroactive substances within the circuit. These substances

include classical neurotransmitters (e.g., serotonin/5-HT, noradrenaline/NA and ATP)

and neuromodulators (e.g., neuropeptides, ref. 33, 58). These molecules are the material

substrates that mediate chemosensory signal transmission and modulation. This notion is

supported by numerous studies. In TB, ATP was recently identified to be a crucial factor

for neurotransmission between TRCs and gustatory nerves by acting on the P2X

purinergic receptors expressed on the afferent nerve terminals (24). Meanwhile, a series

3 of studies demonstrated that by activating the P2Y purinergic receptors expressed on

TRCs, ATP could also play a modulatory role in affecting the excitability of TRCs themselves (5, 13, 38). In CB, co-release of acetylcholine (ACh) and ATP was found to be the main mechanism mediating transmission of hypoxic (low PO2), acidic (low pH), and hypercapnic (high PCO2) signals. Both ACh and ATP, however, may also act as

neuromodulators of CB function. ACh could activate presynaptic autoreceptors while

ATP could exert modulatory influences on type I cells indirectly via affecting type II

cells within the same cell cluster (58). Overall, ample evidence clearly point to the

predominant role of chemical communication via neuroactive agents in modulating the

signal transduction process in peripheral chemosensory organs such as the taste bud and

the carotid body.

1.2. Communication between TRCs in taste bud

Recent studies exploring signal transduction mechanisms of TRCs also provided several lines of evidence necessitating occurrence of intrinsic neuromodulation within TB by way of cell-to-cell communication.

TRCs are differentiated epithelial cells with great heterogeneity. Anatomically, only a small portion of TRCs in a single bud are found to form unambiguous synapses with afferent nerve fibers. However, many more TRCs have access to the oral cavity and can respond to taste stimuli (33). At the molecular level, members of machineries essential for gustatory signal transduction, such as the T2R family of bitter receptors, the taste-specific G-protein gustducin, phospholipase C β2 (PLCβ2), type III inositol 1,4,5- trisphosphate receptor (IP3R3), and a member of the transient receptor potential (TRP)

4 channel family, TRPM5, are all localized in subsets of TRCs that lack synaptic connection with gustatory afferent nerve fibers (1, 65, 93, 94). Therefore, indirect mechanisms must exist for TRCs containing signaling molecules to excite the afferent nerve after stimulated by tastants. The most straightforward solution would be electrical coupling and/or neuroactive substances-mediated synaptic or paracrine communication between TRCs.

Even more persuasive evidence suggesting the existence of intrinsic neuromodulation by way of cell-to-cell communication within TB come from a wealth of studies showing an array of physiological actions of endogenous neuroactive agents on

TRCs (Table 1.1), which reveal multiple routes of chemical communication within the taste buds (33).

It is well known that TRCs not only express multiple neurotransmitters

(serotonin/5-HT, noradrenaline/NA, glutamate/Glu, adenosine triphosphate/ATP, acetylcholine/ACh and gamma-aminobutyric acid/GABA) and neuropeptides

(cholecystokinin/CCK, neuropeptide Y/NPY, vasoactive intestinal polypeptide/VIP and somatostatin/SS), but also express various subtypes of receptors for 5-HT, NA, Glu, ATP,

ACh, CCK, and NPY (3, 5, 7, 12-21, 28-35, 37, 38, 41, 47, 50, 52, 56, 59, 61, 72, 83, 96).

More importantly, different receptor subtypes were demonstrated to mediate physiological responses of TRCs (e.g. alteration of cell excitability) following exogenous application of corresponding agonists such as 5-HT (28, 31), NA (29, 32), ATP (5, 13,

38), ACh (61), CCK (34, 52, 72), and NPY (96), strongly suggesting functional roles of neurotransmitters and neuropeptides in affecting taste signaling within the taste bud during sapid stimulation.

5 1.2.1. Neurotransmitters in TB: 5-HT, NA, glutamate and ATP

Serotonin/5-HT is so far the most extensively studied neurotransmitter in taste

buds. Apart from serotonin’s putative role as the neurotransmitter between TRC and the

afferent nerve (e.g., 35), a series of observations has established a serotonergic paracrine

pathway among TRCs within the bud (28, 31). First, TRCs were found to respond to

serotonin under physiological conditions. Application of micromolar concentrations of

serotonin inhibited two types of currents involved in formation of action potentials in

TRCs: calcium-activated potassium currents and voltage-dependent sodium currents.

Therefore serotonin released during active stimulation may influence taste cell signal

processing by affecting their electrical properties. It was then shown that the inhibitory

effects of serotonin could be mimicked by 8OH-DPAT, a specific 5-HT1A receptor agonist, but not by phenylbiguanide, a specific 5-HT3 receptor agonist, indicating that 5-

HT1A receptor mediated TRC’s response. In a subsequent RT-PCR survey of fourteen serotonin receptor subtypes, the mRNA of 5-HT1A and 5-HT3 receptors was observed in

taste buds. Cellular localization of these two receptor subtypes was studied using

immunocytochemical technique with specific antibodies. Immunoreactivity of 5-HT1A receptor was observed in subsets of TRCs whereas 5-HT3 receptor-immunoreactivity was observed only in ganglion cells and nerve fibers in the dermal core of the papillae, confirming that 5-HT1A receptor is the major mediator of serotonin’s actions in TRCs.

Finally, double labeling experiments demonstrated that serotonin- and 5-HT1A receptor- immunoreactivity were present in exclusively non-overlapping TRC populations, indicating that serotonin released by one set of TRCs may affect neighboring 5-HT1A receptor-expressing cells via a paracrine route.

6 Another classic neurotransmitter, noradrenaline/NA, has been suggested as a

neurotransmitter in the peripheral gustatory system for several decades. Yet only recently

direct evidence was provided to demonstrate adrenergic signaling between TRCs (29, 32).

A putative paracrine role of NA is supported by studies showing that NA and its

receptors are expressed by subsets of TRCs and adrenergic receptors mediate NA or its

receptor agonists’ actions on TRCs under physiological conditions. Adrenergic actions on

TRCs are thought to occur during a state of excitation, given the facts that 1) NA was observed to inhibit outward potassium currents in TRCs, which would increase the duration of action potentials and cause the cell to remain in a depolarized state for a

prolonged time. This inhibitory effect could be mimicked by both α and β adrenergic

receptor agonists and abolished by receptor antagonist. 2) The α adrenergic receptor

agonists clonidine and phenylephrine both elevated intracellular calcium levels in TRCs,

an additional event correlated with cell excitation. Since there is considerable overlap in

the second messenger systems utilized by adrenergic transmission in non-gustatory

tissues and those identified in gustatory signal transduction pathways of TRCs, it is very

likely that NA may modulate peripheral gustatory signal transduction in TRCs by acting

as a paracrine mediator. This modulation would be based on the convergence of the

second messenger systems.

Glutamate has a variety of modulatory effects on a diverse range of cell types. In

the taste bud, glutamate may also play complicated roles since it is considered as both a

neurotransmitter and a taste stimulus (referred to as umami, ref. 7, 12, 14-21, 37, 47, 50,

56, 63, 83, 91, 98). Sources of glutamate may include food, taste bud cells, and primary

sensory neurons that innervate the taste buds. Several types of glutamate receptors have 7 been shown to be present in rodent TRCs, including ionotropic receptors, metabotropic receptors (mGluR), a novel truncated variant of mGluR4, and a heterodimer of T1R1 and

T1R3 of the T1R receptor family that may serve as an amino acid receptor. Besides transducing umami taste, these receptors may also mediate modulatory effects of glutamate on TRC activities. Physiological actions of glutamate on altering taste cell

membrane electrical properties have been reported to involve NMDA receptors and

metabotropic glutamate receptors (14, 50). Although some responses of TRCs to

glutamate seen in these studies are thought to represent taste transduction mechanism, it

can not be ruled out that glutamate may act as a neuromodulator in these cases. In

addition, it was recently identified that kainate GluRs are the main non-NMDA

ionotropic glutamate receptors present in the taste bud (20). Given their known

modulatory functions in the central nervous system, it is highly probable that kainate

GluRs may mediate similar effects inside the taste bud such as regulating

neurotransmitter release.

Adenosine triphosphate/ATP has been well recognized as a neurotransmitter and

neuromodulator in the nervous system in addition to its central role in cellular energy

metabolism. Yet not until recently were purinergic receptor isoforms identified in the

rodent taste bud, with the P2X receptors solely expressed in intragemmal sensory fibers

whereas P2Y receptors expressed exclusively in TRCs located throughout the tongue (5,

13, 24, 38). Modulation of TRC excitability by ATP is thought to mainly involve

activating G-protein coupled P2Y receptors to mobilize intracellular Ca2+, which then led

to opening of Ca2+-dependent Cl- channels (5, 13, 38).

8 1.2.2. Neuropeptides in TB: CCK and NPY

Neuropeptides has long been known to co-localize with classic small molecule neurotransmitters and act as neuromodulators. This appears to be the case in taste buds as well. Two brain-gut peptides, cholecystokinin/CCK and neuropeptide Y/NPY, have been demonstrated to be expressed by subsets of TRCs. Recently, CCK and NPY were shown to participate in communication within taste buds with striking features hence fulfilling their modulatory roles (34, 52, 72, 96). First, CCK and NPY elicit antagonistic physiological responses in rat TRCs. When applied exogenously, CCK was observed to exert three complimentary excitatory physiological actions on TRCs - inhibition of outward potassium currents (delayed-rectifier), inwardly rectifying potassium currents

(KIR), and elevation of intracellular calcium levels from intracellular stores. These actions would act in concert to place the cell into a more excitatory state. CCK-A receptor appeared to be responsible for mediating these effects. In contrast, NPY specifically enhanced KIR via NPY-1 receptors in rat TRCs. KIR is commonly known to contribute to

the resting membrane potential (81). Therefore enhancement of this current by NPY

would stabilize TRC’s resting potential, causing it to be less excitable, whereas inhibition

of KIR by CCK would produce the opposite effect. When released from TRCs (possibly following activation by gustatory stimulation), CCK and NPY may act simultaneously to excite and inhibit different sets of TRCs or could act sequentially to excite then inhibit the same sets of cells. To further elucidate how CCK and NPY may play roles in peripheral gustatory signal transduction, it is necessary to study the phenotype and localization patterns of peptide- and peptide receptor-expressing TRCs. So far little is known regarding characteristics of NPY- and NPY-1-expressing TRCs. On the other

9 hand, studies have shown that the majority of CCK-expressing TRCs are also bitter-

responsive cells that co-express bitter transducing G-protein gustducin, implying

involvement of CCK in bitter transduction. In contrary to the case of serotonin and 5-

HT1A receptor, which are expressed in virtually non-overlapping TRC populations, CCK

and CCK-A receptor immunoreactivity were observed in the same set of cells, suggesting

an autocrine pathway for CCK in taste buds (33). Another remarkable finding is the

colocalization patterns of these neuropeptides in taste buds. Nearly all TRCs that express

NPY were also found to co-express CCK, though a minority of CCK-expressing cells

may not express NPY (72). Taken together this large overlapping coexpression pattern

with the correlation of CCK-expression and bitter responsiveness, it is hypothesized that

CCK and NPY may act through competing pathways to participate signal processing

within taste buds during gustatory stimulation.

1.2.3. Other neuroactive substances in TB

Much less evidence has been reported regarding other neurotransmitters

(acetylcholine/ACh, and gamma-aminobutyric acid/GABA) and neuropeptides

(vasoactive intestinal polypeptide/VIP and somatostatin/SS) expressed in taste buds. ACh was recently observed to cause a large transient increase of intracellular calcium levels followed by a sustained phase of calcium increase in rat TRCs (52, 61). The transient increase of intracellular calcium is thought to be mediated by activating muscarinic ACh receptors that lead to mobilization of intracellular stores whereas the sustained phase of intracellular calcium increase is thought to involve calcium store-operated channels. The

M1 subtype of muscarinic receptors was reported to be present in both rat and mudpuppy

10 TRCs. As for the neuropeptides VIP and SS, although their presence in TRCs has been

studied (3, 30, 72), their functional roles in taste buds remain to be explored.

1.3. GABA as a candidate for mediating neuromodulation in taste bud

Research work presented in this dissertation concerns γ-aminobutyric acid/GABA, which is well-recognized as a neurotransmitter and a neuromodulator in the central nervous system (CNS, ref. 10, 62, 53, 57, 75, 76, 90) but has been poorly studied in taste buds.

1.3.1. GABA synthesis

GABA is the major inhibitory neurotransmitter in the mammalian nervous system

(90). In the CNS, GABA is synthesized from its precursor glutamate, an important excitatory neurotransmitter. The rate-limiting enzyme in GABA synthesis, glutamate decarboxylase(GAD), catalyzes the decarboxylation reaction in which glutamate is converted to GABA. In brain, GAD is found almost exclusively in the inhibitory neurons that release GABA as a neurotransmitter, therefore it is commonly used as a protein marker for GABAergic cells. Two isoforms of GAD have been identified in mammalian species: GAD65 and GAD67, where the numbers refer to their molecular weight in kilodaltons. Genetic studies indicate that GAD65 and GAD67 are products of two distinct genes (e.g. 54). Morphological studies reveal that even though many GABAergic neurons express both isoforms of GAD, there are differences in their subcellular distribution.

GAD67, which is responsible for synthesis of over 90% of GABA in the brain, is a soluble protein present in cytosol. On the other hand, GAD65 is preferentially localized near neuronal synaptic vesicles. Thus distinct roles are proposed for the two isoforms:

11 GAD67 is responsible for maintaining basal levels of GABA in the brain whereas

GAD65 contributes to GABA release from nerve terminal in response of environmental

signals. Although GABA has been detected in subsets of TRCs, presence of either

isoform of GAD in TB has not been reported in the literature.

1.3.2. GABA transporters

As commonly seen with other neurotransmitters, specific membrane transporters are responsible for GABA removal from the synaptic cleft (10, 70). GABA transporters belong to the same Na+/neurotransmitter symporter family of transporters for NA,

dopamine, and serotonin, with 60% to 70% identical in their amino acid sequences. In rat,

at least three GABA transporters (GAT1-3) have been identified by molecular cloning,

pharmacological and kinetic studies. Different distribution patterns of GAT subtypes

were observed: GAT1 and GAT3 showed neuronal tissue-specific expression with GAT1

present both in neuronal terminals and in glial processes while GAT3 localized only in

the glial processes. GAT2 was found to be widely distributed in various tissues. In rat

lingual epithelium, GAT3 was found only in taste bud cells of posterior tongue whereas

GAT2 was seen in nerve fibers in all areas of the tongue (59). No GAT1 expression was

detected in the tongue. Therefore GABAergic neurotransmission in the taste buds is

thought to be terminated by uptake of GABA through GAT3.

1.3.3. GABA receptors

In general, GABA is considered an inhibitory neurotransmitter in the mature

vertebrate brain since its binding with all three classes of GABA receptors so far

identified elicits inhibitory effects (10, 53, 57, 62, 75, 76, 90). GABAA and GABAC receptors belong to the ligand-gated chloride channel superfamily and are believed to

12 mediate the fast inhibitory activity of GABA. GABAB receptor is a member of the large family of G protein-coupled receptors (GPCRs) and regulates certain potassium and calcium channels that mediate the long-term inhibitory actions of GABA.

Being members of the same superfamily as nicotinic ACh receptor, glycine receptors, and serotonin receptors, GABAA and GABAC receptors share the same feature of being pentamers composed by various subunits (53, 62, 75). GABAA receptor subunits are categorized into seven classes: α1-6, β1-4, γ1-3, δ, ε, π, and θ. GABAC receptor is composed of ρ1-3 subunits. These subunits are encoded by unique genes, although some

genes are located on the same chromosome. Each subunit has similar structure but unique

expression pattern in the mammalian brain. The α1, β2, β3, and γ2 subunits are widely distributed and show the same distribution pattern. Among them, the α1 subunit is the most abundant one and is ubiquitously distributed throughout the brain. The α3 subunit is

expressed in regions where the α1 subunit is expressed at low levels. The ρ subunits are

expressed predominantly in the vertebrate retina. The π subunit is found outside the CNS

in organs such as uterus, prostate gland, thymus, and lung. Functional GABAA receptors contain at least one α, one β, and one γ subunit isoform. The most commonly seen

pentameric combinations are 2α2β1γ, 2α1β2γ, or 1α2β2γ. The δ or ε subunits are

thought to be assembled into GABAA receptors in place of γ subunit. The GABA binding sites are assumed to be contained in the N-terminal extracellular domains of both α and β

subunits. GABAA receptor also contains binding sites for several clinically important drugs, including benzodiazepines, barbiturates, and neurosteroids. GABAA and GABAC receptors differ dramatically in their kinetic and pharmacologic characteristics. Compared

13 to GABAA receptors, GABAC receptors have greater sensitivity to GABA, smaller

currents, and do not sensitize. GABAC receptors are insensitive to GABAA receptor antagonist bicucullin and the GABAB receptor agonist baclofen.

