Functional Expression and Characterization of Histamine-gated Chloride Channels Arunesh Saras Ph.D. Dissertation To be presented by permission of the department of Cellphysiology of the Ruhr-University, Bochum

& International Graduate School of Neuroscience

2005

Contents

i

Contents Chapter 1: Introduction

1.1 GABA-receptors and the GABAergic system

1.1.1 The inhibitory γ-aminobutyric acid system 1

- a general overview

1.1.2 GABA-receptors: GABAB and GABAC 1

1.1.3 General properties of GABAA receptors 2

1.1.4 Types of heteromultimeric GABAA receptors 4

and their location and properties

1.1.5 Homomultimeric GABAA receptors 5

1.1.6 Trafficking of GABA-receptors and interacting proteins 6

1.1.7 Potentiation and modulation of GABAA receptors 8

1.1.7.1 Modulation of GABAA receptors by Propofol 8

1.1.7.2 Modulation of GABAA receptors by further 13

chemicals

1.1.8 Function of distinct GABAA subunits in vivo 14

investigated by knockout mice

1.2 Histamine-receptors and the histaminergic system 17

1.2.1 Histamine in the nervous system 17

1.2.2 Metabotropic histamine receptors 18

1.2.3 Interaction of histamine antagonists with GABAA receptors 22

1.2.4 Ionotropic histamine receptors and direct modulatory 22

action of histamine to ion channels

1.2.5 Histamine functions and knockout mice 24

1.2.6 Diseases where histamine is involved 25

1.2.7 Aims of the work 27

Contents

ii

Chapter 2: Materials 28

2.1 Chemicals and enzymes 28

2.2 Drugs used for pharmacological characterizations 29

2.3 Primers 31

2.4 Standards for DNA 32

2.5 Consumption materials 32

2.6 Kits 33

2.7 RNase free materials and chemicals 33

2.8 Frequently used buffer 34

2.9 Bacterial strains 37

2.10 Plasmid vectors 37

2.11 Softwares 38

Chapter 3: Methods 39

3.1 Characterizing, isolating and concentrating nucleic acids 39

3.1.1 Determination of concentrations of nucleic acids 39

3.1.2 Gel electrophoresis 39

3.1.3 Phenol: chloroform extraction of nucleic acids 39

3.1.4 Ethanol precipitation of nucleic acids 40

3.1.5 QIAquick PCR-Purification Kit 41

3.1.6 RNA Extraction 42

3.1.7 Quick preparation of plasmid DNA 42

3.1.8 Maxi preparation of plasmid DNA using the 43

QIAGEN Plasmid Maxi Kit

3.2 PCR Methods 43

3.2.1 Reverse transcription 43

Contents

iii

3.2.3 PCR-based generation of chimeric cDNAs and 45 site-directed mutagenesis

3.2.4 Phosphorylation op PCR primers 46

3.3 Cloning of DNA 46

3.3.1 Restriction 46

3.3.2 Dephosphorylation 46

3.3.3 Polishing of DNA using T4 DNA polymerase and T4 PNK 47

3.3.4 Fill in reaction 47

3.3.5 Ligation of DNA 48

3.3.6 Culturing of bacteria 48

3.3.7 Transformation of plasmid DNA 48

3.3.8 Sequencing of DNA 48

3.4 RNA techniques 48

3.4.1 In vitro transcription 49

3.5 Functional expression of LGICs in Xenopus laevis 50

3.5.1 Surgery 50

3.5.2 Oocyte preparation and injection of cRNA 50

3.5.3 Electrophysiological recording using two-electrode 51

voltage clamp

3.6 Functional Expression of LGICs in HEK 293 cells 52

3.6.1 Culture of HEK 293 cells and transfection 52

3.6.2 Patch clamp investigation of GABA receptors 53

expressed in HEK 293 cells

Chapter 4: Results I 55

4.1 Bioinformatical search for histamine-gated channels 55

4.2 Construction of expression vectors for GABAA receptors 56

4.3 Establishing functional expression GABAA receptors in 61

Xenopus oocytes

4.4 Direct effects of histamine on heteromultimeric GABAA receptors 65

4.5 Modulation of heteromultimeric GABAA receptors by histamine 65

Contents

iv

4.5.1 Potentiation of α1β1 GABAA receptors by histamine 65

4.6.1 Potentiation of α1β1 GABAA receptors by histidine 69

4.6.2 Characterization of histidine potentiation 70

4.6.3 Dependence of average histamine and histidine 72

potentiation on GABA concentration

4.6.4 Histidine does not alter the I/V curve and retains 73

selectivity for the permeability of the channel

4.7 Potentiation of GABAA receptors in HEK 293 cells 74

4.8 Dependence of histamine potentiation on GABAA receptor 76

subunit combinations

4.9 Homomultimeric channels of β1 subunit and the effect of 81

histamine and histidine

4.9.1 Homomultimeric channels of β3 subunit and effect of histamine 84

4.10 Molecular cloning of ρ1 subunit of GABAC receptors 84

4.10.1 GABAC receptors: No potentiation by histamine and histidine 86

4.11 Possible mechanisms of the histamine action 87

4.11.1 Histamine binding site is different from pentobarbital 87

binding-site on β3 subunit of GABAA receptors

4.11.2 Histamine binding site is similar to propofol binding- 88

site on β3 subunit of GABAA receptors –

Experiments on homomultimeric β3 subunit

4.11.3 Histamine binding site is similar to propofol binding 90

Site on β3 subunit of GABAA receptors

- Experiments on heteromultimeric α1β1 receptors

4.11.4 Effect of histamine on EC50 of GABA on GABAA receptors 91

4.11.5 Molecular cloning of point mutation in β1 subunits 96

4.11.6 β1(M286W) mutation completely abolishes potentitaion 96

mediated by histamine

4.11.7 Histamine and propofol have similar binding sites 98

4.11.8 β1 (M286W) mutation completely abolishes potentitaion 101

mediated by histidine

4.11.9 Sequence alignment with GABAC receptors depicts that 101

histamine has similar binding site to propofol

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v

4.11.10 β1 (M286W) mutation does not interfere with the 102

potentiaion mediated by other modulators

Chapter 5: Results II 106

5. Characterization of homomultimeric β3 channels 106

5.1 Homomultimeric β3 receptors behave like histamine-gated ion channels 106

5.2 Histamine gated homomultimeric β3 receptors behave like typical 106

ligand-gated chloride channels

5.3 Relative comparison of various agonist of histamine with GABA 110

- relative agonists efficacy compared to GABA

5.4 Pharmacological characterization of histamine-gated homomultimeric 112

β3 receptors

5.4.1 Thioperamide acts as a competitive blocker for 113

histamine-evoked current

5.5 Comparison of L- and D-histidine action on β3 homomultimeric receptors 116

5.6 Inhibition of Propofol-induced current by thioperamide on 118

homomultimeric β3 receptors

Chapter 6: Results III 121

6. Characterization of homomulitmeric β2 channels 121

6.1 Homomultimeric β2 receptors behave like histamine-gated ion channels 121

6.2 Histamine gated homomultimeric β2 receptors behave like typical ligand- 122

gated chloride channels.