GABAB receptor shares structure similarities with metabotropic glutamate

receptors and vomeronasal receptors (pheromone and taste receptors). Two GABAB receptor subunits (R1 and R2) have been identified and are believed to be encoded by distinct genes. Heterodimer of R1 and R2 formed through an interaction between their intracellular C-terminal tails is thought to be necessary for formation of a functional

GABAB receptor. GABAA and GABAB receptors have distinct distribution patterns in the brain, though in many regions both receptors are present in comparable amounts.

Both GABAA and GABAB receptors can be found either pre- or postsynaptically.

In most cases postsynaptic GABAA receptor activation results in membrane hyperpolarization, and therefore decreases cell excitability. Presynaptic GABAA receptors have been seen in sensory afferent terminals, where increase of chloride conductance through GABAA receptor channel may reduce neurotransmitter release by shunting the presynaptic membrane. GABAB receptors predominantly couple to Gi/o type

G proteins. It is now well established that the βγ subunits are responsible for opening KIR channels postsynaptically and inhibiting voltage-gated calcium channels presynaptically

to inhibit neurotransmitter release whereas Gαi/o subunits mediate inhibition of adenylyl

cyclase. GABAB receptors may act synergistically with Gαs-coupled GPCRs such as β-

adrenergic receptors and mGluR1 due to G protein cross-talk.

14 1.3.4. GABA in the gustatory system

Being the major inhibitory neurotransmitter in the central nervous system,

GABA is well documented to play key roles in diverse neural functions including sensory signal processing. As in the visual and auditory systems, accumulating evidence indicates that GABA exerts its functions at each level of the gustatory system from the periphery to the cerebral cortex (e.g. 26, 42-44, 46, 48, 51, 59, 60, 73, 77-80, 82).

GABA has been postulated to participate in peripheral gustatory signaling in

TB for several decades. In an early electrophysiological study (82), GABA did not inhibit taste cell response elicited by NaCl; instead, it depressed the neural response of glossopharyngeal nerve in frog. Therefore GABA is thought to affect transmission of excitations from TRCs to afferent nerve endings. GABA expression has been shown to be restricted to nerve fibers that innervate the taste buds in Necturus (37). Only until recently cells in rat taste buds were identified as the endogenous source of GABA as well as the membrane transporters responsible for GABA-uptake (59), suggesting involvement of GABA neurotransmission in peripheral gustatory signal processing. However, details are lacking regarding the expression profile of GABA in mammalian taste buds.

Furthermore, other information necessary for firmly establishing GABA’s role as a neurotransmitter in taste buds remain elusive. This includes characteristics of distribution and functions of GABA receptors as well as vesicular storage and release of GABA in taste buds.

GABAergic projections and interneurons are abundant throughout the CNS, including an array of gustatory relay nuclei and the cortical taste area (e.g. 26, 42, 43, 48,

51, 60, 77-79). For example, both GABA-containing neurons and synaptic terminals are

15 richly present in the gustatory portion of rodent nucleus of the solitary tract (gNST). In

fact, the majority of taste-responsive neurons of the gNST are maintained under tonic

GABAergic inhibition, which provides a mechanism to increase and sharpen the breadth of responsiveness of taste-sensitive gNST neurons (77). In another study, in vivo

responses of gNST neurons to taste stimuli (sucrose, NaCl or KCl), while recorded

extracellularly, could be inhibited by local microinjection of GABA (79). Additionally,

GABA was demonstrated to mediate the differential inhibitory effects of corticofugal

input on gustatory afferent activity in gNST (78). Therefore, GABA plays important roles

in processing gustatory information in the gNST. Modulation of gustatory afferent

activities by GABA in gNST involves both GABAA and GABAB receptors activation (42,

48, 77-79).

Considering the wide distribution of GABA in gustatory system, it is not surprise

to see that GABA is able to influence different taste-guided behaviors such as

consumption and aversion (e.g. 9, 64, 73, 80). For example, in a study using mice

deficient in GAD65 (73), which is the key synthetic enzyme responsible for activity-

induced GABA production, consumption of sucrose-quinine mixture but not

preference/aversion responses to four basic taste stimuli was altered in the null mutant

mice when compared to the wild-type mice. This suggests that GAD65-generated GABA

may participate in complex processing of taste information rather than simple detection

and discrimination of tastants. In another study, mice lacking the α1 subunit of the

GABAA receptor showed reduction of saccharin and ethanol consumption, increased

ethanol-conditioned taste aversion and apparent ethanol-stimulated motor activity (9).

16 Other studies have shown that the GABAA receptor agonist benzodiazepine stimulate feeding in animals by enhancing the palatability of food (e.g. 64, 80).

Based on all of the above information, this dissertation proposes that GABA plays

a modulatory role in peripheral gustatory signal processing by mediating cell-to-cell

communication within taste buds. Evidence gathered for supporting this notion includes

the following aspects: 1) expression, cellular localization, and distribution patterns of

GABA, GABA synthetic enzyme GAD, GABA receptor subtypes GABAA and GABAB in taste buds; 2) phenotypes of GABAergic TRCs and GABA receptor-expressing TRCs;

3) actions of GABA and receptor agonists on TRCs under physiological conditions.

Neurotransmitter/ Physiological effects on TRC Receptors Neuromodulator involved

2+ + Serotonin/5-HT Inhibition of Ca -dependent K currents 5-HT1A (Ref. 28, 31) Inhibition of voltage-dependent Na+ currents

Noradrenaline/ Inhibition of outward K+ currents α & β NA Elevation of intracellular Ca2+ levels α (Ref. 29, 32) Enhancement of outward Cl- currents β

Glutamate Elevation of intracellular Ca2+ levels NMDA & (Ref. 50, 14) kinate Increase in holding current NMDA Decrease in holding current mGluR

Acetylcholine/ACh Elevation of intracellular Ca2+ levels M1 (Ref. 61)

Adenosine Elevation of intracellular Ca2+ levels P2Y triphosphate/ATP (Ref. 13-5)

Cholecystokinin/ Inhibition of outward K+ currents CCK-A CCK Inhibition of inwardly rectifying K+ currents (Ref. 34-72) Elevation of intracellular Ca2+ levels

Neuropeptide Y/ Enhancement of inwardly rectifying K+ currents Y1 NPY (Ref. 96)

Table 1.1. Examples of neurotransmitters/neuromodulators as well as their actions in TB

CHAPTER 2

METHODS

2.1. Immunocytochemistry

2.1.1. Tissue preparation

Circumvallate or foliate papillae were quickly dissected and fixed by immersion

in 4% paraformaldehyde or Bouin’s fixative (71% saturated picric acid, 24% glacial acetic acid, and 5% formalin) for 5 hours (h) at 4°C. For frozen sections, tissue blocks were cryoprotected in 30% sucrose and frozen in methylbutane and dry ice before sectioning at 8 µm thickness on cryostat. Alternatively, tissue blocks were dehydrated, embedded in paraffin, and sectioned on a rotary microtome at 8 µm thickness. All

sections were collected onto Fisher Superfrost Plus slides.

2.1.2. Conventional immunocytochemistry protocol

Paraffin sections were deparaffinized and rehydrated before subsequent processing. Frozen sections were air-dried for 30 min at room temperature (RT) before the following procedures. In all experiments, sections were rinsed in 0.01 M phosphate buffered saline (PBS, pH 7.4). For horseradish peroxidase (HRP)-involved reactions,

sections were first blocked with 0.5% H2O2 in methanol for 30 min to eliminate

endogenous peroxidase activity. Subsequent blocking with 5% dry milk, 0.75% gelatin,

19 and 10% normal goat serum (all dissolved in PBS) for 30 to 60 min at RT were

performed to reduce nonspecific antibody binding. Sections were then incubated in

primary antiserum at the appropriate dilutions (Table 2.1). Slides were housed in a closed moist chamber for 36 h at 4°C.

Subsequently, the sections were rinsed in PBS (3 x 10 min) before incubated for 1 h at RT in secondary biotinylated goat-anti-rabbit IgG (1:400), rinsed again in PBS, and then incubated for 1 h in avidin-biotin-peroxidase complex (Vectastain "Elite ABC" kit,

Vector Laboratories, Burlingame, CA) at a dilution of 1:50. Tissue-bound peroxidase was

visualized by incubating sections for 3 min in a freshly prepared solution containing

0.01% hydrogen peroxide and 0.05% 3-3'-diaminobenzidine tetrahydrochloride (DAB,

Sigma) in 0.05 M Tris buffer, pH 7.6. Subsequently, sections were rinsed in deionized

water, dehydrated through a series of ethanol, cleared in xylene, and coverslipped with

Permont (Fisher Scientific, Fair Lawn, NJ).

A similar protocol was employed for immunocytochemical examination using epifluroescence. The peroxidase blocking step was omitted. After incubation in the primary antibody, sections were rinsed in PBS and then detected using Cy3-conjugated

goat anti-rabbit IgG serum (1:400, RT, 1 h, in dark). Slides were mounted in Cytoseal 60

(Electron Microscopy Sciences, Washington, PA) or Gel Mount (Sigma).

Immunofluorescence was observed under a Nikon microscope and digital photos were

processed using Photoshop software.

2.1.3. TSA-amplified immunocytochemistry protocol

Paraffin or frozen sections were first incubated with a solution of 0.5% hydrogen

peroxide in methanol for 30 min to eliminate endogenous peroxidase activity. To reduce

20 nonspecific antibody binding, sections were then incubated for 1 h at RT in PBS containing 10% normal goat serum and 0.3% Triton X-100. Primary antiserum (Table 2.1)

was subsequently applied to the sections and the slides were housed in a closed moist chamber for 36 h at 4°C. After PBS washing (3 x 10 min), sections were incubated with biotin-streptavidin-conjugated goat anti-rabbit (or donkey anti-guinea pig) IgG Fab

fragment (1:800, Jackson ImmunoResearch Laboratories, West Grove, PA) at RT for 1 h.

In the following steps, sections were rinsed in TNT buffer (0.1 M Tris, 0.15 M NaCl,

0.05% Tween-20, pH 7.5). After incubated for 30 min at RT with TNB buffer (TNT with

0.5% blocking reagent provided in the TSA kit; Indirect NEL 700A, NEN Life Science

Products, Boston, MA), sections were incubated with HRP-conjugated streptavidin

(diluted 1:100 in TNB buffer, RT, 30 min, in dark, provided in the TSA kit). Sections

were rinsed in TNT and then incubated with biotinyl tyramide (1:50 in amplification diluent, provided in the TSA kit) for 10 min (RT, in dark). After being washed in PBS, immunoreactivity was visualized with streptavidin-fluorescein (1:400, Jackson

ImmunoResearch Labs). Slides were mounted in Cytoseal 60 or Gel Mount.

Omission of primary antibody was performed as negative control and no positive staining was observed in either conventional or TSA amplified immunocytochemistry experiments.

2.1.4. Double-labeling immunocytochemistry protocol

An indirect immunofluorescence double-labeling protocol was modified to allow localization of two antigens in the same preparation when both primary antibodies are raised in the same species. This protocol relies on a combination of the methods in previously published papers and involves using TSA with a Fab fragment secondary

21 antibody for detection of the first primary antibody (6, 36, 74, 89). With the use of TSA,

the first primary antibody can be used at very low concentration so that the antigen can

only be detected by TSA but not by a conventional fluorophore-conjugated secondary

antibody, which prevents the cross-reaction between the first primary antibody and the

second secondary antibody (referred to as interference I), while the use of a Fab fragment

instead of the whole IgG molecule or F(ab)2 fragment as the first secondary antibody prevents the capture of the second primary antibody by the first secondary antibody

(interference II). Therefore, this modified protocol prevents cross-reactions between the primary and the unintended secondary antibodies.

Two control experiments were performed to ensure this cross reactivity did not

occur. To control for interference I (the first primary with the second secondary), after incubation with the first primary, sections are reacted using the second secondary antibody and the standard (non-TSA) protocol (Cy3-conjugated goat anti-rabbit IgG;

1:800, RT, 1 h). No fluorescence was observed, indicating that the first primary was too dilute to be detected with unamplified means. To control for interference II (the first secondary with the second primary), a substitute second primary (also from rabbit) whose antigen is not expressed in lingual tissue was employed. Rabbit anti-Iba (ionized calcium

binding adaptor molecule-1), which is expressed in microglia and macrophages but not

lingual epithelium, was chosen. If there were interference binding, the second primary would be visualized. This was not evident in control experiments. Parallel experiments were performed in inverted order (i.e., switching primary 1 and primary 2) and produced equivalent results.

22 Experiments were done as the following protocol:

Fixed frozen 8-µm sections containing taste buds were rinsed in 0.01 M PBS (pH

7.4). Sections were first incubated with a solution of 0.5% hydrogen peroxide in methanol for 30 min to eliminate endogenous peroxidase activity. To reduce nonspecific antibody binding, the sections then were incubated for 1 h at RT in blocking solution (PBS containing 10% normal goat serum and 0.3% Triton X-100). The first primary antiserum was applied at a dilution that would not allow the antigen to be detected by Cy3- conjugated goat anti-rabbit IgG serum (1:800, RT, 1 h) in the conventional immunofluorescence method but was still detectable after TSA. The slides were housed in a closed moist chamber for 36 h at 4°C.

Subsequently, the sections were rinsed in PBS (3 x 10 min) and then incubated with biotin-streptavidin-conjugated goat anti rabbit IgG Fab fragment (1:1,000, RT, 1 h).

After being rinsed in TNT (3 x 5 min), the sections were incubated for 30 min at room temperature with TNB buffer. The sections were then incubated with HRP-conjugated streptavidin (1:500 in TNB buffer, RT, 30 min, in dark). The sections were rinsed in TNT and then incubated with biotinyl tyramide (1:50 in amplification diluent) for 10 min (RT, in dark). After being washed in TNT, streptavidin-fluorescein (1:800) was applied for 1 h at room temperature in the dark. Following PBS rinsing, the sections were incubated for

36 h at 4°C in the dark with the second primary antibody later detected by Cy3- conjugated goat anti-rabbit IgG serum (1:800, RT, 1 h, in dark). Slides were mounted in

Cytoseal 60 or Gel Mount. To control for the ability of Cy3-conjugated secondary antibodies to detect the first primary antiserum, after the TSA and streptavidin steps,

Cy3-conjugated secondary antibodies diluted 1:800 in PBS were applied for 1 h at room

23 temperature. It was observed that with omission of the second primary antibody, no signal showing red fluorescence could be detected.

For double labeling experiments using primary antibodies raised in different species, the following protocol was performed: after overnight incubation of the first primary antibody (rabbit anti-rat antigen, Table 2.1) at 4°C, the sections were rinsed in

PBS (3 x 5 min) and then incubated with biotin-streptavidin-conjugated goat anti rabbit

(or donkey anti-guinea pig) IgG Fab fragment (1:400, 4°C, overnight). The following day, sections were washed then kept in dark for incubation with streptavidin-fluorescein

(1:400) as well as the second primary antibody (mouse anti-SNAP-25 1:200, Chemicon

International MAB331, Temecula, CA) for 2 h at RT. After subsequent rinses, sections were incubated in dark with Cy3-conjugated donkey anti-mouse IgG (1:200, Jackson

ImmunoResearch Laboratories) for 1 h at RT before final washes and coverslipping with

Gel Mount.

2.1.5. Cell counting and data analysis

Immunocytochemical data were analyzed quantitatively by cell counting. Sections were chosen from at least 3 independent experiments repeated identically using different animals. To ensure that a taste bud was not counted twice, one out of every 12 to 40 consecutive 8 µm sections (i.e., sections spaced by greater than 80 µm) were selected and adjacent sections were never chosen for analysis. Using either single- or double-labeling protocols, positively stained TRCs were typically observed in a great majority of the taste buds identified on each section. Individual buds were selected for analysis and only those labeled cells that displayed apparent apical processes and/or perinuclear region were counted.

24 To ensure the consistency between the conventional and the amplification

protocols, dilution series of the primary antibodies were performed. The most appropriate

dilutions for each antibody were chosen (Table 2.1) so that when comparing the numbers

of labeled cells (per cross section of a taste bud) obtained from either protocol using the

same primary antibody, no significant difference was observed between the two.

Cell counting results of the experiments using both foliate and circumvallate

papillae were pooled together since there was no significant difference observed between

data gathered from the two areas in any of the independent experiments.

2.2. Western Blotting

Excised tissue blocks (including both gustatory and non-gustatory epithelium as well as a thin muscle layer underlying the epithelium) were immediately homogenized

using IKA Ultra-Turrex T8 homogenizer (Janke & Kunkel GmbH, Germany) in

homogenization buffer (15 µl/mg wet weight; buffer consists of 50 mM β- glycerophosphate, 1.5 mM EGTA, 0.1 mM Na3VO4, 1.0 mM dithiothreitol, 10 µg/ml aprotinin, 1 µg/ml pepstatin, 10 µg/ml leupeptin, 1 µM phenylmethanesulfonyl fluoride), diluted 50:50 with 2X sample buffer (0.1 M TRIS, pH 6.8, 10% 2-mercaptoethanol, 20% glycerol, 2% sodium dodecyl sulfate (SDS)), heated (95oC, 3 min), and centrifuged

(14,000 rpm, 10 min, 4oC). Supernatants were then transferred to clean tubes and stored at -20oC.

In membrane preparation, tissue blocks were homogenized on ice. Homogenates

were first centrifuged at 1960g at 4oC for 10 min. Supernatants were transferred to clean

tubes then centrifuged again at 20800g at 4oC for 1 hr. Supernatants (containing 25 cytoplasmic proteins) were transferred to clean tubes and stored at -20oC. Pellets

(containing membranous proteins) were resuspended in homogenization buffer and stored

at -20oC.

Protein concentration was determined using the Bio-Rad (Hercules, CA) RC-DC

protein assay kit. Equivalent amounts of total protein were loaded onto 12% gels for

electrophoresis. Proteins were transferred to 0.22 µm polyvinylidene fluoride (PVDF) membranes (100V, 45 min). Membranes were washed with TRIS-buffered saline containing 0.1% Tween-20 (TBST), blocked with 5% non-fat dry milk in TBST (1 h, room temperature) and incubated in primary antibody (Table 2.1) overnight at 4oC. Blots were visualized using a chemiluminecent detection kit (SuperSignal West Femto

Maximum Sensitivity Substrate, Pierce Biotechnology, Rockford, IL) and exposed to X- ray films (Hyperfilm MP, Amersham Biosciences, Piscataway, NJ).

2.3. Patch clamp recording

2.3.1. Dissociation of rat taste receptor cells

Patch-clamp recordings were performed on taste receptor cells isolated from the

circumvallate and foliate papillae of adult male Sprague-Dawley rats. All procedures

were approved by the Ohio State University's Laboratory Animal Care and Use

Committee and adhered to the National Institutes of Health Guide for the Care and Use of

Laboratory Animals. Animals were brought to a surgical level of anesthesia by intraperitoneal injection of a 0.09 ml/100 g body wt ketamine (91 mg/ml; Fort Dodge

Laboratories)-acepromazine (0.09 mg/ml; Butler Laboratories) mixture before decapitation and excision of foliate and circumvallate papillae. The papillae were then

26 o incubated for several hours at 32 C in 5% CO2/95% air with cysteine-activated (1 mg/ml) papain (14 U/ml) in divalent-free solution composed of (mM): 80 NaCl, 5 KCl, 26

NaHCO3, 2.5 NaH2PO4•1H2O, 20 D-glucose and 1 EDTA. Cells were dissociated by

gentle agitation in the standard extracellular fluid (ECF) comprised (mM): 126 NaCl,

1.25 NaH2PO4•1H2O, 5 KCl, 5 NaHepes, 2 MgCl2, 2 CaCl2 and 10 glucose, to pH 7.4 with NaOH. Some papillae were maintained in ECF solution at 4 oC for later dissociation.

2.3.2. Solutions for patch clamp recordings

Both whole cell and perforated patch clamp recording were performed. In whole cell configuration, the pseudo-intracellular fluid (ICF) used for filling the recording pipette comprised (mM): 140 KCl, 2 MgCl2, 1 CaCl2, 11 EGTA, 10 Hepes and 4 ATP

(disodium salt). Perforated patch clamp recording was performed using amphotericin B

(Sigma, St Louis, MO) as the ionophore (400 µg/ml in the ICF). The composition of the

perforated patch ICF was (in mM): 55 KCl, 75 K2SO4, 8 MgCl2, and 10 HEPES.

To record isolated chloride currents, a potassium-free ECF was employed for bath

solution in both whole cell and perforated patch clamp recording modes. This ECF

consisted of (in mM) 126 NaCl, 5 HEPES, 2 CaCl2, 2 MgCl2, and 10 glucose with a final chloride concentration of 134 mM. In whole cell mode, ICF consisted of (in mM)

24 CsCl, 116 methanesulfonic acid (cesium salt), 2 MgCl2, 1 CaCl2, 11 EGTA,

10 HEPES, and 4 ATP (magnesium salt); this solution was adjusted to a final pH of

7.2 with CsOH and yielded a final chloride concentration of 30 mM. In perforated patch mode, ICF consisted of (in mM) 55 CsCl, 75 Cs2SO4, 8 MgCl2, and 10 HEPES and contained a final chloride concentration of 71 mM.

27 To record inwardly rectifying potassium currents, final extracellular potassium

concentration in the standard ECF recipe was elevated to 30 mM with the replacement of

126 mM NaCl and 5 mM KCl by 101 mM NaCl and 30 mM KCl. ICF for recording the inwardly rectifying potassium currents consisted of 10 mM KCl and 130 mM potassium salt of gluconic acid, instead of 140 mM KCl in the standard ICF.

2.3.3. Patch clamp recording procedures

The micropipettes used for recording were pulled from 1.5 mm o.d. borosilicate glass (World Precision Instruments, Sarasota, FL, USA) with typical resistances of 4-7

MΩ when filled with ICF and measured in ECF. The pipette tip was positioned to contact the cell membrane, and negative pressure was applied to its interior to facilitate gigaseal

formation. Junction potentials were corrected before the electrode contacted the cell. Seal resistances were typically on the order of several decades of gigaohms. Additional negative pressure was applied to enter whole-cell recording mode. For perforated patch recordings, approximately 30 minutes (min) were required to reach a stable level of recording after gigaseal formation. Fast and slow capacitance compensation was used as necessary with amplifier controls. Cell membrane capacitance and uncompensated series

resistance were adjusted to produce optimal transient balancing. Membrane capacitance

was 3-6 pF; series resistance averaged 10 MΩ in conventional whole-cell mode and 20-

50 MΩ in most amphotericin B-perforated patch-clamp recordings. Low-pass filtering

caused by resistance-capacitance coupling was considered minimal. The product of these factors produces a time constant of 30-300 µsec or a cutoff frequency (1/2πRC) of 1.6-

16.6 kHz.

28 Data were acquired with high-impedance amplifiers (Axopatch 200-A or B; Axon

Instruments, Union City, CA), Pentium-based 300 or 450 MHz computers, a 12 bit 330

kHz A/D converter (Digidata 1200; Axon Instruments) and a commercial software

program (pCLAMP, versions 7.0 or 8.01; Axon Instruments). Membrane currents were

acquired after low-pass filtering with a cut-off frequency of 5 or10 kHz (at -3 dB). A

software-driven D/A converter generated the voltage protocols. Recordings were made at

room temperature.

In most situations, membrane potential was held at -50 mV under voltage clamp

mode. Ramp protocols were used with command voltage varied linearly from -100 to 100

mV for recording voltage-dependent potassium currents or from -140 to 10 mV for

recording KIR. To record chloride currents, a holding potential of 0 mV was used with a ramp protocol in which command potential continuously changed from -130 mV to 120 mV.

Exchange of the bathing solution was accomplished with a gravity-fed perfusion system. All drugs were focally applied through a pipette positioned ~500 µm from the recorded cell. GABA was obtained from Calbiochem (La Jolla, CA) and Sigma. All other reagents were purchased from Sigma. Stock solutions were made with deionized water and diluted in ECF before application.

Data were analyzed with a combination of off-line software programs that included a software acquisition suite (pCLAMP, Axon Instruments), Microsoft Excel, and a technical graphics/analysis program (Origin 6.1; MicroCal Software). Data were normalized to the value of the current magnitude before drug application and are presented as mean ± SE.

Primary Antibody Dilution Experiment Rabbit anti-GABA (Sigma A2052) 1:1000 Single labeling ICC, conventional method Rabbit anti-GAD65/67 (Chemicon 1:200 Single and double labeling ICC, AB1511, Temecula, CA) 1:500 conventional method 1:1000 Western blotting

Rabbit anti-GABAA α1 subunit 1:50 Single and double labeling ICC, (Chemicon AB5592) TSA amplification method 1:200 Western blotting

Rabbit anti-GABAA α3 subunit 1:50 Single and double labeling ICC, (Chemicon AB5594) TSA amplification method 1:200 Western blotting

Guinea pig anti-GABAB R1 subunit 1:1000 Single and double labeling ICC, (Chemicon AB1531) 1:2000 TSA amplification method Rabbit anti-NCAM (Chemicon 1:1000 Double labeling ICC, conventional AB5032) 1:20,000 and TSA amplification methods Rabbit anti-gustducin α subunit (Santa 1:1000 Double labeling ICC, conventional Cruz Biotechnology sc-395, Santa 1:20,000 and TSA amplification methods Cruz, CA) Mouse anti-SNAP-25 (Chemicon 1:200 Double labeling ICC, conventional MAB331) method Rabbit anti-PGP9.5 (Biogenesis, 1:1000 Double labeling ICC, conventional Kingston, NH) 1:20,000 and TSA amplification methods Rabbit anti-cholecystokinin (CCK) 1:500 Double labeling ICC, conventional octapeptide (Chemicon AB1973) 1:8000 and TSA amplification methods Rabbit anti-vasoactive intestinal 1:1000 Double labeling ICC, conventional peptide (VIP, Diasorin #20077, 1:40,000 and TSA amplification methods Stillwater, MN)

Table 2.1. List of primary antibodies

CHAPTER 3

RESULTS

Morphological and functional studies have suggested that “cross-talk” occurs

among TRCs in the peripheral taste organ, the taste bud. Communications among TRCs

largely rely on chemical messengers (neurotransmitters and neuropeptides) present in the taste bud. By interacting with specific receptors expressed by TRCs, endogenous neuroactive molecules in the taste bud may contribute to processing of the initial sensory signals and help modulate the final output of the chemosensory TRCs to afferent nerves.

To understand how and to what extend this intrinsic neuromodulation actually occurs, it is necessary to determine the identity of the neuroactive agents in taste buds and examine the mechanisms by which their receptors activation may modify the physiological

functions of TRCs. Data presented in this chapter are results from a series of experiments

that were designed to investigate the distribution patterns and physiological actions of

one of the neurotransmitters in taste bud, GABA.

31 3.1. Distribution of GABAergic TRCs in rat posterior lingual epithelium

3.1.1. Expression of GABA and the GABA synthetic enzyme GAD65/67 in TRC

GABA has been postulated as one of the neurotransmitters involved in peripheral taste signal processing. Presence of GABA as well as the membrane transporter for

GABA-uptake, GAT3, has been demonstrated in subsets of taste bud cells at rat circumvallate papillae (59). In the present study, expression of GABA in rat posterior lingual epithelium was confirmed using an immunocytochemical technique with an antibody directed against GABA (Fig. 3.1). GABA-immunoreactivity was observed in populations of TRCs located in both the foliate and the circumvallate papillae, indicating that these cells were the endogenous source of GABA in the taste buds. Positively stained

TRCs typically displayed a smooth outline of an elongate, spindle shape. A large round or oval nucleus was commonly recognized situating at the middle level of the cell while tapered processes on both ends could generally span the whole length of the taste bud from the apical taste pore to the basement membrane. Immunoreactivity of GABA typically dispersed evenly throughout the cytoplasmic area of the cell whereas the nuclear region was devoid of staining. Correspondingly, on a transverse section, GABA- immunopositive cells appeared as circular or oval rings of immunofluoresence. On a cross-section of either foliate or circumvallate papillae at 8 µm thickness, over eighty percent of the identified taste buds contained positively stained TRCs; an average of 1.56

± 0.04 (mean ± standard error/SE) GABA-immunoreactive TRCs could be recognized in a single taste bud (Table 3.1). GABA-immunoreactive TRCs showed scattering distribution within the taste buds with a majority of them situated in the central area of

the bud. No immunoreactivity was observed in nerve fibers inside or around the taste

32 buds. No specifically labeled cell was observed in the surrounding epithelium outside the taste buds. No difference in any of the above morphological features of GABA- immunoreactive TRCs was found between the foliate and the circumvallate papillae.

To further confirm the presence of GABAergic TRCs in taste buds of rat posterior

lingual epithelium, expression of the key synthetic enzyme for GABA, GAD65/67, was

examined using western blotting and immunocytochemisty. In western blotting

experiments, immunoreactive bands to GAD65/67 were recognized in total protein

extracts from taste papillae of the anterior as well as the posterior regions of the tongue

(Fig. 3.2a). This result not only revealed the predicted existence of GAD65/67 in taste

papillae of the tongue but also verified the specificity of the antibody used in the present

study. Cellular distribution of GAD65/67 was examined in cross sections of rat foliate and circumvallate papillae (Fig. 3.2b-g). GAD65/67-immunopositive TRCs exhibited morphological characteristics in complete agreement with those of the GABA- immunoreactive TRCs. While comparing the cell counting results of GABA versus

GAD65/67 immunocytochmical experiments, only slight differences between the two were noticed regarding the percentage of taste buds containing positively stained cells as well as the number of immunoreactive cells in an 8 µm cross section of an individual taste bud (Table 3.1), which may reflect different inherent properties of the two antibodies utilized.

3.1.2. Phenotypes of GABAergic TRCs with respects to TRC population-specific protein markers

According to their morphological features (e.g., spindle shape, round nucleus, and an apical process extending to the taste pore), GABA-containing cells in rat taste buds

33 resemble TRCs expressing molecules of gustatory signaling cascades. However, little is

known regarding the exact phenotypes of GABAergic TRCs. Double labeling immunocytochemical studies were carried out to investigate co-localization patterns of

GAD65/67 with TRC population-specific protein markers.

Several protein markers, including the taste-specific G-protein gustducin α

subunit (Gα-gust), neural cell adhesion molecule (NCAM) and protein gene product 9.5

(PGP 9.5), have been widely employed to characterize distinct populations of TRCs (e.g.

25). Gα-gust has been implicated in transduction of both sweet and bitter-tasting substances (e.g. 71). Gα-gust-immunoreactivity could be detected in subsets of TRCs in

all taste bud regions of the rodent tongue (e.g. 11, 94). TRCs containing Gα-gust were spindle-shaped, with circular cross-sections and apical processes that extended to the taste pore, the latter being crucial for the chemoreceptive functions of these cells.

NCAM is expressed in about 20% of the cells within an adult rat circumvallate

taste bud (e.g. 67). This group of cells is separate from the population of Gα-gust-

containing TRCs. NCAM-expressing TRCs have been suggested to represent a distinct

group that synapses directly with gustatory afferent nerve fibers in the taste buds (e.g. 95).

Morphological characteristics of NCAM-immunoreactive TRCs include a slender elongate shape and a relatively narrow nuclear region. Most NCAM-containing TRCs also express the neurotransmitter 5-HT (95).