6.3 Relative comparison of various agonists of histamine with GABA 122

6.4 Pharmacological characterization of histamine-gated homomultimeric 122

β2 receptors

6.5 Relative comparison of homomultimeric β2 and β3 homomultimeric 126

receptors

Contents

vi

Chapter 7: Results IV 132

7. Characterization of homomultimeric ε subunit 132

7.1 Characterization of ε subunit containing receptors 132

7.2 ε contianing receptors behave like histamine-gated ion channels 133

7.3 Pharmacology of ε subunit 135

7.4 Expression of α1β1ε, β1ε and ε in HEK 293 cells 136

7.5 Molecular cloning of GFP-tagged subunits of GABAA receptors 136

7.5.1 Molecular cloning of GFP-tagged α1 and β1 subunits of 137

GABAA receptors

7.5.2 Molecular cloning of GFP-tagged ε subunit of 137

GABAA receptor

7.6 Expression of α1, β1 and ε in HEK 293 cells 142

Chapter 8: Results V 145

8.1 Properties of α1β2γ2 receptors and direct activation by histamine 145

8.2 Homomultimeric γ2 receptors: Activation by histamine and its 147

metabolites

Chapter 9: Discussion 150

Chapter 10: Summary 167

Chapter 11: References 169

Appendix: 189

Appendix I 189

Appendix II 190

Acknowledgements 191

Curriculum Vitae 192

Chapter 1 Introduction

1

1. INTRODUCTION

1.1: GABA-receptors and the GABAergic system

1.1.1: The inhibitory γ-aminobutyric acid system - a general overview

Gamma–aminobutyric acid (GABA) is the major inhibitory neurotransmitter in the

mammalian central nervous system. It regulates many physiological functions and

emotional and cognitive behaviors through neurosynaptic contacts widespread in the

brain (Costa, 1982). In the mammalian brain the GABA is synthesized primarily from

glutamate in a reaction which is catalyzed by two glutamic acid decarboxylase (GAD)

enzymes, GAD65 and GAD67 (Bloom and Iversen, 1971). In the synaptic vesicle GABA

is loaded by a vesicular neurotransporter (VGAT) (Fon and Edwards, 2001) and it is

liberated into nerve terminal by calcium dependent exocytosis. However, no-vesicular

GABA secretion is being described and might play a role during development (Attwell et

al., 1993; Taylor and Gordon-Weeks, 1991). The effect of GABA can be mediated by

ionotropic or metabotropic receptors, which are localized post - or presynaptically. The

termination of GABA activation can happen either by its reuptake into the nerve

terminals or surrounding glial cells by a class of plasma membrane GABA transporters

(GATs) (Cherubini and Conti, 2001). Thereafter, GABA is metabolized by a

transamination reaction that is catalyzed by GABA transaminase (GABA-T). GABA acts

on 3 types of receptors which are phylogenitically conserved across different species:

GABAA, GABAB and GABAC receptors (Friedl et al., 1988).

1.1.2: GABA-receptors: GABAB and GABAC

GABAB receptors are bicuculline insensitive, chloride independent, metabotropic

receptors (Hill and Bowery, 1981; Bowery et al., 1980; Nicoll, 1988) and belong to the

superfamily of G-protein coupled receptors. GABAB receptors were shown to mediate

Chapter 1 Introduction

2

presynaptic inhibition on some nerve endings and postsynaptic inhibition on some cell

bodies or dendrites. GABAB receptors exist as GABAB1a / GABAB2 and GABAB1b /

GABAB2 and are associated with G–proteins. They have seven transmembrane domains.

GABAB receptors are localized both pre- and postsynaptically and they use different

mechanisms at these locations to regulate cell excitability. Presynaptic inhibition occurs

through a GABAB receptor mediated reduction in calcium current at the nerve terminal

and a subsequent reduction in transmitter release, whereas postsynaptic inhibition occurs

by GABAB receptor mediated activation of potassium currents that hyperpolarize the

neuron (Connors et al., 1988).

Like GABAA receptors, GABAC receptors are ligand-gated ion channel receptors (Sigel,

1995; Johnston, 1996; Enz and Cutting, 1998). This receptor is a chloride-selective ion

channel, but is insensitive to the GABAA receptor antagonist bicuculline (Bormann and

Feigenspan, 1995). GABAC receptors are believed to be homo - or heteropentameric

proteins that are composed of a single or multiple ρ subunits. They are also different from

GABAB receptors being insensitive to baclofen but are responsive to cis-4-aminocrotonic

acid, a structural analogue of GABA. GABAC receptors can be considered as

pharmacological variants of GABAA receptors (Mehta and Ticku, 1999; Bormann, 2000).

1.1.3: General properties of GABAA receptors

The GABAA receptors are members of the ligand-gated ion channel superfamily, which

also includes nicotinic acetylcholine, glycine and serotonin (5-HT3) receptors. GABAA

receptors are the primary mediators of GABA-induced rapid inhibitory neurotransmission

(Sieghart, 1995) and are believed to be heteropentameric proteins that are constructed

from subunits derived from several related genes or gene families (Macdonald and Olsen,

1994). At present, six α subunits, three β subunits, three γ subunits, one δ subunit, one ε

subunit, one π subunit and one θ subunit have been identified in mammals (Macdonald

and Olsen, 1994; Schofield et al., 1987; Mehta and Ticku, 1999). All the subunits are

related to each other and have molecular weights of about 50 kD. These various subunits

provide enormous subunit combinations but only certain subunit combinations are

Chapter 1 Introduction

3

preferred (McKernan and Whiting, 1996). Native receptors contain at least one α, one β

and one γ subunit. The δ, ε, π and θ subunits able to substitute for the γ–subunit

(McKernan and Whiting, 1996). GABAA receptors are integral membrane proteins,

which are formed by assembly of five homologous subunits around a central ion channel

(Chang et al., 1996). Each subunit has a large extracellular N-terminal domain and a C-

terminal domain containing four transmembrane segments, designated M1-M4, and

connected by relatively short loops. The extracellular N-terminal domains are believed to

form the agonist binding sites, whereas the transmembrane domains form the channel;

with the five M2 domains being the primary lining of the ion-conducting pore of the

receptor (Xu and Akabas, 1996). The M2 domain is thought to be a key channel-lining

component, which determines channel properties such as conductance, rectification, and

desensitization. Determined by the pore-forming M2-region, GABAA receptors carry

primarily chloride ions, however other anions, such as bicarbonate (HCO3-) can also

permeate the channel pore, although less efficiently (Kaila, 1994; Bormann et al., 1987;

Moss and Smart 2001).

Fig. 1.1: Structure of GABAA receptors. Proposed structure of a ligand gated ion channel. A receptor subunit contains four hydrophobic transmembrane (TM) domains. TM2 is believed to form the lining of the ion channel. The large amino terminal domain is located extracellularly and believed to incorporate neurotransmitter and some modulators binding sites. The intracellular domain in between TM3 and TM4 comprises ~ 10 % of the mass of each subunit. This domain is the most divergent part of individual receptor subunits and contains numerous consensus sites for the action of both serine/threonine and lysine protein kinases. Adapted from (Moss and Smart, 2001).

Chapter 1 Introduction

4

1.1.4: Types of heteromultimeric GABAA receptors and their location and

properties

Although molecular biology revealed seven types of homologous GABAA subunit types

by now, the subunit composition and the arrangement of subunits within a functional

GABAA receptor in the brain remains unknown in detail. Of the many subunit

combinations that are theoretically possible, only a few dozen have been shown to exist,

reflecting the differential distribution of subunit types among brain regions (Wisden et

al., 1992; Fritschy and Mohler, 1995; Pirker et al., 2000). The most abundantly

expressed receptor subtype in the brain is formed from α1, β2 and γ2 subunits (Sieghart

and Sperk, 2002; McKernan and Whiting, 1996; Whiting, 2003). The likely

stoichiometry is two α, two β and one γ subunit (Tretter et al., 1997; Farrar et al., 1999),

with the subunits arranged around the ion channel pore in the sequence γ-β-α-β-α,

(Baumann et al., 2002). Other common assemblies also contain α, β and γ2 subunits (for

example, α2β3γ2, α3β3γ2, α4βxγ2, α5β3γ2 and α6βxγ2), whereas receptors in which the γ2

subunit is replaced by γ1, γ3, or δ are less abundant. Further variability arises from the fact

that individual pentamers might contain two different α or two different β subunit

isoforms (Sieghart and Sperk, 2002). In some cases, the γ subunit can be replaced by α, ε,

α or π subunit, and the π and θ subunits might also be capable of co-assembling with α, β

and γ subunits to form receptors that contain subunits from four families (Neelands and

Macdonald, 1999; Bonnert et al., 1999). This molecular heterogeneity has important

functional consequences for GABAA receptor subtypes: subunit composition dictates not

only the properties of the receptors, but also their cell surface distribution and dynamic

regulation (Luscher and Keller, 2004; Sieghart and Sperk, 2002; Hevers and Luddens,

1998).

A combination of several methods allowed more precise subcellular localization of

GABAA receptors, and enrichment of the α1, α2, α3, α6, β2, β3 and γ2 subunits within the

postsynaptic membrane of GABAergic synapses. Each of these receptor subunits was

also found in extrasynaptic plasma membranes, and no GABAA receptor subunit type has

Chapter 1 Introduction

5

yet been found to have an exclusively synaptic location. Even in the case of α1β2γ2

GABAA receptors, which are highly enriched in synapses, more receptors are found

outside than inside synaptic junctions. Some GABAA receptors do not seem to

accumulate at synaptic junctions; for example, the δ subunit was shown to be present

exclusively in the extra - synaptic somatic and dendritic membranes of cerebellar granule

cells (Nusser et al., 1995) and at extra-synaptic and peri-synaptic locations in

hippocampal dentate gyrus granule cells (Wei et al., 2003). The lack of a γ subunit is

probably responsible for δ subunit failure to be incorporated at the synapse and δ subunit

containing receptors seems to be purely extra - synaptic. In general, receptors containing

a γ2 subunit in association with α1, α2, α3 subunits are the predominant receptor subtypes

that mediate synaptic inhibition and receptors that contain α4 or α6 subunits in

combination with δ subunits are predominantly or exclusively extra synaptic.

A vital property of a ligand-gated ion channel is its sensitivity to endogenous agonists.

For recombinant receptors that contain α, β and γ subunit, sensitivity to GABA is most

strongly affected by the type of α subunit that is present, with α3 subunits conferring the

highest and α6 subunits the lowest EC50 values (Knoflach et al., 1996; Fisher and

Macdonald, 1997; Bohme et al., 2004; 2004; Minier and Sigel, 2004). The absolute

EC50 values for specific subunit combinations reported by different groups is

considerably variable, but, in studies in which α subunits have been compared, the rank

order was shown to be α6 < α1 < α2 < α4 < α5 < α3, Bohme et al., 2004). Replacing the

γ2 subunit in α4β3γ2 assemblies with a δ subunit decreases the EC50 for GABA (Brown et

al. 2002). Overall α6β3γ2 or α6β3δ combinations have the lowest EC50s for GABA (~0.3–

0.7 µM), whereas for α1β3γ2 or α2β3γ2 subtypes they are an order of magnitude higher

(~6–14 µM).

1.1.5: Homomultimeric GABAA receptors

It is reported that some GABAA receptor subunits indeed form homomultimeric channels.