PGP 9.5, an ubiquitin carboxyl-terminal hydrolase, has been broadly recognized as a protein marker for sensory paraneurons. PGP 9.5-immunoreactivity is present in a subset of TRCs that demonstrates NCAM-immunoreactivity and forms synapses with afferent nerves but does not display 5-HT-immunoreactivity. A separate group of TRCs

34 that does not make synaptic contacts with intragemmal gustatory nerve endings exhibits

PGP 9.5-immunoreactivity but does not express Gα-gust. Overall, TRCs expressing the protein markers Gα-gust, NCAM or PGP 9.5 represent distinctive cell populations in taste buds yet these cells all share morphological similarities to the GABAergic TRCs described in session 3.1.1.

As demonstrated in Figure 3.3, there was virtually no overlapping of GAD65/67- immunoreactivity with either NCAM- or PGP 9.5-immunoreactivity in TRCs on all cross sections of rat foliate and circumvallate papillae examined. In contrast, over half of the

GAD65/67-immunopositive TRCs also displayed Gα-gust-immunoreactivity (Table 3.2 and Figure 3.4). This result implies that GABAergic TRCs are unlikely to be the chemosensory cells that make direct synaptic contacts with gustatory afferent nerve fibers inside the taste bud according to their non-overlapping distribution pattern with NCAM- immunopositive TRCs. Moreover, GABA and 5-HT are not likely to be co-expressed by the same cell in taste buds since 5-HT always co-localizes with NCAM. On the other hand, co-localization of GAD65/67- and Gα-gust-immunoreactivity in TRCs that was observed in the present study suggests that some of the GABAergic TRCs are capable of transducing taste signals such as sweet or bitter stimuli. It has been shown that all Gα- gust-containing cells also contain the amiloride-sensitive sodium channel (ASNaC) that is implicated in the detection of sodium salts by rats (49). Therefore GABAergic TRCs may be involved in transduction of signals belonging to multiple taste modalities.

35 3.1.3. Co-localization of GAD65/67 with the presynaptic membrane-specific protein

marker SNAP-25

Chemical synapses existing within the taste buds may be formed between TRCs

and afferent nerve fibers as well as between individual TRCs (e.g. 33). Although

GABAergic TRCs are mostly unlikely to form synapses directly with afferent gustatory nerve endings in taste buds, it is highly probable that they make synaptic contacts with adjacent TRCs in the same bud and/or they signal neighboring cells via regulated vesicular release of paracrine substances.

Typical synaptic neurotransmission in the nervous system involves regulated vesicular release of neuroactive substances. Several proteins are believed to play critical roles during regulated synaptic vesicle docking, membrane fusion and exocytosis of neurotransmitters. Among these proteins are vesicle-associated membrane protein synaptobrevin (VAMP) as well as the presynaptic membrane-specific proteins SNAP-25

(synaptosomal associated protein of 25 kD) and syntaxin. Existence of these three proteins has been demonstrated in rat taste buds (66, 92, 93).

Synaptobrevin-immunoreactivity was detected in approximately 35% of the cells of rat circumvallate taste buds (93). By using colloidal gold immunoelectron microscopy, it was found that synaptobrevin-immunoreactivity was associated with synaptic vesicles at the synapses of rat TRCs. All of the synapses observed from TRCs onto nerve processes displayed synaptobrevin-immunoreactivity. Some of the synaptobrevin- immunopositive TRCs also express SNAP-25, 5-HT, PGP 9.5 and IP3R3 (not necessarily

in the same cell). Notably, IP3R3 was found to co-localize almost completely with

PLCβ2 and mostly with Gα-gust in TRCs. TRCs co-expressing these signal transduction

36 molecules are capable of responding to various taste stimuli but are not believed to

synapse directly onto sensory nerves. Correspondingly, some TRCs without synaptic

contacts with afferent nerves were observed to be immunoreactive to synaptobrevin.

These cells may release autocrine or paracrine factors to influence neighboring cells

through regulated exocytosis in response to taste stimuli.

Similarly, the presynaptic membrane-specific protein SNAP-25 has been

demonstrated to be associated with nearly all morphologically identified synapses between TRCs and nerve processes in rat circumvallate papillae therefore is designated as a marker for TRCs with synapses (92). SNAP-25-immunoreactivity is present in a subset of elongate TRCs and most nerve processes inside and surrounding the taste buds. Some of the SNAP-25-immunoreactive TRCs also contain NCAM- and 5-HT- immunoreactivity. They are thought to represent the group of chemosensory TRCs that form synaptic contracts onto gustatory afferent nerves. Additionally, a partial overlapping pattern was observed between immunoreactivity of Gα-gust and the synaptic proteins synaptobrevin or SNAP-25 in TRCs of rat circumvallate papillae (66), implying that regulated vesicular release of neuroactive agents may be employed by TRCs with taste signal transduction functions but without synaptic contacts with nerve processes.

Endogenous neuroactive agents released in this process are likely to be the principle mediators for communication between the two groups of TRCs with and without direct synaptic contacts with afferent nerves, allowing TRCs stimulated by tastants to indirectly excite the afferent nerves.

As demonstrated in Table 3.2 and Figure 3.5, over two-thirds of the GAD65/67- immunoreactive TRCs were also labeled with an antibody directed against SNAP-25,

37 suggesting that most GABAergic TRCs are able to utilize regulated exocytosis as a

mechanism for releasing endogenous neuroactive substances. On the other hand, merely

forty percent of the SNAP-25-immunopositive TRCs were double-labeled with

GAD65/67 antibody, implying that only a minority of TRCs with synapses may utilize

GABA as a neurotransmitter. This evident heterogeneity of the neurochemical content of

synapses-containing TRCs is consistent with numerous reports revealing the greatly

diversified biochemical composition of these cells (e.g. 25).

3.1.4. Co-localization of GAD65/67 with neuropeptides CCK and VIP

It has been widely appreciated that many, if not all, GABAergic cells in the

nervous system contain other neurotransmitters or neuropeptides. These neuroactive

substances may act at autoreceptors and modulate GABA release, they may act on axons

of adjacent neurons to modulate their transmitter release, or they may modulate

postsynaptic responses to GABA or other transmitters (84). The four neuropeptides

identified in taste bud cells, CCK, VIP, NPY and somatostatin, have been shown

individually to be co-expressed with GABA in neurons throughout the brain. For

example, GABA was recognized to co-localize with a wide variety of cytochemically

identified neuropeptides and amines in hypothalamic neurons (e.g. 84). Particularly, in the suprachiasmatic and arcuate nuclei, GABA was found in neurons that also contain

VIP, NPY, or somatostatin, among many other neuropeptides. These peptides have

profound effects on hypothalamic function. A primary mechanism underlying these

effects appears to be modulation of GABA neurotransmission. In the cerebral cortex and

the hippocampus, the neuropeptide CCK also co-localizes with GABA (e.g. 85).

38 To investigate whether GABA co-localizes with neuropeptides in the taste buds

(as the case in the brain), the GAD65/67 antibody was paired with another antibody directed against either CCK or VIP in a double-labeling immunofluorescent study performed on cross sections of rat foliate or circumvallate papillae. Indeed, considerable overlapping of GAD65/67-immunoreactivity with either CCK- or VIP-immunoreactivity was observed in TRCs on rat posterior lingual epithelium (Figure 3.6), indicating that co-

transmission of GABA and neuropeptides may occur in taste buds. As summarized in

Table 3.2, the majority of GAD65/67-immunopositive cells identified were double-

labeled with either CCK or VIP antiserum, whereas only about two-fifths of the

peptidergic TRCs also contained the GABA synthetic enzyme, suggesting heterogeneity

of GABAergic and peptidergic TRCs in terms of neurochemical composition.

Furthermore, peptide-containing TRCs were generally observed to outnumber those

immunopositive to GAD65/67 to a large extent, consistent with the difference noticed

between the two groups regarding the percentages of double labeled cells. The above

characteristics of the overlapping expression pattern of GABA and neuropeptides in taste

buds are in concert with the fact that immunoreactivities of GAD65/67, CCK or VIP have

all been found to partially co-localize with that of Gα-gust (session 3.1.1, ref. 72).

Therefore, TRCs with gustatory signal transduction functions have the potential ability to transmit complex information encoded by various endogenously-expressed neurotransmitters and neuropeptides.

39 3.2. Distributions of GABA receptors in taste buds of rat posterior lingual epithelium

3.2.1. Expression of the GABAA receptor α1 subunit in TRCs

Neurotransmitters like GABA exert their functions by activating a diversity of receptor subtypes. The ionotropic GABAA receptor subtype is thought to mediate the fast

inhibitory effects of GABA in the central nervous system (e.g. 53). Among all subunits

identified to compose functional GABAA receptors, the α1 subunit is the most abundant one in the mammalian brain while the α3 subunit is expressed in brain regions with low

levels of the α1 subunit (75). Antibodies against either of the two subunits were obtained

to examine possible expression of GABAA receptors as well as their subunit composition in taste buds of rat foliate and circumvallate papillae.

In western blotting experiments, immunoreactive bands to the antibody against the α1 subunit, but not the α3 subunit, was detected in membrane protein extracts of both anterior and posterior taste papillae (Figure 3.7). Consistently, cellular distribution of the

α1 subunit-immunoreactivity was observed in a small population of TRCs on the cross

sections of rat posterior lingual epithelium (Figure 3.7a), whereas the α3 subunit-

immunoreactivity was absent in any of the taste buds examined. Forebrain sections that

served as positive controls for localization of the α3 subunit-immunoreactivity were

processed in parallel with tongue sections of the same rats. Despite of failure to detect

any α3 subunit-immunoreactivity in the tongue sections, numerous α3 subunit-

immunoreactive neurons were observed in the forebrain sections (Figure 3.7b), validating

the α3 subunit antibody and the experimental procedure used in the present study.

Collectively, it is believed that subsets of TRCs express the GABAA receptor subtype,

40 whose subunit composition includes the α1 subunit; however, the α3 subunit is unlikely

to be present in taste buds.

On average, about two-thirds of the taste buds identified on an 8 µm cross section

of either foliate or circumvallate papillae contained cells positively labeled by the

GABAA receptor α1 subunit antibody (Table 3.1). Additionally, an average of 1.62 ±

0.08 (mean ± SE) cells in a 8 µm cross sectional area of a single taste bud exhibited

immunoreactivity to the GABAA receptor α1 subunit. As shown in Figure 3.7a, GABAA receptor α1 subunit-immunopositive TRCs typically exhibited morphological features similar to those of the TRCs identified to express signal transduction molecules. That is, these TRCs positively labeled with the α1 subunit antibody were elongate cells comprising a circular or oval nuclear region and processes extending from the apical end to the basement membrane of the taste buds. Immunoreactivity to the GABAA receptor

α1 subunit was diffusely distributed in the cytoplasmic area of TRCs but was absent in

the nuclear region. Positively stained cells were observed mostly in the central area of the

buds. No significant immnoreactivity to the GABAA receptor α1 subunit was recognized

in any of the nerve fibers inside or under the taste buds. The epithelium surrounding the

taste buds lacked specific immunostaining with the α1 subunit antibody.

3.2.2. Expression of the GABAB receptor R1 subunit in TRCs

The metabotropic GABAB receptors are believed to mediate the long-term inhibitory effects of GABA in the central nervous system. GABAB receptors belong to a particular group of G-protein coupled receptors due to the fact that heterodimer formation by the R1 and R2 subunits is required for the receptor to be physiologically functional.

41 Possible expression of GABAB receptors in rat taste buds was investigated using an antibody directed against the R1 subunit of the GABAB receptor. Immunoreactivity to the

GABAB receptor R1 subunit was observed exclusively in the taste buds in spindle-shaped cells demonstrating similar morphological characteristics as those of GABA-,

GAD65/67-, or GABAA receptor α1 subunit-immunoreactive TRCs (Figure 3.8). Only about half of the taste buds on each 8 µm cross section of posterior lingual epithelium displayed cells immunopositive to the GABAB receptor R1 subunit (Table 3.1). Moreover, fewer positively labeled cells were observed in a single taste bud cross sectional area when compared to that of GABA-, GAD65/67-, or GABAA receptor α1 subunit-

immunoreactive TRCs. Therefore, TRCs in adult rat posterior lingual epithelium appear

to differentially express GABA as well as essential subunits of its receptor subtypes,

GABAA and GABAB.

3.2.3. Distribution of GABAergic TRCs with respect to GABAB receptor subtype

Both GABAA and GABAB receptor subtypes could be localized either presynaptically or postsynaptically in the central nervous system. Presumably, GABA

may act as an autocrine or a paracrine factor inside the taste buds as well. Due to the

difficulty in finding an antibody of the GABAA receptor α1 subunit suitable for double

labeling immunocytochemical experiments using rat tongue sections, only the

distribution pattern of the GABAB receptor R1 subunit with respect to GABAergic TRCs was examined in cross sections of rat foliate and circumvallate papillae.

As revealed in Figure 3.9, GAD65/67-immunoreactive TRCs appeared as a separate group of cells from the one exhibiting immunoreactivity to the GABAB receptor

42 R1 subunit, indicating a paracrine role of GABA mediated by the GABAB receptor subtype in the taste buds.

3.2.4. Co-localization patterns of the GABAB receptor R1 subunit with SNAP-25, Gα-gust as well as neuropeptides CCK and VIP

To elucidate phenotypes of TRCs expressing the GABAB receptor R1 subunit, its co-localization patterns with SNAP-25, Gα-gust as well as neuropeptides CCK and VIP in rat posterior lingual epithelium was investigated.

On a typical cross section of either foliate or circumvallate papillae, there were clearly much more TRCs labeled with SNAP-25 antibody than those immunopositive to the GABAB receptor R1 subunit (Figure 3.10). Over seventy percent of the TRCs

displaying immunoreactivity to the GABAB receptor R1 subunit also demonstrated

SNAP-25-immunoreactivity, whereas no more than one fourth of the SNAP-25- expressing TRCs also expressed the GABAB receptor R1 subunit (Table 3.3). Therefore, most of the GABAB receptor R1 subunit in the taste buds appeared to be expressed by

TRCs with synapses and capable of releasing neuroactive substances via regulated

exocytosis. However, GABAB receptor-expressing cells only constituted a small

percentage in the group of TRCs with synapses.

Based on the previous findings that GAD65/67-immunoreactivity displayed a

non-overlapping pattern with the GABAB receptor R1 subunit-immunoreactivity but extensive overlap with immunoreactivities of Gα-gust, CCK or VIP, little co-expression of these three molecules was expected to be found with the GABAB receptor R1 subunit.

Interestingly, this is not completely true according to the results of double labeling

immunofluorescent experiments. As predicted, GABAB receptor R1 subunit-

43 immunoreactivity was not observed to co-localize with immunoreactivity to Gα-gust or

VIP (Figure 3.11). Surprisingly, a number of TRCs were discovered to exhibit double labeling with antibodies of both CCK and the GABAB receptor R1 subunit (Figure 3.12).

About seventy five percent of the GABAB receptor R1 subunit-immunoreactive TRCs

was also positively labeled with CCK antibody, while only about twenty five percent of

the CCK-immunopositive TRCs expressed immunoreactivity to the GABAB receptor R1 subunit (Table 3.3). This result suggests that a small percentage of CCK-expressing

TRCs may receive inhibitory input mediated by the GABAB receptor. This modulation is most likely to originate from neighboring GABAergic TRCs since GABA and the

GABAB receptor has been shown to be expressed by separate groups of cells in taste buds

in the previous session. Furthermore, these results suggest that peptidergic TRCs may be

differentially modulated by GABAB receptor-mediated mechanisms in taste buds. In a recent studies (96), the neuropeptide NPY was demonstrated to almost completely co- localize with either CCK or VIP in taste buds. Consistently, CCK and VIP overlapped

with each other partially. Considering that the GABAB receptor R1 subunit was partly

localized only in some CCK-expressing TRCs but not in any of the VIP-expressing cells,

it is reasonable to infer that NPY-expressing cells are unlikely to contain immunoreactivity to the GABAB receptor R1 subunit. Overall, CCK-expressing TRCs

may be modulated by GABA in a paracrine pathway through the GABAB receptor, whereas this would not be the case for other peptidergic cells containing either VIP or

NPY.

44 3.3. Physiological actions of GABA on TRCs in rat posterior lingual epithelium

According to our results of the immunocytochemical studies, GABA as well as the GABA receptor subtypes, GABAA and GABAB, could all be localized in subsets of

TRCs in the taste buds. Therefore it is hypothesized that endogenously released GABA in the taste buds may influence TRCs’ functions by activating different receptor subtypes.

To test this hypothesis, patch clamp recordings were performed on TRCs acutely dissociated from adult rat foliate and circumvallate papillae.

TRCs are excitable cells like neurons in the CNS. That is, these cells display a vast array of voltage-dependent currents including sodium, potassium, calcium, and chloride currents (18, 27, 29, 81). These currents not only set the resting membrane potential, participate in forming receptor potentials and action potentials in response to taste stimulation, but also determine the excitability and sensitivity of TRCs. Since voltage-dependent currents are often involved in the early stages of taste transduction (e.g.

19, 97), regulation of these currents serves as one of the primary mechanisms for neuroactive substances to influence TRC’s signaling functions. Indeed, most neurotransmitters (e.g. 5-HT and NA) and neuropeptides (e.g. CCK and NPY) found in the taste buds have been demonstrated to affect different ion conductances of TRCs to exert their physiological actions (summarized in Table 1.1). It is likely that GABA may also modulate TRC functions through regulation of different voltage-dependent ion channels as would be predicted based on inherent properties of different GABA receptor subtypes shown in the CNS. Possible physiological effects of GABA and receptor subtypes involved were tested using patch clamp recordings of different ion currents exhibited by TRCs.

45 3.3.1. Effects of GABA and the GABAA agonists on chloride currents recorded from TRCs

In vertebrate brains, the GABAA receptor subtype has been recognized as a ligand-gated anion channel mainly permeable to Cl-. Therefore isolated chloride currents were recorded from TRCs to explore possible GABAA receptor-mediated actions of

GABA on TRCs. With potassium-free extracellular and intracellular recording solutions, chloride currents displaying apparent outward rectification were recorded from all acutely dissociated rat posterior TRCs that satisfied criteria for either whole cell or perforated patch clamp recording mode (Figure 3.13a&b). When evoked by a ramp command potential that changing continuously between -130 to 120 mV from a holding potential of

0 mV, chloride currents typically displayed maximal outward amplitude less than 200 pA.

Recorded chloride currents could be effectively inhibited by various chloride channel blockers including 4-acteamido-4'-isothiocyanastilbene-2,2'-disuldonic acid (SITS), niflumic acid, or 9-anthracene carboxylic acid (9-AC), each tested at 500 µM, replicating our previous studies (29). Application of the calcium channel blocker cadmium (1 mM) was also effective in reducing the magnitude of chloride currents, suggesting the presence of a calcium-dependent component in these chloride currents. Modulation of chloride currents by neurotransmitters in the taste buds has been shown in a previous study demonstrating adrenergic enhancement of outward chloride currents mediated by β- adrenergic receptors (29). Results of the present study revealed that GABA and the

GABAA receptor agonists at various concentrations enhanced both outward and inward

portions of chloride currents recorded from TRCs (Figure 3.13b).

When GABA was applied focally to the recorded TRCs at a concentration series

ranging from 30 µM to 2 mM, it elicited concentration-dependent enhancement of

46 chloride currents that was reversible with washout of GABA from the bathing solution

(Table 3.4 and Figure 3.13). With increasing concentrations (30 µM ~ 2 mM), responses to GABA were observed in more TRCs tested (Figure 3.13d). For example, at 30 µM,

GABA induced enhancement of chloride currents amplitude in less than 15% (3 out of 21) of TRCs tested, whereas over 30% (7 out of 22) of the tested TRCs responded to 2 mM

GABA application. Additionally, percentages of current amplitude increase were around

25% with GABA applied at either low or high concentrations but peaked (approximately

35% increase) at GABA concentration of 300 µM (Figure 3.13c). Similar to adrenergic modulation of chloride currents, GABA at lower concentrations (e.g. 30 µM) appeared to affect the outwardly rectifying portion of the chloride currents (Figure 3.13a). On the other hand, enhancement of the inward fraction was more significant when GABA was applied at higher concentrations (e.g. 500 µM, Figure 3.13b).

To examine possible involvement of the GABAA receptor in the enhancing effects of GABA on chloride currents recorded from TRCs, the GABAA receptor-specific agonists muscimol and isoguvacine were applied at various concentrations. Muscimol is a naturally occurring GABA analog isolated from the psychoactive mushroom Amanita muscaria (45). It is a potent and specific agonist at GABAA receptors and has been a valuable tool for pharmacological studies. Particularly, muscimol has been extensively used as a lead for the design of different classes of GABA analogs. The conversion of muscimol into 4,5,6,7-tetrahydroisoxazolo[5,4-c]pyridin-3-ol (THIP) led to development of isoguvacine, one of the specific monoheterocyclic GABAA receptor agonists. While recording isolated chloride currents of TRCs, application of both muscimol (Figure 3.14) and isoguvacine (Figure 3.15) were able to induce elevation of currents amplitude in a 47 concentration-dependent manner (Table 3.4). Consistent with that observed following

GABA application, both agonists were able to enhance not only the outward portion but

also the inward portion of the chloride currents recorded from TRCs, implying

involvement of a GABAA receptor-mediated mechanism underlying GABA’s effects.

3.3.2. Effects of GABA and the GABAB agonist Baclofen on inwardly rectifying potassium

currents recorded from TRCs

In the mammalian brain, the metabotropic GABAB receptor is believed to couple to G proteins to modulate potassium channels (in most cases the inwardly rectifying potassium/KIR channels) postsynaptically and to modulate voltage-dependent calcium

channels presynaptically (mostly as autoreceptors). Since previous results demonstrated

that GABA and the GABAB receptor were expressed in distinct TRC populations, it is

unlikely that the GABAB receptor may act as presynaptic autoreceptors in the taste buds.

Therefore inwardly rectifying potassium currents were recorded from TRCs to test

possible postsynaptic effects of GABA mediated by the GABAB receptors.

KIR channels are ubiquitously expressed in TRCs of rat posterior taste buds (81).

They display pharmacological and eletrophysiological features commonly reported for this class of channel. These features include: activation by hyperpolarization, strong

inward rectification, dependence on voltage and extracellular potassium, and blockage by

cesium or barium. As in many other cell types, KIR may contribute to the resting potential of TRCs and help to stabilize membrane potential. Regulation of KIR conductance therefore may affect membrane excitability of TRCs. Two neuropeptides expressed in the taste buds, CCK and NPY, have been demonstrated to exert their modulatory effects on

TRC’s physiological functions by affecting KIR in opposite ways (34, 96). While CCK

48 inhibited KIR to elevate TRC’s excitability, NPY enhanced KIR to stabilize TRC’s

membrane potential thus causing it to be less excitable. Based on the reports of GABAB receptor-mediated inhibitory effects in the CNS, GABA is expected to produce an enhancing effect on KIR by activating the GABAB receptors expressed on TRCs, leading to membrane potential stabilization similar to that induced by NPY.

When applied focally, exogenous GABA at concentrations ranging from 500 µM to 2 mM effectively elevated the current magnitude at the inwardly rectifying portion of

KIR recorded in a subset of TRCs (Table 3.5, Figure 3.16). Enhancement of KIR was most significant at the highest GABA concentration (2 mM) tested (Figure 3.16c). Moreover,

highest percentage (30.43%, 7 out of 23) of responding TRCs was observed with 2 mM

GABA (Figure 3.16d).

To examine involvement of specific receptor subtypes, both GABAB receptor- specific agonist and antagonist were tested. Baclofen, a synthetic analog of GABA, is a potent GABAB receptor-specific agonist. When applied at 500 µM, baclofen produced

similar effects on KIR in approximately 30% of the TRCs recorded (Table 3.5, Figure

3.17). When baclofen (500 µM) was applied with the presence of the GABAB receptor-

specific antagonist CGP 35348 (500 µM), only 1 out of 10 (i.e. 10%) recorded cells still displayed increased current magnitude of KIR (Figure 3.18b). In contrast, none of the

other nine cells exhibited any alteration of recorded KIR while baclofen and CGP 35348 were applied together, even though all of them exhibited KIR elevation when baclofen was applied alone before and/or after addition of CGP 35348 (Figure 3.18a). Therefore activation of the GABAB receptor appeared to underlie GABA’s modulatory effect on

KIR expressed by TRCs. 49 In the CNS, the GABAB receptor is thought to couple to G proteins to indirectly

modulate potassium channels including KIR channels. Involvement of G protein in

GABAB receptor-mediated enhancement of KIR recorded from TRCs was verified as pretreatment with GDP-βs (2 mM), a nonhydrolyzable GTP analogue that inactivates G proteins by irreversibly binding to the alpha subunit, completely abolished baclofen’s enhancing effects on KIR in all of the 19 cells tested (Figure 3.18c&d). Therefore it appears that GABA released in the taste buds may affect TRC’s excitability by activating

G protein-regulated KIR channels.

Receptor desensitization is a general feature for G protein-coupled receptors such

as the GABAB receptor. This appeared to be no exception in the taste buds as significant reduction of response amplitude was observed when GABA (Figure 3.16e) or baclofen

(Figure 3.17d) was applied repeatedly at the same or different concentrations onto the same TRCs. Obvious desensitization of the GABAB receptor demonstrated in the present study implies uniformity between centrally and peripherally expressed GABAB receptors.

The mechanisms underlying desensitization of the GABAB receptor expressed by TRCs remain undiscovered.

In summary, GABA may influence TRC’s physiological functions by modulating

TRC’s excitability, which may be achieved by activating specific GABA receptor subtypes to regulate different ion conductance expressed by TRCs.

Antibody Total TBs Stained TBs % S/T Stained TRCs Mean (C/S) ± SE GABA 681 562 82.53 844 1.56 ± 0.04 GAD65/67 1083 831 76.73 1453 1.80 ± 0.05 GABAA α1 301 202 67.11 316 1.62 ± 0.08 GABAB R1 942 477 50.64 581 1.22 ± 0.02 Abbreviations: TBs – 8 µm cross section of taste buds, S – TBs containing stained cells, T – total TBs, TRCs – taste receptor cells, C – stained TRCs, SE – standard error, GABAB R1 – receptor R1 subunit, GABAA α1 – GABAA receptor α1 subunit.

Table 3.1. Cell counting results of single labeling immunocytochemistry.

Mean Mean Total GAD Gα-gust DL (% DL/GAD/TBs) (% DL/Gα-gust/TBs) TBs ± SE ± SE GAD & 147 269 531 146 55.49 ± 3.24 31.32 ± 2.30 Gα-gust Mean Mean Total GAD SNAP-25 DL (% DL/GAD/TBs) (% DL/SNAP-25/TBs) TBs ± SE ± SE GAD & 98 192 347 138 76.19 ± 2.63 42.47 ± 2.01 SNAP-25 Mean Mean Total GAD CCK DL (% DL/GAD/TBs) (% DL/CCK/TBs) TBs ± SE ± SE GAD & 78 157 281 113 77.24 ± 3.23 46.70 ± 3.14 CCK Mean Mean Total GAD VIP DL (% DL/GAD/TBs) (% DL/VIP/TBs) TBs ± SE ± SE GAD & 100 202 398 163 84.52 ± 2.27 43.77 ± 2.05 VIP Abbreviations: TBs – 8 µm cross section of taste buds, GAD – glutamate decarboxylase, Gα-gust – G protein gustducin α subunit, DL – double labeled, SE – standard error.

Table 3.2. Cell counting results of GAD65/67 double labeling immunocytochemical experiments.

Mean Mean Total R1 SNAP-25 DL (% DL/R1/TBs) (% DL/SNAP-25/TBs) TBs ± SE ± SE R1 & 90 109 328 80 73.33 ± 4.34 24.81 ± 1.74 SNAP-25 Mean Mean Total R1 CCK DL (% DL/R1/TBs) (% DL/CCK/TBs) TBs ± SE ± SE R1 & 52 67 205 50 76.92 ± 5.04 25.16 ± 1.86 CCK

Abbreviations: R1 – GABAB receptor R1 subunit, TBs – 8 µm cross section of taste buds, DL – double labeled cells, SE – standard error.

Table 3.3. Cell counting results of GABAB receptor R1 subunit double labeling immunocytochemical experiments.

Number of Number (%) of Responding value Drug Concentration TRCs recorded TRCs responded (% control value) 10 µM 22 0 (0%) − 30 µM 21 3 (14.29%) 122.71 ± 2.02 100 µM 44 7 (15.91%) 122.6 ± 2.11 GABA 300 µM 32 6 (18.75%) 134.99 ± 3.36 500 µM 118 31 (26.27%) 129.69 ± 3.26 1 mM 21 6 (28.57%) 122.69 ± 3.27 2 mM 22 7 (31.82%) 125.06 ± 4.56 10 µM 21 3 (14.29%) 121.55 ± 1.55 Muscimol 100 µM 60 19 (31.67%) 129.87 ± 3.79 500 µM 37 10 (27.03%) 129.53 ± 4.69 100 µM 34 14 (41.18%) 119.01 ± 1.42 Isoguvacine 500 µM 63 25 (39.68%) 139.87 ± 7.75

Table 3.4. Enhancing effects of GABA and the GABAA receptor agonists (muscimol and isoguvacine) on isolated chloride currents recorded from dissociated TRCs.

Number of TRCs Number (%) of Responding value Drug Concentration recorded TRCs responded (% control value) 500 µM 14 3 (21.43%) 121.33 ± 4.85 GABA 1 mM 24 5 (20.83%) 123.47 ± 6.27 2 mM 23 7 (30.43%) 130.48 ± 5.67 500 µM 59 19 (32.2%) 131.09 ± 3.55 Baclofen 1 mM 12 6 (50%) 120.93 ± 3.23

Table 3.5. Enhancing effects of GABA and the GABAB receptor agonist baclofen on inwardly rectifying potassium currents recorded from dissociated TRCs.

Number of TRCs Number (%) of TRCs Drug recorded responded Baclofen 500 µM + CGP35348 500 µM 10 1 (10%) Baclofen 500 µM + GDP-βs 2 mM 19 0 (0%)

Table 3.6. Effects of the GABAB receptor antagonist CGP35348 and the G protein

inhibitor GDP-βs on blocking baclofen’s enhancement of inwardly rectifying potassium

currents recorded from dissociated TRC

53 a. c. e.

b. d. f.

Figure 3.1. Localization of GABA-immunoreactivity in rat posterior TRCs. a & b, Foliate papillae. c & d, Circumvallate papillae. e & f, Cerebellum. Scale bar = 10 µm.

54 a. PT AT CTX CE - 65 kD b. c. d.

e. f. g.

Figure 3.2. Localization of GABA synthetic enzyme GAD65/67- immunoreactivity in rat posterior TRCs. a, Immunoreactive bands in western blot. PT, posterior tongue. AT, anterior tongue. CTX, cortex. CE, cerebellum. b-d, Foliate papillae. e-g, Circumvallate papillae. Scale bar = 10 µm.

55 GAD65/67 NCAM

GAD65/67 PGP9.5

Figure 3.3. Non-overlapping of GABA synthetic enzyme GAD65/67-immunoreactivity with immunoreactivity of NCAM (neural cell adhesion molecule) or PGP 9.5 (protein gene product 9.5) in rat posterior TRCs. Arrows indicate immunoreactivity to GAD65/67. Arrow heads indicate immunoreactivity to NCAM or PGP 9.5. Scale bar = 10 µm.

56 GAD65/67 Gα-gust

Figure 3.4. Overlapping of GABA synthetic enzyme GAD65/67- immunoreactivity with immunoreactivity of Gα-gust (G protein gustducin α subunit) in rat posterior TRCs. Arrows indicate double labeled TRCs. Arrow heads indicate single labeled GAD65/67-immunoreactive cells. Scale bar = 10 µm.