Among these subunits, the β subunit is thought to be a key component to assemble

Chapter 1 Introduction

6

heteromultimeric functional ion channels, to play a central role in determining the

subcellular locations of GABAA receptors (Connolly et al., 1996) and to bear binding

sites for agonists (Sigel et al., 1990; Amin and Weiss, 1993) and some clinically

important drugs such as general anesthetics (Cestari et al., 1996; Hill-Venning et al.,

1997). The β subunits are found to be capable of forming homomultimeric functional

channels when expressed in Xenopus oocytes or mammalian cells (Sigel et al., 1989;

Krishek et al., 1996). Channels composed of β1 subunits are constitutively active and

show spontaneous currents whereas GABAA receptors that contain β3 subunits are

inactive in the absence of GABA but they also form homomulitmeric channels, in which

the GABA current can be potentiated by pentobarbital and propofol. It was shown

(Martinez-Torres and Miledi, 2004) that the human γ2 subunit could also form

homomultimeric channels with an EC50 of 300 µM. The γ2 receptors were blocked by

bicuculline and were potentiated by pentobarbital and flunitrazepam. The other possible

homomultimeric receptors are suspected to be retained in the endoplasmic reticulum by

interactions with the Ig-binding protein BiP or calnexin and are then rapidly degraded

(Bollan et al., 2003; Gorrie et al., 1997).

1.1.6: Trafficking of GABA-receptors and interacting proteins

It is documented that GABAA receptors can be inserted and removed rapidly at synapses

(Kittler and Moss, 2003). This process is important in the synaptic inhibition and causes

the enhancement in the amplitude of miniature postsynaptic currents (mIPSC) (Wan et

al., 1997). Insulin induces the rapid insertion of GABAA receptors in to the synaptic

membrane by phosphorylating β subunits through Phosphoinositide-3 Kinase (PI3K)

(Wang et al., 2003). Conversely, removal of the receptors occurs by the activity of brain

derived neurotrophic factors (NFS) leading to suppression of mIPSC (Jovanovic et al.,

2004). Like glutamate receptors, there are both relatively immobile and highly mobile

GABAA receptors on the surface of neurons (Velazquez et al., 1989), with certain

subunits (for example, α1 and α6) being responsible for anchoring at the surface (Peran et

al., 2004). Several proteins have been identified that bind directly to GABAA to regulate

Chapter 1 Introduction

7

their trafficking and to determine the role of phosphorylation in this process (Moss and

Smart, 2001).

Fig. 1.2: Dynamic control of GABAA receptor expression at the cell surface. GABAA receptors composed of α, β and γ subunits are clustered at synaptic sites by both gephyrin-dependent and independent mechanisms, the details of which remain unresolved. GABAA receptors are intimately associated with both protein kinsaes C βII (PKC-βII) and RACK 1 (receptor associated C kinase), which together mediate the phophorylation of β receptor subunits, facilitating functional modulation of GABAA receptors. In addition, GABAA receptors at synaptic sites undergo constitutive dynamin-dependent, clathrin-medited exocytosis. This process is mediated by the interaction of receptor α β γ subunits with α and β adaptins, proteins that are essential for the recruitment of cargo in to cathrin-coated pits. Internalized receptor enters the endosomal system, where they can be returned to the cell surface. Interaction with the GABA receptor γ2 subunit might facilitate membrane interaction of GABAA receptors. This protein specifically localized into intracellular membrane including the Golgi net, Moreover GABARAP can bind NSF (N-ethylmaleimide sensitive factor), a key protein in membrane fusion that is critical for intracellular transport. Adapted from (Moss and Smart, 2001).

Chapter 1 Introduction

8

Direct binding partners include GABA-receptor-associated protein (GABARAP) (Wang

and Olsen, 2000; Wang et al., 1999), receptor for activated C-kinase (RACK1) (Brandon

et al., 1999), Src and AP2 (Mochly-Rosen and Gordon, 1998; Chang et al., 1998;

Yarwood et al., 1999). The GABA receptors are retained in the Golgi complex by

GABARAP protein (Wang et al., 1999) and their exit from this compartment could

involve interactions between GABARAP and N-ethyl maleimide sensitive factor (NSF)

and / or catalytically inactive phospholipase C (PLC)-related protein. Once inserted at

synapses, GABAA receptors are stabilized by their interaction with gephyrin and other

clustering molecules (Kneussel et al., 2001). Endocytosis of GABAA receptors might also

involve an interaction with (ubiquitin related protein), Plic-1, which could protect them

from degradation (Bedford et al., 2001).

1.1.7: Potentiation and modulation of GABAA receptors

The GABAA receptors are modulated by various chemical agents like benzodiazepines

(Sigel, 2002; Boileau and Czajkowski, 1999), neurosteroids (Rick et al., 1998),

barbiturates (Olsen et al., 1986), anesthetics (Krasowski et al., 1998) and alcohol (Mihic

et al., 1997). In total, GABAA receptors incorporate more than ten distinct binding sites

which have made this receptor a well recognized target for drug development (Korpi,

1994). In this study, it turned out that the modulatory site for propofol was the most

important one; therefore it is described in greater detail.

1.1.7.1: Modulation of GABAA receptors by Propofol

Propofol belongs to the class of general anesthetics enhancing GABAA-receptor function.

General anesthetic administration induces a state characterized by loss of consciousness,

amnesia, analgesia and immobility (Yamakura et al., 2001). At the level of ion channels,

intravenous anesthetic effects on GABAA receptors are concentration dependent. At low

concentrations, GABA-active anesthetic potentiate submaximal GABA-induced currents.

At higher concentration, they directly open channels in the absence of GABA

Chapter 1 Introduction

9

(Yamakura et al., 2001; Belleli et al., 1999). At even higher concentrations, some

anesthetics inhibit currents.

Fig. 1.3: Modulation of GABAA receptors by various chemicals. Several classes of CNS depressant drugs like benzodiazepines, barbiturates, neurosteroids, anesthetic, alcohol and propofol have been suggested to modulate GABAA receptors. In particular, the anxiolytic effects of benzodiazepines are shown to result from potentiation of GABA action on GABAA receptors. Barbiturates and related sedatives also enhance GABAA receptor-mediated inhibition, and their pharmacological spectrum overlaps with that of the benzodiazepines and related substances. The two classes of drugs have a different mechanism of action at the molecular channel level; barbiturates prolong the lifetime of GABA currents, in addition to gating channels directly at high concentrations, whereas benzodiazepines increase the frequency of opening of GABA receptor channels and do not directly open channels in the absence of GABA. Long-chain alcohols have anesthetic activity, as does ethanol at high doses (greater than 100 mM), whereas the intoxicating effects at lower concentrations (10 to 100 mM) have been suggested to involve enhancement of GABAA receptors. GABAA receptor function is further modulated by neurosteroids (Progesterone etc.). The neuroactive steroids act principally by binding directly to membrane GABAA receptors and enhancing their function in a manner resembling the barbiturates. Propofol is an alkylphenol derivative (2, 6 - diisopropylphenol), which is a fast-acting, short-lived intravenous anesthetic. It has similar mechanism of action like benzodiazepines. Adapted from (Rudolph and Antkowiak, 2004).

Chapter 1 Introduction

10

Propofol is an alkylphenol derivative (2, 6 - diisopropylphenol), which is a fast-acting,

short-lived intravenous anesthetic. The behavioral actions of propofol cover a large

concentration range. High concentration produces sleep, sedation, hypnosis and

immobility, whereas mild sedation and impairment of memory occurs at lower

concentration (around 3 % of those needed to induce immobility) (Veselis et al., 2002;

Smith et al., 1994). At sedative concentration, propofol reduces neuronal activity

prominently in cortical networks. At higher, hypnotic concentration, subcortical

structures, including the thalamus, midbrain reticular formation and possibly the

hypothalamus, are also affected (Rudolph and Antkowiak, 2004). Interestingly, there is a

linear relationship between the regional benzodiazepine binding site densities, consistent

with a similar mechanism of action of propofol and benzodiazpines (Alkire and Haier,

2001).

During propofol-induced hypnosis, global cerebral blood flow and glucose metabolism

seem to be significantly decreased, and some brain areas show a markedly higher degree

of depression than others. These regions are localized in diverse cortical areas, and also in

the thalamus and midbrain (Fiset et al., 1999; Alkire, 1998). Electroencephalography

(EEG) (Alkire, 1998) has provided evidence that thalamic structures are inhibited at

hypnotic propofol concentrations. In an elegant approach (Hofbauer et al., 2004) showed

that at mildly sedating concentration, human subjects ratings of thermal pains were

increased, and there was a corresponding increase in evoked activity in the thalamus and

somatosensory cortex. When subjects lost consciousness, noxious stimuli evoked

thalamic responses were abolished. Bonhomme used a similar experimental design,

tactile stimuli were applied during sedative and hypnotic propofol

concentration (Bonhomme et al., 2001). With hypnotic concentrations of propofol,

thalamic and cortical responses ceased. Magoun and Moruzzi found that several nuclei in

the midbrain reticular formation are involved in arousal, wakefulness and sleep and these

structures are plausible targets for general anesthetic to produce some of their sedative

and hypnotic effects (Moruzzi and Magon, 1949). General anesthesia and sleep share

some common features like depression of sensory input and motor output and similar

EEG patterns. Moreover, similar to sleep a recovery process takes place in anesthesia

Chapter 1 Introduction

11

(Tung et al., 2004). So, hypothalamic networks that are involved in sleep regulation

might have a key role in mediating anesthetic-induced hypnosis. The hypnotic effects of

several anesthetic applied to TM nucleus-hypothalamic region involved in regulation of

sleep and wakefulness, are consistent with such a mechanism (Nelson et al., 2002).

Fig. 1.4: Propofol anesthesia in humans. (a) Correlation between propofol and plasma concentration and anesthetic depth. Symbols indicate values reported in different studies to cause light and moderate sedation, hypnosis and immobility. Horizontal bars represent mean values. (b) Sites in the CNS that are thought to be involved in the sedative, hypnotic and immobilizing actions of propofol. The concentration-dependent depression of CNS functions by propofol seems to be in accordance with the classical idea that phylogenitically older parts of the CNS are more resistant to anesthetic treatment than those that appeared later in evolution. Adapted from (Rudolph and Antkowiak, 2004).

Neuroanatomical substrates that are relevant for the modulation of working memory by

general anesthetic have been identified by functional imaging studies, in which human

subject were asked to memorize words during administration of propofol (Veselis et al.,

2002). Propofol causes similar concentration-dependent depression of regional cerebral

blood flow (rCBF) and oxidative metabolism in the brain so it is reasonable to assume

that propofol induced depression in rCBF is closely linked to a depression in neuronal

activity (Kaisti et al., 2002).

Chapter 1 Introduction

12

The GABAA receptors have attracted considerable attention as a target for anesthetic

agents. Using knock-in point mutations in mice, (Jurd et al., 2003) have provided

definitive evidence that specific GABAA receptors are involved in the actions of propofol.

Whereas sites on both α and β subunits are crucial for volatile anesthetic action (Mihic et

al., 1997) for example α1-S270, α1-A291, β2-N265 and β2-M286, only the sites on β

subunits have been found to be relevant for the actions of the intravenous anesthetic

propofol (Krasowski et al., 1998). Two groups have recently reported the introduction of

point mutation in to β subunits on the GABAA receptors. Jurd showed the generation and

analysis of β3 (N265M) mice (Jurd et al., 2003). This point mutation abolished the

modulatory and direct effect of etomidate and propofol in vitro, and subsequently

reduced the modulatory actions of enflurane, whereas the modulatory actions of

neuroactive steroid alphaxalone was preserved (Siegwart et al., 2002). The duration of

the loss of the righting reflex in response to etomidate and propofol was reduced in β3

(N265M) mice compared with wild-type mice, indicating that the hypnotic activity is

mediated in part by GABAA receptors that contain the β3 subunit and in part by other

targets, possibly GABAA receptors that contain the β2 subunit. A point mutation in β1

subunit (M286W) abolished potentiation of GABA by propofol but did not alter direct

activation of the receptor by higher concentrations of propofol (Krasowski et al., 1998).

This point mutation in M3 of the β1 subunit (M286W) eliminated GABA potentiation by

1 µM propofol. In fact, submaximal GABA currents at the α2β1 (M286W) mutant

receptor were not enhanced by propofol at concentration up to 10 µM. Cysteine

substituted for these residues was used to determine whether propofol could protect it by

sulfhydryl – reactive reagents p-chloromercuribenzensulfonate (pCMBS) (Bali and

Akabas, 2004). The pCMBS reaction rate with an engineered Cys depends on two major

factors: first accessibility of the Cys to bulk solution and second reactivity of the Cys

with sulfhydrylreagents. Accessibilty depends on stearic and electrostatic factors in the

access pathway from bulk solution to the site of Cys. Their results showed that propofol

protected the substituted Cys at β2 M286W by a stearic effect caused by the local

presence of propofol and hence they concluded that this residue lies near the propofol-

binding site. The other β2 subunit residue, β2N265C was not protected by propofol.

Chapter 1 Introduction

13

Methionine oocupies a volume 43 Aº greater than asparagine and is more hydrophobic.