57 GAD65/67 SNAP-25

Figure 3.5. Overlapping of GABA synthetic enzyme GAD65/67- immunoreactivity with immunoreactivity of the presynaptic membrane-specific protein SNAP-25 in rat posterior TRCs. Arrows indicate double labeled TRCs. Arrow heads indicate single labeled GAD65/67-immunoreactive cells. Scale bar = 10 µm.

58 GAD65/67 CCK

GAD65/67 VIP

Figure 3.6. Overlapping of GABA synthetic enzyme GAD65/67- immunoreactivity with immunoreactivity of the neuropeptides CCK (cholecystokinin) or VIP (vasoactive intestinal peptide) in rat posterior TRCs. Arrows indicate double labeled TRCs. Arrow heads indicate single labeled GAD65/67-immunoreactive cells. Scale bar = 10 µm.

59 a. GABAA receptor α1 subunit CTX CE CV Fol AT - 50 kD

CE CV

b. GABAA receptor α3 subunit CTX CV Fol AT - 50 kD

Forebrain

Figure 3.7. Localization of the GABAA receptor α1 & α3 subunits in rat posterior TRCs. CTX, cortex. CE, cerebellum. CV, circumvallate papillae. Fol, foliate papillae. AT, anterior tongue. Arrow indicates a positively labeled cell. Scale bar = 20 µm.

60 CV Fol Forebrain Cerebellum

Figure 3.8. Localization of the GABAB receptor R1 subunit- immunoreactivity in rat posterior TRCs. Arrows indicate positively labeled cells. Scale bar = 10 µm.

61 GAD65/67 GABAB R1

Figure 3.9. Non-overlapping of GABA synthetic enzyme GAD65/67-immunoreactivity with immunoreactivity of the

GABAB receptor R1 subunit in rat posterior TRCs. Arrow heads indicate immunoreactivity to GAD65/67. Arrows indicate immunoreactivity to GABAB R1. Scale bar = 5 µm.

62 GABAB R1 SNAP-25

Figure 3.10. Overlapping of GABAB receptor R1 subunit- immunoreactivity with immunoreactivity of the presynaptic membrane-specific protein SNAP-25 in rat posterior TRCs. Arrows indicate double labeled TRCs. Arrow heads indicate single labeled GABAB R1 -immunoreactive cells. Scale bar = 5 µm.

63 GABAB R1 Gα-gust

GABAB R1 VIP

Figure 3.11. Non-overlapping of GABAB receptor R1 subunit- immunoreactivity with immunoreactivity of Gα-gust and the neuropeptide VIP in rat posterior TRCs. Arrows indicate immunoreactivity to GABAB R1. Scale bar = 5 µm.

64 GABAB R1 CCK

Figure 3.12. Overlapping of GABAB receptor R1 subunit- immunoreactivity with immunoreactivity of the neuropeptide CCK in rat posterior TRCs. Arrows indicate double labeled TRCs. Scale bar = 5 µm.

65 b. a. 200 I (pA) 400 I (pA) Before 30 µm GABA Rinse 150 Before 300 500 µM GABA 120 mV 100 Rinse 200 0 mV 0 mV

-130 mV 50 100

-120 -80 40 80 120 -120 -80 -40 80 120 0 0 V (mV) V (mV) -50 -100

-100 -200 c. d. 140 100

130 80 lue

a 60 V 120 rol s responding

l 40 110 6/21 7/22 Cont 31/118

% 6/32 % Cel 20 3/21 7/44 100 0 10 100 1000 30 100 300 500 1000 2000 GABA Conentration (µM) GABA Concentration (µM)

Figure 3.13. GABA’s enhancing effects on chloride currents recorded from TRCs. a & b, sample traces of chloride currents recorded from TRCs showing effects of GABA. c, concentration- response curve of GABA application. d, number of cells responding to various concentrations of GABA.

66 a. b. I (pA) 250 136

200 128 Before

150 e 100 µM Muscimol lu Rinse 100 a 120 l V o

50 r

-120 -80 -40 80 120 n 112 0 o V (mV)

-50 % C 104

-100 96 -150 1 10 100 1000 Muscimol Concentration (µM) c. 100

60 ponding

Res 40 s ll e 20 % C 3/21 19/60 10/37 0 10 100 500 Muscimol Concentration (µM)

Figure 3.14. Effects of the GABAA receptor agonists muscimol on chloride currents recorded from TRCs. a, sample traces of chloride currents recorded from TRCs showing effects of muscimol. b, concentration-response curve of muscimol application. c, number of cells responding to various concentrations of muscimol.

67 a. b.

200 I (pA) 150 n=25

Before 150 140 100 µM Isoguvacine e lu

Rinse a 100 130 V

l o 50 r n=14

nt 120 -120 -80 -40 40 80 120 0

V (mV) % Co 110

-50 100 100 500 -100 Isoguvacine Concentration (µM) c. 100

80 ng ndi 60 spo e 40 s R ll e

C 20 % 14/34 25/63 0 100 500 Isoguvacine Concentration (µM)

Figure 3.15. Effects of the GABAA receptor agonists isoguvacine on chloride currents recorded from TRCs. a, sample traces of chloride currents recorded from TRCs showing effects of isoguvacine. b, TRCs’ responding values with isoguvacine application. c, number of cells responding to two concentrations of isoguvacine.

68 a. b. I (pA) I (pA) 10 mV 200 150 -50 mV -50 mV 100 -140 mV V (mV) V (mV) 0 0 -140 -120 -100 -80 10 -140 -120 -100 -80 -60 10 -100

-200 -150

-300

-400 -300 Before Before 500 µM GABA -500 1 mM GABA Rinse Rinse -600 -450 c. d. 140 100

e 130 lu

a 60 V sponding

120 e

R 40

110

% Control 20 % Cells 3/14 5/24 7/23 100 0 0.51.01.52.0 500 µM 1 mM 2 mM GABA Concentration (mM) GABA Concentration e. 1st GABA application 170 2nd GABA application

160

150

Value 140

rol 130

120

% Cont 110

100

90 TRC#1 TRC#2 TRC#3

Figure 3.16. Effects of GABA on KIR recorded from TRCs. a & b, sample traces of KIR currents recorded from TRCs showing effects of GABA. c, concentration-response curve of GABA application. d, number of TRCs responding to GABA applied at various concentrations. e, receptor desensitization to repeated application of GABA.

69 a. I (pA) b. 400 150

200 e 140 u

l n=19 a

0 V 130 -140 -120 -100 -80 -60 10 n=6

V (mV) -200 120 % Control -400 110 Before 500 µM Baclofen -600 100 Rinse 500 µM 1 mM Baclofen Concentration c. d. 1st Baclofen application 100 150 2nd Baclofen application

g 80 140 e 130

60 Valu 120 40 ontrol 110

20 % C 100 % Cells Respondin 19/59 6/12 0 90 500 µM 1 mM TRC#1 TRC#2 Baclofen Concentration

Figure 3.17. Effects of the GABAB receptor agonist baclofen on KIR recorded from TRCs. a, sample traces of KIR currents recorded from TRCs showing effects of baclofen. b, responding values of baclofen application. c, number of TRCs responding to balcofen applied at two concentrations. e, receptor desensitization to repeated application of baclofen.

70 a. b. 150 100

140

n=19 ing 80 e

lu 130

Va 60

120 Respond

ntrol 40 110 lls

% Co n=9 20 100 % Ce 19/59 1/10 90 0 Baclofen Baclofen + Baclofen Baclofen + CGP 35348 CGP 35348 c. d.

150 100

140 n=19 80

130 60 Value sponding

120 e R 40 ntrol 110 Co n=19 20 %

100 % Cells 0 19/59 0/19 90 Baclofen Baclofen + Baclofen Baclofen + GDP-βs GDP-βs

Figure 3.18. The GABAB receptor antagonist CGP 35348 and the G protein blocker GDP-βs abolished effects of the GABAB receptor agonist baclofen on KIR recorded from TRCs. a & b, effects of balcofen with the presence of the GABAB receptor antagonist CGP 35348. c & d, effects of balcofen with the presence of the G protein blocker GDP-βs.

CHAPTER 4

DISCUSSION

The discovery of GABA in the brain over fifty years ago has inspired countless

studies that led to the unquestionable recognition of GABA’s critical role as the major

inhibitory neurotransmitter in virtually every region of the nervous system. Naturally,

GABA has been implicated in an enormous variety of functions and disorders. In the

sensory systems such as the visual and auditory systems, extensive studies have

suggested involvement of GABA in information processing at each level from the

periphery to the cerebral cortex. In the gustatory system, however, studies of GABA have

been mainly focused on the neuronal circuits in the central nervous system (e.g. 26, 42-44,

46, 48, 51, 60, 77-80), leaving GABA’s functional role in the periphery illusive. The

present study is the first to characterize the distribution patterns and functional

consequences of GABA in the peripheral end organ of the sense of taste, the taste buds.

Results presented in this study identified GABA as one of the neurotransmitters produced

endogenously by subsets of TRCs in the taste buds. Morphological features of

GABAergic TRCs were then recognized. Furthermore, distribution profiles of GABA receptor subtypes in the taste buds were demonstrated. Finally, functional studies revealed that GABA was able to modulate different electrical properties of TRCs by

72 acting on specific receptor subtypes. Therefore the present study provides valuable

information into current understandings of the role of GABA in peripheral gustatory signal processing.

4.1. Presence of GABAergic TRCs in the taste buds

4.1.1. GABA as a neurotransmitter in the taste buds

To establish a role of a neuroactive substance as a neurotransmitter in a neuronal circuit, several criteria have to be fulfilled (86). First, the substance must be generated in neurons within the circuit and stored in vesicles in the presynaptic terminals, from which it is released upon depolarization. Second, the released neurotransmitter must be able to act on its target cell to elicit postsynaptic responses, which are mediated by specific receptor subtypes. Third, removal (e.g. uptake) or inactivation (e.g. degradation) mechanisms must exist to effectively clear the neurotransmitters from the synaptic cleft.

Last, exogenous application of the substance to the native circuit must produce effects comparable to those induced by the naturally occurring neurotransmitter.

Results in the present study verified that GABA in the taste buds met nearly all of the above criteria therefore could be classified as a neurotransmitter that is able to participate in peripheral gustatory signal processing. First, the present study provided strong evidence confirming GABA’s origin in TRCs. GABA’s presence in the taste bud has been demonstrated in a previous study showing cellular localization of immunoreactivity to a GABA antibody in TRCs (59). In the present study, immunoreactivity not only to GABA itself, but also to the key synthetic enzyme for

GABA, GAD65/67, were localized in subsets of TRCs in rat posterior lingual epithelium,

73 indicating that TRCs are indeed the endogenous source of GABA in the taste buds.

Additionally, GAD65/67-immunoreactivity was found to largely co-localize with

immunoreactivity to the presynaptic membrane-specific protein SNAP-25 in a group of

TRCs. SNAP-25, together with the synaptic vesicle-associated protein, synaptobrevin,

are essential members of the protein complex formed during regulated vesicular release

of neurotransmitters. Since all SNAP-25-immunoreactive TRCs also displayed

immunoreactivity to synaptobrevin (93), it is reasonable to deduce that the majority of

GABAergic TRCs that express SNAP-25 may also express synaptobrevin. Together with

the findings that all synapse-bearing TRCs contained numerous synaptic vesicles (both

small clear ones and large dense-cored ones) and exhibited SNAP-25-immunoreactivity

(92), it is highly probable that GABA originated in most GABAergic TRCs may be

released from synaptic vesicles via regulated exocytosis upon cell activation.

Second, both GABA receptor subtypes, GABAA and GABAB receptors, were localized in subsets of TRCs. Moreover, under physiological conditions, GABA was able to induce enhancement of two classes of currents recorded from TRCs, chloride currents and KIR, which was mimicked by specific GABAA and GABAB receptor agonists, respectively, suggesting involvement of mechanisms mediated by different receptor subtypes. By affecting the chloride currents and KIR, GABA may influence TRC’s excitability and possibly alter functional consequences of TRCs during sapid stimulation.

Although the actual biological effects of GABA on TRCs were not demonstrated in the

present study, the results implied a functional role for GABA to modulate TRC

physiology in vivo thus provided important insights that may guide future research.

74 Finally, one of the membrane transporters for GABA uptake, GAT3, was found in

populations of TRCs (59), demonstrating the existence of GABA-removal mechanisms in

the taste buds. Collectively, results from the present study and the previous one all point

to a role for GABA to act as a neurotransmitter in the taste buds.

4.1.2. Phenotypes of GABAergic TRCs

TRCs are well-known to display incredible heterogeneity in terms of

morphological features and neurochemical composition, both of which have been

correlated to functional types of TRCs (e.g. 25). For example, crucial members of TRC

signal transduction machineries, including the three families of G protein coupled taste

receptors (i.e. the T1R2+3 heterodimer of sweet receptor, the T1R1+3 heterodimer of

amino acids receptor, and the T2R family of bitter receptor), the taste-specific G protein

gustducin, the downstream effector proteins PLCβ2, IP3R3 and TRPM5, as well as the

amiloride-sensitive sodium channel for the detection of sodium salts by rats, were all

localized in subsets of TRCs exhibited similar morphological characteristics, i.e., a

smooth outline of an elongate or spindle shape, a large clear round or oval nucleus

situated mostly in the middle part of the cell, and processes at the two ends extending the

whole length of the bud from the apical taste pore to the basement membrane.

Furthermore, taste receptor families are selectively expressed in subsets of TRCs so that

different groups of TRCs respond to compounds with different taste modalities,

indicating functional segregation of TRCs containing different neurochemical contents.

On the other hand, although these cells are known to be able to respond to various

gustatory stimuli by transmitting the presence of tastants into electrical signals that ultimately elicit transmitter release, they are not believed to be directly connected with

75 afferent nerve fibers inside the bud via synapses. Whereas another group of cells, which

could be identified by a slender elongate shape, a somewhat irregular-shaped nucleus that usually lies in the basal half of the taste bud, and a mostly central rather than peripheral location in the bud, is not believed to express the signal transduction molecules mentioned above but has been shown to form synapses onto nerve endings present inside the bud. Overall, morphological and neurochemical characterization of TRCs expressing a certain type of neuroactive substance is prerequisite and necessary for elucidating the

substance’s functional roles in peripheral gustatory signal processing.

The TRC population-specific protein markers chosen in the present study (i.e.,

NCAM, PGP 9.5, Gα-gust and SNAP-25) offer excellent representations of distinct

groups of TRCs. Collectively, populations of TRCs may be distinguished by these

markers signifying either the occurrence of synaptic connections either between adjacent

cells or with afferent nerve endings, the presence of the neurotransmitter 5-HT, or the

responsiveness to gustatory stimuli. Therefore double labeling immunocytochemistry

studies with these markers provided valuable information for identifying the phenotypes

of GABAergic TRCs as well as predicting their functional roles in peripheral taste signal

processing.

Based on the information gathered from the present study (summarized in Table

4.1), GABA is expressed by a subset of TRCs that mostly contain Gα-gust and SNAP-25,

but not NCAM or PGP 9.5. Together with other results of the present study revealing

GABAergic TRC’s morphological features and distribution profiles, it is suggested that

most of the GABAergic TRCs may be able to respond to gustatory stimuli (such as sweet

or bitter substances) but would not form synapses with the afferent nerves directly. 76 Therefore GABAergic TRCs may be active participants of gustatory signal processing in

the taste buds though they may not be the chemosensory cells which directly transmit signals to the brain.

4.1.3. Co-transmission in GABAergic TRCs

Co-transmission of neurotransmitters and neuropeptides in the nervous system has

been fully established for many years. Neuropeptides are thought to play important roles

in modulating various pre- or postsynaptic events therefore modulating the functions of

co-expressed neurotransmitters. This modulation greatly enriches the diversity of

synaptic signaling. This is likely to occur in the taste buds based on results obtained from

the present study regarding co-localization of GAD with neuropeptides CCK and VIP.

CCK was one of the first gastrointestinal peptides detected in the mammalian

brain (4, 85). It plays a key role in facilitating digestion within the small intestine. In the

brain, CCK is widely and abundantly distributed. In fact, CCK remains one of the most

abundant peptides in cerebral cortex, striatum and hippocampus. CCK is present in

many important neuronal pathways and is co-localized with several classic

neurotransmitters including GABA. Among its many biological effects, CCK has been

suggested to be responsible for the satiety signals to the brain. Behaviorally, CCK

injected into the ventricles of animal brain reduces food intake as a result of satiety

induction (e.g. 22). In addition, CCK is among the few neuropeptides closely related to

the peripheral gustatory system. CCK has been localized in subsets of TRCs and found

to exert an array of excitatory effects on TRCs by activating the CCK-A receptor (34, 52).