Thus, stearic bulk at position 265 can alter propofol binding, perhaps by inducing a

conformational change at the propofol binding site equivalent 100 Aº away.

Fig. 1.5: Proposed roles of GABAA-receptor subtypes and other target in propofol actions. Propofol acts on various targets including voltage-dependent Na+ channels to various β subunits of GABAA

receptors. Action of propofol on β2 and β3 subunits of GABAA receptors leads to sedation, hypnosis and immobility whereas its activity on glycine receptors, L-type Ca2+ channels is unclear. Adapted from (Rudolph and Antkowiak, 2004).

1.1.7.2: Modulation of GABAA receptors by further chemicals

Several classes of CNS depressant drugs had for some time been suggested to modulate

GABAA receptors. In particular, the anxiolytic effects of benzodiazepines were shown to

result from potentiation of GABA action on GABAA receptors (Costa et al., 1975). The

classical benzodiazepines such as diazepam (Valium) have had a tremendous history in

psychopharmacology primarily for clinical anxiety (Zorumski and Isenberg, 1991).

Chapter 1 Introduction

14

Other uses of benzodiazepines include sedation, muscle relaxation, and a significant

utilization for treatment of panic (Biggio et al., 1995). Barbiturates and related sedatives also

enhance GABAA receptor-mediated inhibition, and their pharmacological spectrum

overlaps with that of the benzodiazepines and related substances. The selective actions of

benzodiazepines not shown by barbiturates or vice versa arise from heterogeneity in

GABA receptor sensitivity to the drugs, and corresponding heterogeneity in brain regions

and functions. Some GABA-receptors are insensitive to benzodiazepines but not to

barbiturates. In addition, the two classes of drugs have a different mechanism of action at

the molecular channel level; barbiturates prolong the lifetime of GABA currents, in

addition to gating channels directly at high concentrations, whereas benzodiazepines

increase the frequency of opening of GABA receptor channels and do not directly open

channels in the absence of GABA (Study and Barker, 1981). Alcohols are CNS

depressants known to enhance GABAA receptor currents with a pharmacological spectrum

of action overlapping those of the benzodiazepines and barbiturates. Long-chain alcohols

have anesthetic activity, as does ethanol at high doses (greater than 100 mM), whereas the

intoxicating effects at lower concentrations (10 to 100 mM) have been suggested to

involve enhancement of GABAA receptors (Suzdak et al., 1986). GABAA receptor function

is further modulated by neurosteroids. The neurosteroids are endogenous steroid hormone

metabolites that have direct and rapid actions on cells not involving steroid hormone

receptors or regulation of gene expression. Progesterone was shown to produce rapid

sedative activity. Progesterone has anxiolytic and anticonvulsant activity; discontinuation

after long-term administration leads to withdrawal signs that are clearly CNS mediated.

The neuroactive steroids act principally by binding directly to membrane GABAA

receptors and enhancing their function in a manner resembling the barbiturates (Lambert et

al., 1995).

1.1.8: Function of distinct GABAA subunits in vivo investigated by knockout

mice

Gene targeting and transgenic mice have demonstrated several important roles for GABA in

the CNS. Knockouts of both GAD67 and GABAA receptor subunit β3 lead to early

Chapter 1 Introduction

15

neonatal lethality (Asada et al., 1997). GAD65 knockout mice show increased anxiety,

increased sensitivity to benzodiazepines (Kash et al., 1999). Epilepsy results from knockout

of GAD65, GABA β3, and GABA receptor δ subunit. Mice targeted for this subunit have

a phenotype remarkably similar to Angelman syndrome, especially the epilepsy, but also

including the cognitive, motor and sleep impairment (DeLorey et al., 1998). The γ2 subunit

knockout mice show early neonatal lethality (Gunther et al., 1995), without cleft palate,

involving impaired clustering of GABAA receptors at synapses (Essrich et al., 1998).

Because GABA receptors are important drug targets, some GABA receptors subunit

knockout mice have impaired sensitivity to drugs, such as decreased response to

benzodiazepines in γ2 homozygous knockouts. Increased response to benzodiazepines is

seen in γ2 heterozygous knockouts or in γ2L null mutants (Quinlan et al., 2000). Reduced

sensitivity to anesthetics was seen in β3 but not α6 knockouts, and reduced sensitivity to

neuroactive steroids is observed in the δ subunit knockout (Mihalek et al., 1999).

Gene targeting in mice also has been employed to ‘‘knock in’’ a mutation of the α1 subunit

H101N, which prevents benzodiazepine binding to GABA receptors containing this subunit

(McKernan et al., 2000). The resulting animals have greatly impaired sensitivity to the

sedative but not the anxiolytic actions of the benzodiazepines, whereas anticonvulsant

activity is partially reduced. This finding indicates that the subtypes of GABA receptors

containing the α1 subunit and the brain circuits in which they function are the substrates

for benzodiazepine-stimulated sedation.

α1 subunit-containing GABAA receptors in forebrain contribute to the effect of inhaled

anesthetics on conditioned fear. Knockout mice were 75 to 145 % less sensitive to the

amnestic effects of the inhaled anesthetic isoflurane. These results indicate that α1-

containing GABAA receptors in the hippocampus, amygdala, and / or cortex influence the

amnestic effects of inhaled anesthetics (Sonner et al., 2005). Also α1 knockout mice

show impaired dendritic spine maturation. There was a concomitant decreased density of

mature mushroom-shaped spines, which became more pronounced in adults. In contrast,

dendritic arborization was not altered in these mice (Heinen et al., 2003). α5 knockout

Chapter 1 Introduction

16

mice showed enhanced learning and memory and altered GABAergic synaptic

transmission (Collinson et al., 2002). In the CA1 region of hippocampal brain slices from

α5 knockout mice, the amplitude of the IPSCs was decreased, and paired-pulse

facilitation of field EPSP (fEPSP) amplitudes was enhanced indicating α5 containing

GABAA receptors play a key role in cognitive processes by controlling a component of

synaptic transmission in the CA1 region of the hippocampus.

Requirement of α5 GABAA receptors for the development of tolerance to the sedative

action of diazepam in knock-in mice, in which the α1, α2, α3, or α5 GABAA receptors had

been rendered insensitive to diazepam by histidine-arginine point (van Rijnsoever et al.,

2004). A reduction in α5 subunit-containing gamma-aminobutyric acid GABAA receptors

has been reported to enhance some forms of learning in mutant mouse models (Yee et al.,

2004). Moreover, the largely extrasynaptic α5 GABAA receptors in hippocampal

pyramidal cells are implicated as control elements of the temporal association of threat

cues in trace fear conditioning (Crestani et al., 2002).

Wild type, α2 (H101R), and α3 (H126R) mice showed a robust diminution of the motor-

depressant drug action. In contrast, α5 (H105R) mice failed to display any sedative

tolerance. α1 (H101R) mice showed no alteration of motor activity with chronic

diazepam treatment. Thus, the chronic activation of α5 GABAA receptors is crucial for the

normal development of sedative tolerance to diazepam, which manifests itself in

conjunction with α1 GABAA receptors. To identify the molecular and neuronal target

mediating the anxiolytic action of benzodiazepines, (Low et al., 2000) generated and

analyzed two mouse lines in which the α2 or α3 GABAA receptors, respectively, were

rendered insensitive to diazepam by a knock-in point mutation. The anxiolytic action of

diazepam was absent in mice with the α2 (H101R) point mutation but present in mice

with the α3 (H126R) point mutation. These findings indicate that the anxiolytic effect of

benzodiazepine drugs is mediated by α2 GABAA receptors, which are largely expressed

in the limbic system, but not by α3 GABAA receptors, which predominate in the reticular

activating system. In another study it was shown that by introducing a histidine-to-

Chapter 1 Introduction

17

arginine point mutation at position 101 of the murine alpha1-subunit gene, that α1-type

GABAA receptors, are rendered insensitive to allosteric modulation by benzodiazepine-

site ligands, whilst regulation by the physiological neurotransmitter gamma-aminobutyric

acid is preserved (Rudolph et al., 1999). Alpha1(H101R) mice failed to show the

sedative, amnesic and partly the anticonvulsant action of diazepam. In contrast, the

anxiolytic-like, myorelaxant, motor-impairing and ethanol-potentiating effects were fully

retained, and are attributed to the nonmutated GABAA receptors found in the limbic

system (α2, α5), in monoaminergic neurons (α3) and in motoneurons (α2, α5).

1.2: Histamine-receptors and the histaminergic system

1.2.1: Histamine in the nervous system

Histamine is one of the aminergic neurotransmitters, playing an important role in the

regulation of several physiological processes. Histamine is synthesized and transported to

brains of almost all animal species. The content of histamine varies between species,

being higher in lower vertebrates and to be a lower level in mammals (Reite, 1972;

Almeida and Beaven, 1981). Histamine containing nerve cells in the brain are found

exclusively in the tubomamillary nucleus of the hypothalamus (TM nucleus) and they

project throughout the brain and to all fields of hippocampus (Schwartz et al., 1991). In

all mammals, the cerebral cortex, amygdala, substantia niagra and striatum receive

moderate or dense histaminergic innervations. The density of projections in the

hippocampus and thalamus varies, and the retina and spinal cord also receive

histaminergic fibers from the TM nucleus. Also, afferent projections to TM neurons are

wide spread and come from prominent sources like infralimbic cortex, lateral septum and

preoptic nucleus (Ericson et al., 1991). The brain stem innervations in to TM nucleus

include, the adrenergic cell group C1-C3, from noradrenergic groups A1-A3, and from

serotonergic group B5-B9 also, only few fibers from locus coeruleus and the

dopaminergic groups of substantia nigra and ventral tegmentum innervates TM nucleus.

Chapter 1 Introduction

18

Fig. 1.6: Distribution of histaminergic neurons in the brain. The histamine-producing neurons, located in the tuberomamillary nucleus of the human brain, innervate all of the major parts of the cerebrum, cerebellum, posterior pituitary and the spinal cord. Adapted from (Haas and Panula, 2003).

Histamine is synthesized from histidine, which is transported in to neurons by L-amino

acid transporter. Histidine decarboxylase converts histidine in to histamine. Histamine is

then taken up in to vesicles by the vesicular monoamine – transporter VMAT-2. After

release into the synaptic cleft, histamine is methylated by histamine methyltransferase –

which is located postsynaptically and in glia – to tele-methylhistamine (t-MHA), a

metabolite that does not show any histamine like activity (Haas and Panula, 2003).

1.2.2: Metabotropic histamine receptors

Histamine is a ubiquitous chemical messenger that exerts numerous functions mediated

through at least four pharmacologically distinct receptors (H1-H4), which are all members

of the G-protein-coupled receptor family (Hill et al., 1997).

Chapter 1 Introduction

19

Fig. 1.7: Metabolism of Histamine in the neurons The L-amino-acid transporter brings histidine into neurons where histamine is synthesized by the specific enzyme histidien decarboxylase. Histamine is then taken up into vesicles by the vesicular monoamine-transporter VMAT-2. After release, histamine is methylated by histamine-methyltransferase which is located postsynaptically and in glia to tele-methylhistamine, a metabolite that does not show any histamine-like activity. Adapted from (Haas and Panula 2003).

The H1 receptor is expressed in the brain, endothelial cells, and smooth muscle cells. The

most characteristic roles for H1 receptor activation are smooth muscle contraction and

increases in vascular permeability (Ash and Schild, 1966). The H1 receptor is a 486-491

amino acid protein encoded by an intronless gene (Yamashita et al., 1991). H1 receptors

mediate excitatory actions on whole brain activity. At the cellular level excitation is

achieved by the activation of the Gq/11 heterotrimeric G-protein and its downstream

effector phospholipase C (PLC). Stimulation of the Gq/11 - PLC pathway by the H1-

receptor results in the synthesis of inositol-1, 4,5-trisphosphate and 1,2-diacylglycerol,

which in turn stimulate an increase in intracellular Ca2+ and the activation of protein

kinase C (PKC). H1 receptor activation can lead to activation of several other signaling

pathways like stimulation of nitric oxide synthase activity (via a Ca2+/calmodulin-

dependent pathway) and subsequent activation of soluble guanylyl cyclase in a variety of

different cell types (Leurs et al., 1991; Casale et al., 1985; Duncan et al., 1980).