Interestingly, CCK and CCK-A receptor were discovered to be expressed in the same

cells, indicating an autocrine role of CCK (33). With some of the CCK-expressing cells

77 also express GABA, as demonstrated by the double labeling experiment in the present

study, CCK may act at its autoreceptors (i.e. the CCK-A receptor) to modulate GABA

release from the same cell. On the other hand, some of the CCK-expressing TRCs also

express the GABAB receptor, implying possible modulatory effect of GABA on CCK

release. Either way of modulation has been shown in various brain regions (4, 85). Since

some of the CCK-responding cells (which are likely to be CCK-expressing cells as well)

express Gα-gust and are bitter-responsive TRCs (52, 72), interactions between CCK and

GABA may modulate taste responses of these cells to participate in gustatory signal

processing in the taste buds. Additionally, CCK-expressing cells may receive complex

modulation from different neurotransmitters at different circumstances. Most CCK-

expressing TRCs were also sensitive to cholinergic stimulation (52), implying that ACh

may also be involved in modulating functions of these cells. It is possible that some of

the CCK-expressing cells may receive excitatory input from ACh in certain

circumstances while receive inhibitory input from GABA under other situations.

VIP is another widely distributed neuropeptide in the central and peripheral nervous system (87). As CCK, VIP often colocalizes with additional neuromodulators and neurotransmitters in various brain regions. Colocalization has been described with

GABAergic interneurons and CCK-containing neurons of the hippocampus. In the ventral thalamus colocalization occurs with CCK and in the reticular thalamic nucleus with GABA. Colocalization of VIP with either GABA or CCK (96) was also observed in subsets of the TRCs in rat posterior tongue, although functional consequences are hard to predict based on the current result.

NPY is another neuropeptide that is closely related to feeding and digestion. In

78 contrast to the case of CCK, injection of NPY into the hypothalamus potently increases

food intake (88). Like CCK, cellular distribution and physiological functions of NPY has

also been characterized in the taste buds (96). According to the discovery that NPY

colocalizes almost completely with either CCK or VIP in taste buds, it is highly likely

that NPY colocalizes with GABA in a group of TRCs as well, since GABA co-localizes

extensively with either CCK or VIP. Colocalization of GABA and NPY has been

described in many brain areas such as the hypothalamus, the striatum, the basal forebrain,

the hippocampus and the amygdale (88). Particularly, NPY exerts profound modulatory

effects in hypothalamus mainly through interaction with GABA or glutamate.

Modulatory interactions between NPY and GABA may also occur in the taste buds.

Future experiments are needed to fully verify this speculation.

Co-existence of GABA with 5-HT in TRCs is unlikely since GABA does not co-

localize with NCAM while 5-HT mostly co-localizes with NCAM. Evidence is lacking

about co-expression of their receptors in taste buds. Co-localization of neuropeptides and

5-HT is also unlikely since neither CCK nor VIP co-localizes with NCAM. 5-HT-

expressing TRCs clearly form a distinct group apart from that of TRCs express GABA,

neuropeptides, and taste signal transduction molecules.

4.2. GABA receptors in the taste buds

The present study demonstrated, for the first time, expression of two GABA receptor subtypes, GABAA and GABAB, in TRCs of adult rat posterior lingual epithelium.

Furthermore, the present study characterized the morphological features and distribution patterns of GABA receptor-expressing TRCs. More importantly, co-localization patterns

79 of the GABAB receptor with GAD, SNAP-25, Gα-gust, CCK or VIP were described,

which provided valuable information necessary for fully understanding GABAB receptor- mediated functional roles of GABA in taste buds.

GABAA and GABAB receptors belong to the inotropic and metabotropic receptor

superfamilies, respectively. Therefore, the GABAA receptor is thought to mediate the fast inhibitory neurotransmission effects of GABA whereas the GABAB receptor is thought to

mediate the slow and long-lasting neuromodulatory effects of GABA. As in many brain

regions, both GABA receptor subtypes are present in the same circuit, the taste bud.

However, it is difficult to speculate any interactions between the two in the taste bud

since colocalization and functional studies are lacking at the present time.

The two GABA receptor subtypes found in the taste buds both appeared to be

expressed in a manner resembles TRCs with known taste signaling functions, suggesting their possible involvement in peripheral gustatory signal processing. Even though double labeling experiments were only feasible with the antibody against GABAB receptor R1 subunit but not any of the subunits of the GABAA receptor, results from the present study revealed important implications of how GABA exerts its functions and what functional roles GABA may play in the taste buds. First, GABA is thought to act on the GABAB receptor in a paracrine pathway according to the non-overlapping distribution pattern of immunoreactivity to GAD and the GABAB receptor R1 subunit. Second, the GABAB

receptor did not appear to be co-expressed in Gα-gust-containing TRCs, suggesting

minimal involvement of GABAB receptor-mediated modulation in signal transduction processes of sweet or bitter substances in these cells. Finally, GABAB receptor-mediated

effects of GABA may differentially modulate properties of peptidergic TRCs, as 80 demonstrated by the differential co-expression patterns of the GABAB receptor with CCK or VIP. Collectively, GABAB receptor-mediated actions of GABA seems to mainly influence CCK-expressing TRCs that do not express GABA, VIP, NPY, or Gα-gust,

therefore are unlikely to directly modulate signal transduction of sweet or bitter

substances. It is possible, however, that GABAB receptor-mediated actions of GABA may modulate signal transduction of other taste modalities. Further morphological and functional characterization of GABA receptor-expressing TRCs are needed to fully understand their roles in peripheral gustatory signal processing in the taste buds.

4.3. Physiological actions of GABA on TRCs

As shown in the present study, GABA induced physiological responses of TRCs in patch clamp experiments, as previously described for the neurotransmitters 5-HT, NA, glutamate, ATP and ACh, as well as the neuropeptides CCK and NPY (Table 1.1).

Response profiles of GABA are consistent with those identified in the central nervous system and mainly include enhancement of chloride currents through a GABAA receptor- mediated mechanism and enhancement of KIR through a GABAB receptor-mediated mechanism.

Chloride currents and KIR can both serve to stabilize cell’s membrane potential.

Both classes of channels are expressed ubiquitously in TRCs (29, 81). Alteration of these channels has been shown with noradrenergic modulation of chloride currents as well as

CCK and NPY’s antagonist effects on KIR in rat TRCs (Table 1.1). The present study

now adds GABA to the list of neurotransmitters and neuropeptides with characterized

effects on the electrical properties of TRCs. By enhancing chloride currents via the

81 GABAA receptor subtype or KIR via the GABAB receptor subtype, GABA may result in

hyperpolarization of the TRC membrane potential, causing the cell to be less excitable.

Therefore, GABA exerts inhibitory effects on TRCs in the taste buds as it does to neurons

in the CNS. Presumably, GABA may affect TRC’s sensitivity to gustatory stimuli by

modulating cell excitability, fulfilling its role as an inhibitory intrinsic neuromodulatory

factor in the taste buds.

4.4. GABA’s role in peripheral gustatory signal processing

Sensory systems including the gustatory system endow us with the ability to perceive and appreciate the world around us. With further processing, sensory perception

then directs generation of appropriate behaviors in response to the environment. To

accomplish these goals, every sensory system needs to be highly selective to the

exceeding amount of information available to us, therefore modulation at each level of

the sensory pathway, from the peripheral receptor to the cerebral cortex, plays important

roles in filtering out redundant information, improving signal-to-noise ratios, and

ensuring optimal extraction of information in changing conditions (55). In these cases,

neuromodulators allow sensory inputs to be integrated in a way most favorable for

eliciting proper behaviors and allow perception to be influenced by a broad array of

factors including levels of arousal, attention, emotional stress and experience.

Modulation of sensory systems begins in the periphery, which may include both

intrinsic and extrinsic neuromodulation. Sharing many similarities with the carotid body,

the taste bud serves as an excellent model of intrinsic neuromodulation. Numerous

neurotransmitters and neuropeptides endogenously expressed in the taste buds may be the

82 main substrates carrying out intrinsic neuromodulatory functions by mediating communications between TRCs within the bud. As in the brain, where the overall activity

of neurons is basically determined by two superior functions – excitation by the major

excitatory amino acid transmitter glutamate as well as inhibition by the major inhibitory

neurotransmitter GABA, intrinsic neuromodulation in the taste buds may include

excitatory actions of 5-HT, NA, and CCK; as well as inhibitory actions mediated by

GABA or NPY. The present study provided first-hand information in favor of a role for the inhibitory neurotransmitter GABA in intrinsic neuromodulation within the taste bud.

First, the presence of GABA, GABA receptors, as well as their distribution

patterns in the taste buds, as discovered in the present study, all point to a paracrine role

for GABA to modulate TRCs functions, instead of a role for mediating direct sensory

transduction to the afferent nerves.

Second, as demonstrated in the present study as well as in the previous studies,

neurotransmitters and neuropeptides are expressed in complex distribution and co-

localization patterns in the taste buds. Adding another level of complexity, their receptor

subtypes are expressed differentially in the taste buds as well. Moreover, co-expression of

neurotransmitters and neuropeptides with signal transduction molecules occur extensively

in the taste buds. Most importantly, neurotransmitters or neuropeptides are able to exert

excitatory or inhibitory actions on TRCs by activating specific receptor subtypes

expressed in the taste buds. Taken together, a complicated network consisting of a great

diversity of neuroactive substances as well as their receptors seems to be present in the

taste buds. This network overlaps with another one consisting members of taste signal

transduction cascades in taste buds; thus is capable to execute modulatory functions for

83 fine tuning of the final sensory output of the latter one. GABA is undoubtedly one of the active members of this network since it fulfills every requirement (such as endogenous origin, specific receptors expression, and physiological actions) for this functional role as demonstrated in the present study. For example, bitter-responsive TRCs may contain

CCK and GABA as co-neurotransmitters. In response to bitter stimuli, these TRCs may release CCK to act on CCK-A autoreceptor to enhance excitability of themselves as a feed-forward mechanism. They may also release GABA to suppress neighboring GABAB receptor-expressing TRCs, which are not bitter-responsive, to increase signal-to-noise ratio of inputs to the chemosensory TRCs, as the mechanism of lateral inhibition seen in the visual system.

4.5. Concluding remarks and future directions

GABA is by now the least studied neurotransmitter in the peripheral taste organ, the taste bud. The present study for the first time characterized in details the morphological features, distribution patterns and physiological actions of GABA in TRCs of adult rat posterior lingual epithelium. GABA’s role as a neurotransmitter was established in the taste buds based on the results of the present study. Functional roles of

GABA in peripheral gustatory signal processing were explored by studying the phenotypes of GABAergic and GABA receptor-expressing TRCs as well as physiological responses of TRCs to exogenously applied GABA and specific receptor agonists. Results implied that GABA may participate in peripheral gustatory signal processing by mediating communication between neighboring TRCs. Therefore, GABA may act as an intrinsic neuromodulator in the taste buds to help shape the final output of chemosensory

84 TRCs. These results may serve as guidance for future studies. For example, nerve recording and behavioral studies may be designed and performed based on the results of the present study to provide more direct evidence of GABA’s involvement in peripheral gustatory signal processing.

85 GAD R1 Gα-gust NCAM PGP 9.5 SNAP-25 CCK VIP 5-HT GAD – + – – + + + –* R1 – – + + – Gα- + – –* – + + + –* gust NCAM – –* + + – – + PGP – – + +* – – – 9.5 SNAP- + + + + +* + + + 25 CCK + + + – – + + –* VIP + – + – – + + –* 5-HT –* –* + – + –* –*

Abbreviations: GAD – glutamate decarboxylase, R1 – GABAB receptor R1 subunit, Gα- gust – G protein gustducin α subunit, NCAM – neural cell adhesion molecule, PGP 9.5 – protein gene product 9.5, CCK – cholecystokinin, VIP – vasoactive intestinal peptide. “ * ” – inferred from experimental data

Table 4.1. Summary of protein co-localization in taste buds.

LIST OF REFERENCES

1. Adler E, Hoon A, Mueller KL, Chandrashekar J, Ryba NJP, Zuker CS. 2000. A novel family of mammalian taste receptors. Cell 100:693-702

2. Akisaka T, Oda M. 1978. Taste buds in the vallate papillae of the rat studied with freeze-fracture preparation. Arch Histol Jpn 41(1):87-98

3. Astback J, Arvidson K, Johansson O. 1995. Neurochemical markers of human fungiform papillae and taste buds. Regul Pept 59(3):389-98

4. Beinfeld MC. 2001. An introduction to neuronal cholecystokinin. Peptides 22(8):1197- 200

5. Baryshnikov SG, Rogachevskaja OA, Kolesnikov SS. 2003. Calcium signaling mediated by P2Y receptors in mouse taste cells. J Neurophysiol 90(5):3283-94

6. Bergner AJ, Murphy SM, and Anderson CR. 2000. After axotomy, substances P and vasoactive intestinal peptide expression occurs in pilomotor neurons in the rat superior cervical ganglion. Neuroscience 96: 611-618

7. Bigiani A, Delay RJ, Chaudhari N, Kinnamon SC, Roper SD. 1997. Responses to glutamate in rat taste cells. J Neurophysiol 77(6):3048-59

8. Bigiani A, Roper SD. 1994 Reduction of electrical coupling between Necturus TRCs, a possible role in acid taste. Neurosci Lett 176(2):212-6

87 9. Blednov YA, Walker D, Alva H, Creech K, Findlay G, Harris RA. 2003. GABAA receptor alpha 1 and beta 2 subunit null mutant mice: behavioral responses to ethanol. J Pharmacol Exp Ther 305(3):854-63

10. Borden LA. 1996. GABA transporter heterogeneity: pharmacology and cellular localization. Neurochem Int 29(4):335-56

11. Boughter JD Jr, Pumplin DW, Yu C, Christy RC, Smith DV. 1997. Differential expression of alpha-gustducin in taste bud populations of the rat and hamster. J Neurosci 17(8):2852-8

12. Bradley RM, King MS, Wang L and Shu X. 1996. Neurotransmitter and neuromodulator activity in the gustatory zone of the nucleus tractus solitarius. Chem Senses 21:377–385

13. Bystrova MF, Yatzenko YE, Fedorov IV, Rogachevskaja OA, Kolesnikov SS. 2006. P2Y isoforms operative in mouse taste cells. Cell Tissue Res 323(3):377-82

14. Caicedo A, Jafri MS, Roper SD. 2000. In situ Ca2+ imaging reveals neurotransmitter receptors for glutamate in TRCs. J Neurosci 20(21):7978-85

15. Caicedo A, Kim KN, Roper SD. 2000. Glutamate-induced cobalt uptake reveals non- NMDA receptors in rat taste cells. J Comp Neurol 417(3):315-24

16. Chaudhari N, Landin AM, Roper SD. 2000. A metabotropic glutamate receptor variant functions as a taste receptor. Nat Neurosci 3(2):113-9

17. Chaudhari N, Yang H, Lamp C, Delay E, Cartford C, Than T, Roper S. 1996. The taste of monosodium glutamate: membrane receptors in taste buds. J Neurosci 16(12):3817-26

18. Chen Y, Sun XD, Herness S. 1996. Characteristics of action potentials and their underlying outward currents in rat taste receptor cells. J Neurophysiol 75(2):820-31

88 19. Chen Y, Herness MS. 1997. Electrophysiological actions of quinine on voltage- dependent currents in dissociated rat taste cells. Pflugers Arch 434(3):215-26

20. Chung KM, Lee SB, Heur R, Cho YK, Lee CH, Jung HY, Chung SH, Lee SP, Kim KN. 2005. Glutamate-induced cobalt uptake elicited by kainate receptors in rat taste bud. Chem Senses 30(2):137-43

21. Damak S, Rong M, Yasumatsu K, Kokrashvili Z, Varadarajan V, Zou S, Jiang P, Ninomiya Y, Margolskee RF. 2003. Detection of sweet and umami taste in the absence of taste receptor T1r3. Science 301(5634):850-3

22. Della Fera MA, Baile CA. 1979. CCK-octapeptide injected in CSF causes satiety in sheep. Ann Rech Vet 10(2-3):234-6

23. Ewald DA and Roper SD. 1994. Bidirectional synaptic transmission in necturus taste buds. J Neurosci 14(6):3791-804

24. Finger TE, Danilova V, Barrows J, Bartel DL, Vigers AJ, Stone L, Hellekant G, and Kinnamon SC. 2005. ATP Signaling Is Crucial for Communication from Taste Buds to Gustatory Nerve. Science 310 (1495-99)

25. Finger TE, Simon SA. 2000. Cell biology of taste epithelium. In: The Neurobiology of Taste and Smell, Second Edition (Finger TE, Silver WL, Restrepo D eds.), Wiley-Liss, Inc. New York. Ch.12:287-314

26. Hasegawa K, Otawa S, Ikeda I, Ogawa H. 2003. Responsiveness of the cortical taste area neurons to a mixture of the four basic tastants in rats. Chem Senses 28(2):131-40

27. Herness MS, Sun XD. 1995. Voltage-dependent sodium currents recorded from dissociated rat taste cells. J Membr Biol 146(1):73-84

28. Herness MS, Chen Y. 2000. Serotonergic agonists inhibit calcium-activated potassium and voltage-dependent sodium currents in rat TRCs. J Membr Biol 173(2):127- 38

89 29. Herness MS, Sun SD. 1999. Characterization of chloride currents and their noradrenergic modulation in rat TRCs. J Neurophysiol 82:260-71

30. Herness MS. 1989. Vasoactive intestinal peptide-like immunoreactivity in rodent taste cells. Neuroscience 33(2):411-9

31. Herness S, Chen Y. 1997. Serotonin inhibits calcium-activated K+ current in rat TRCs. Neuroreport 8(15):3257-61

32. Herness S, Zhao FL, Kaya N, Lu SG, Shen T, Sun XD. 2002. Adrenergic signaling between rat TRCs. J Physiol 543(Pt 2):601-14

33. Herness S, Zhao FL, Kaya N, Shen T, Lu SG, Cao Y. 2005. Communication routes within the taste bud by neurotransmitters and neuropeptides. Chem Senses Suppl 1:i37- i38.

34. Herness S, Zhao FL, Lu SG, Kaya N, Shen T. 2002. Expression and physiological actions of cholecystokinin in rat TRCs. J Neurosci 22(22):10018-29

35. Huang YJ, Maruyama Y, Lu KS, Pereira E, Plonsky I, Baur JE, Wu D, Roper SD. 2005. Mouse taste buds use serotonin as a neurotransmitter. J Neurosci 25(4):843-7

36. Hunyady B, Krempels K, Harta G, and Mezey E. 1996. Immunohistochemical signal amplification by catalyzed reporter deposition and its application in double immunostaining. J Histochem Cytochem 44: 1353-1362

37. Jain S, Roper SD. 1991. Immunocytochemistry of gamma-aminobutyric acid, glutamate, serotonin, and histamine in Necturus taste buds. J Comp Neurol 307:675–682.

38. Kataoka S, Toyono T, Seta Y, Ogura T, Toyoshima K. 2004. Expression of P2Y1 receptors in rat taste buds. Histochem Cell Biol 121(5):419-26

39. Katz PS and Frost WN. 1996. Intrinsic neuromodulation: altering neuronal circuits from within. Trends Neurosci 19(2):54-61

90 40. Katz PS. 1999. What are we talking about? Modes of neuronal communication. In: Beyond neurotransmission : neuromodulation and its importance for information processing (Katz PS eds.), Oxford University Press. Ch. 1:1-29

41. Kim DJ, Roper SD. 1995. Localization of serotonin in taste buds: a comparative study in four vertebrates. J Comp Neurol 353:364-70

42. King MS. 2003. Distribution of immunoreactive GABA and glutamate receptors in the gustatory portion of the nucleus of the solitary tract in rat. Brain Res Bull 60(3):241- 54

43. Kobashi M, Bradley RM. 1998. Effects of GABA on neurons of the gustatory and visceral zones of the parabrachial nucleus in rats. Brain Res 799(2):323-8

44. Koga T, Bradley RM. 2000. Biophysical properties and responses to neurotransmitters of petrosal and geniculate ganglion neurons innervating the tongue. J Neurophysiol 84(3):1404-13

45. Krogsgaard-Larsen P, Frolund B, Liljefors T. 2002. Specific GABA(A) agonists and partial agonists. Chem Rec 2(6):419-30

46. Lasiter PS, Kachele DL. 1988. Organization of GABA and GABA-transaminase containing neurons in the gustatory zone of the nucleus of the solitary tract. Brain Res Bull 21:623–636.

47. Lawton DM, Furness DN, Lindemann B, Hackney CM. 2000 Localization of the glutamate-aspartate transporter, GLAST, in rat taste buds. Eur J Neurosci 12(9):3163-71

48. Leonard NL, Renehan WE, Schweitzer L. 1999. Structure and function of gustatory neurons in the nucleus of the solitary tract. IV. The morphology and synaptology of GABA-immunoreactive terminals. Neuroscience 92(1):151-62

49. Lin W, Finger TE, Rossier BC, Kinnamon SC. 1999. Epithelial Na+ channel subunits in rat taste cells: localization and regulation by aldosterone. J Comp Neurol 405:406-20

91 50. Lin W, Kinnamon SC. 1999. Physiological evidence for ionotropic and metabotropic glutamate receptors in rat taste cells. J Neurophysiol 82(5):2061-9

51. Lopez-Garcia JC, Bermudez-Rattoni F, Tapia R. 1990. Release of acetylcholine, gamma-aminobutyrate, dopamine and glutamate, and activity of some related enzymes, in rat gustatory neocortex. Brain Res 523(1):100-4

52. Lu SG, Zhao FL, Herness S. 2003. Physiological phenotyping of cholecystokinin- responsive rat taste receptor cells. Neurosci Lett 351(3):157-60

53. Macdonald RL. 1994. GABAA receptor channels. Annu Rev Neurosci 17:569-602

54. Martin DL, Tobin AJ. 2000. Mechanisms Controlling GABA Synthesis and Degradation in the Brian. In: GABA in the Nervous System: The View at Fifty Years (Martin DL, Olsen RW eds.), Lippincott Williams & Wilkins. Ch.2:25-42

55. Mercer AR. 1999. Changing the way we perceive things: sensory systems modulation. In: Beyond neurotransmission : neuromodulation and its importance for information processing (Katz PS eds.), Oxford University Press. Ch. 6:198-240

56. Nagai T, Delay RJ, Welton J, Roper SD. 1998 Uptake and release of neurotransmitter candidates, [3H]serotonin, [3H]glutamate, and [3H]gamma-aminobutyric acid, in taste buds of the mudpuppy, Necturus maculosus. J Comp Neurol 392(2):199-208

57. Nicoll RA. 2004. My close encounter with GABAB receptors. Biochem Pharmacol 68(8):1667-74

58. Nurse CA. 2005. Neurotransmission and neuromodulation in the chemosensory carotid body. Auton Neurosci 120(1-2):1-9

59. Obata H, Shimada K, Sakai N, Saito N. 1997. GABAergic neurotransmission in rat taste buds: immunocytochemical study for GABA and GABA transporter subtypes. Brain Res Mol Brain Res 49(1-2):29-36

92 60. Ogawa H, Hasegawa K, Otawa S, Ikeda I. 1998. GABAergic inhibition and modifications of taste responses in the cortical taste area in rats. Neurosci Res 32(1):85- 95

61. Ogura T. 2002. Acetylcholine increases intracellular Ca2+ in taste cells via activation of muscarinic receptors. J Neurophysiol 87(6):2643-9

62. Olsen RW, MacDonald RL. 2002. GABAA receptor complex: structure and function. In: Glutamate and GABA receptors and transporters: structure, function and pharmacology (Egebjerg J, Schousboe A, Krogsgaard-Larsen P eds), Taylor & Francis. Ch.9: 202-35

63. Ozeck M, Brust P, Xu H, Servant G. 2004. Receptors for bitter, sweet and umami taste couple to inhibitory G protein signaling pathways. Eur J Pharmacol 489(3):139-49

64. Pecina S, Berridge KC. 1996. Brainstem mediates diazepam enhancement of palatability and feeding: microinjections into fourth ventricle versus lateral ventricle. Brain Res 727(1-2):22-30

65. Perez CA, Huang L, Rong M, Kozak JA, Preuss AK, Zhang H, Max M, Margolskee RF. 2002. A transient receptor potential channel expressed in taste receptor cells. Nat Neurosci 5(11):1169-76

66. Pumplin DW, Getschman E. 2000. Synaptic proteins in rat taste bud cells: appearance in the Golgi apparatus and relationship to alpha-gustducin and the Lewis(b) and A antigens. J Comp Neurol 427(2):171-84

67. Pumplin DW, Yu C, Smith DV. 1997. Light and dark cells of rat vallate taste buds are morphologically distinct cell types. J Comp Neurol 378(3):389-410

68. Roach J, Roper SD. 1996. Gap junction proteins in rat vallate taste buds. Society For Neuroscience Abstracts 22:1825

69. Roper SD. 1992. The microphysiology of peripheral taste organs. J Neurosci 12(4):1127-34

93 70. Schousboe A, Kanner B. 2002 GABA transporters: functional and pharmacological properties. In: Glutamate and GABA receptors and transporters: structure, function and pharmacology (Egebjerg J, Schousboe A, Krogsgaard-Larsen P eds), Taylor & Francis. Ch.14: 337-50

71. Scott K. 2005. Taste recognition: food for thought. Neuron 48:455-64

72. Shen T, Kaya N, Zhao FL, Lu SG, Cao Y, Herness S. 2005. Co-expression patterns of the neuropeptides vasoactive intestinal peptide and cholecystokinin with the transduction molecules alpha-gustducin and T1R2 in rat taste receptor cells. Neuroscience 130(1):229- 38

73. Shimura T, Watanabe U, Yanagawa Y, Yamamoto T. 2004. Altered taste function in mice deficient in the 65-kDa isoform of glutamate decarboxylase. Neurosci Lett 356(3):171-4

74. Shindler KS and Roth KA. 1996. Double immunofluorescent staining using two unconjugated primary antisera raised in the same species. J Histochem Cytochem 44: 1331-1335

75. Sieghart W, Sperk G. 2002. Subunit composition, distribution and function of GABA(A) receptor subtypes. Curr Top Med Chem 2(8):795-816.

76. Simeone TA, Donevan SD and Rho JM. 2003. Molecular biology and ontogeny of γ- aminobutytic acid (GABA) receptors in the mammalian central nervous system. J Child Neurol 18(1):39-48

77. Smith DV, Li CS. 1998. Tonic GABAergic inhibition of taste-responsive neurons in the nucleus of the solitary tract. Chem Senses 23(2):159-69

78. Smith DV, Li CS. 2000. GABA-mediated corticofugal inhibition of taste-responsive neurons in the nucleus of the solitary tract. Brain Res 10;858(2):408-15

79. Smith DV, Liu H, Vogt MB. 1994. Neural coding of aversive and appetitive gustatory stimuli: interactions in the hamster brain stem. Physiol Behav 56:1189–96

94 80. Soderpalm AH, Hansen S. 1998. Benzodiazepines enhance the consumption and palatability of alcohol in the rat. Psychopharmacology (Berl) 137(3):215-22

81. Sun XD, Herness MS. 1996. Characterization of inwardly rectifying potassium currents from dissociated rat taste receptor cells. Am J Physiol 271(4 Pt 1):C1221-32

82. Tateda H, Beidler LM. 1964. The receptor potential of the taste cell of the rat. J Gen Physiol 47:479-86

83. Toyono T, Seta Y, Kataoka S, Harada H, Morotomi T, Kawano S, Shigemoto R, Toyoshima K. 2002. Expression of the metabotropic glutamate receptor, mGluR4a, in the taste hairs of taste buds in rat gustatory papillae. Arch Histol Cytol 65(1):91-6

84. Van den Pol AN, Obrietan K. 2000. Neuromodulation of GABA in Developing and Mature Hypothalamic Neurons. In: GABA in the Nervous System: The View at Fifty Years (Martin DL, Olsen RW eds.), Lippincott Williams & Wilkins. Ch.23:409-38

85. Von Bohlen und Halbach O, Dermietzel R. 2002. Cholecystokinin. In: Neurotransmitters and neuromodulators: handbook of receptors and biological effects (Von Bohlen und Halbach O, Dermietzel R eds; language editing by Ballantyne DJ), Wiley-VCH. Ch.4:116-271

86. Von Bohlen und Halbach O, Dermietzel R. 2002. Neurotransmitters. In: Neurotransmitters and neuromodulators: handbook of receptors and biological effects (Von Bohlen und Halbach O, Dermietzel R eds; language editing by Ballantyne DJ), Wiley-VCH. Ch.1:1-18

87. Von Bohlen und Halbach O, Dermietzel R. 2002. Vasoactive intestinal peptide (VIP). In: Neurotransmitters and neuromodulators: handbook of receptors and biological effects (Von Bohlen und Halbach O, Dermietzel R eds; language editing by Ballantyne DJ), Wiley-VCH. Ch.4:116-271

88. Von Bohlen und Halbach O, Dermietzel R. 2002. Neuropeptide Y (NPY). In: Neurotransmitters and neuromodulators: handbook of receptors and biological effects (Von Bohlen und Halbach O, Dermietzel R eds; language editing by Ballantyne DJ), Wiley-VCH. Ch.4:116-271

95 89. Wang G, Achim CL, Hamilton RL, Wiley CA, and Soontornniyomkij V. 1999. Tyramide signal amplification method in multiple-label immunofluorescence confocal microscopy. Methods 18: 459-464

90. Watanabe M, Maemura K, Kanbara K, Tamayama T and Hayasaki H. 2002. GABA and GABA receptors in the central nervous system and other organs. Int Rev Cytol 213:1- 47.

91. Xu H, Staszewski L, Tang H, Adler E, Zoller M, Li X. 2004. Different functional roles of T1R subunits in the heteromeric taste receptors. Proc Natl Acad Sci U S A 101(39):14258-63

92. Yang R, Crowley HH, Rock ME, Kinnamon JC. 2000. Taste cells with synapses in rat circumvallate papillae display SNAP-25-like immunoreactivity. J Comp Neurol 424(2):205-15

93. Yang R, Stoick CL, Kinnamon JC. 2004. Synaptobrevin-2-like immunoreactivity is associated with vesicles at synapses in rat circumvallate taste buds. J Comp Neurol 471(1):59-71

94. Yang R, Tabata S, Crowley HH, Margolskee RF, Kinnamon JC. 2000. Ultrastructural localization of gustducin immunoreactivity in microvilli of type II taste cells in the rat. J Comp Neurol 425(1):139-51

95. Yee CL, Yang R, Bottger B, Finger TE, Kinnamon JC. 2001. “Type III” cells of rat taste buds: immunohistochemical and ultrastructural studies of neuron-specific enolase, protein gene product 9.5, and serotonin. J Comp Neurol 440: 97-108

96. Zhao FL, Shen T, Kaya N, Lu SG, Cao Y, Herness S. 2005. Expression, physiological action, and coexpression patterns of neuropeptide Y in rat taste-bud cells. Proc Natl Acad Sci U S A 102(31):11100-5

97. Zhao FL, Lu SG, Herness S. 2002. Dual actions of caffeine on voltage-dependent currents and intracellular calcium in taste receptor cells. Am J Physiol Regul Integr Comp Physiol 283(1):R115-29

96 98. Zhao GQ, Zhang Y, Hoon MA, Chandrashekar J, Erlenbach I, Ryba NJ, Zuker CS. 2003. The receptors for mammalian sweet and umami taste. Cell 115(3):255-66 cells. Chem Senses 30(2):137-43.

APPENDIX

LOCALIZATION AND FUNCTIONAL INVESTIGATION

OF THE TRANSCRIPTION FACTOR CREB IN TASTE RECEPTOR CELLS

100

101

102

103