Chapter 1 Introduction

20

The H2 receptor was first cloned from dog and later found in several species. H2 receptor

is an intronless gene and protein consists of 358-359 amino acids. The H2 receptor has

been demonstrated to function as a key modulator for gastric acid secretion, and H2

receptor antagonists are widely used for the treatment of gastrointestinal ulcers (Soll and

Walsh, 1979). The direct action on neuronal membranes is usually excitatory or

potentiates excitation. The H2 receptors signals through Gs-G-proteins, adenylyl cyclase

and PKA, which phosphorylates proteins and activates the transcription factor cyclic-

AMP response element binding protein.

The H3 receptor was first characterized as an auto-receptor - regulating histamine

synthesis and release from rat cerebral cortex, striatum, and hippocampus (Arrang et al.,

1983, 1985). H3-receptor-mediated inhibition of histamine release has also been observed

in human cerebral cortex (Arrang et al., 1988). H3 receptor is located presynaptically on

histaminergic neurons. By alternative splicing several isoform of H3 receptors, consisting

of 326-445 amino acids, are derived from a single gene. H3 also provides negative

feedback to the release of other transmitter such as glutamate, acetylcholine and

noradrenaline. H3 receptors are coupled to Gi\o and high voltage activated Ca2+ channels.

The H3 receptors are coupled negatively by cAMP and activates the mitogen activated

protein kinase pathways (Drutel et al., 2001).

The H4 receptor is detected predominantly in the periphery, for example in bone marrow

and leucocytes. The amino acid sequence of the H4 receptor has a 35 % amino acid

homology with the H3 receptor and a much lower homology to H1 and H2 receptors. Very

little is known about the actual biological function of H4 receptor. The H4 receptor can

mediate chemotaxis and calcium influx in mast calls and eosinophils (O'Reilly et al.,

2002; Hofstra et al., 2003).

Chapter 1 Introduction

21

Fig. 1.8: Signal transduction pathways mediated by various metabotropic histamine receptors. Histamine receptors and their coupling to G-proteins are shown on the left; the membrane targets of receptor activation are on the right. The box contains the signaling pathways and intracellular targets. AA, arachdonic acid; AC, adenylyl cyclase; CREB, cyclic-AMP-response element (CRE)-binding protein; DAG, diacylglycerol; GC, guanylyl cyclase, HVCC, high voltage activated Ca2+ -current; Ih, hyperploraization-activated cationic current; IAHP, the small conductance Ca2+-dependent K+ current; IK,leak, leak K+ current; Ins(1,4,5)P3, inositol-1,4,5-triphosphate; MAPK, mitogen activated protein kinase; NCX, Na+-Ca2+ exchanger; NMDA, N-methyl-d-aspartate; NO, nitric oxide; PKA, protein kinase C; PKC protein kinase C; PLA, Phospholipase A; PLC, Phospholipase C. Adapted from (Haas and Panula, 2003).

Chapter 1 Introduction

22

1.2.3: Interaction of histamine antagonists with GABAA receptors

In one report the interaction of histamine H2 receptor antagonists with GABA and

benzodiazepine binding sites in the CNS was analyzed (Lakoski et al., 1983). The

histamine H2-receptor antagonist cimetidine potently inhibited [H3] muscimol and

enhanced [H3] flunitrazepam binding in membranes prepared from several brain regions

in the rat, including the dorsal raphe nucleus. As further examined in cortical membranes,

this effect on both GABA and benzodiazepine binding sites was specific for imidazole-

derived H2 receptor antagonists (potency: cimetidine greater than metiamide greater than

tiotidine) and not observed with either several H1 receptor antagonists or histamine. Their

data indicate a striking similarity between the actions of cimetidine (and other imidazole-

derived H2 receptor antagonists) and GABA on binding parameters at the GABA receptor

complex.

In one report the in vitro antagonism of benzodizepines binding to cerebral receptors by

H1 and H2 histamine antagonists was checked (Speeg et al., 1981). They investigated

about the depressant action of histamine antagonists in CNS. It was demonstrated that

cimetidine and pyrilamine are competitive antagonists of 3H-benzodiazepine binding to

human cerebral receptor in vitro. Therefore, the interaction of antihistamine with CNS

receptors other than histamine receptor may explain, at least in part, the side effect of

sedation.

1.2.4: Ionotropic histamine receptors and direct modulatory actions of histamine to

ion channels

In another study the effect of histamine H2-receptor antagonists on the GABA-responses

of the intestine was investigated. GABA and the GABAA agonist muscimol were applied

to isolated ileal guinea pig preparations in the absence and presence of two H2 receptor

antagonists, famotidine and cimetidine. Both GABA and muscimol produced a

concentration-dependent contractile effect on the guinea pig ileum. Famotidine and

cimetidine modified this contractile effect, either by enhancing or by inhibiting it. The

Chapter 1 Introduction

23

differing results depended not only on the antagonist concentration, but also on the

concentration of GABA or muscimol. In conclusion, the interaction of H2 receptor

antagonists with GABA receptors is not limited to the central nervous system, but it also

extends to the peripheral nervous system. The receptor interaction mainly involves

GABAA receptors and depends on both the specific H2 antagonist and the concentration

used (Koutsoviti-Papadopoulou et al., 2003).

To test the hypothesis that cimetidine-like drugs produce CNS effect like seizure and

analgesia effects via inhibition of GABAA receptors, the actions of these drugs were

studied. The H2 antagonists famotidine and tiotidine produced competitive and reversible

inhibition of GABA-evoked currents in HEK 293 cells transfected. In contrast, the H2

antagonist ranitidine and the cimetidine congener improgan had very weak (if any)

effects. Authors concluded that cimetidine-like drugs do not appear to produce seizures

or analgesia by directly inhibiting GABAA receptors (Cannon et al., 2004).

In contrast to the multiple genes for metabotropic histamine receptors, no genes for

ionotropic histamine receptors have been identified in mammals up to now. There are few

hints from electrophysiological experiments that in mammals such direct activated

channels may also exist and that histamine mediate fast synaptic inhibition of rat

supraoptic oxytocin neurons via chloride conductance activation. Up to now, the ion

channels mediating this action were not identified (Hatton and Yang, 2001). At N-

methyl-D-aspartate (NMDA) receptors, histamine enhances the glutamate-evoked current

by direct binding to the channel protein itself. Histamine causes a direct facilitation of the

NMDA-receptor through its polyamine modulatory sites. When applied to cultured

hippocampal neurons, histamine selectively increased by up to tenfold the amplitude of

the component of synaptic transmission that was mediated by NMDA-receptor (Bekkers,

1993). By selectively enhancing the NMDA component of neurotransmission, histamine

should enhance process in which NMDA currents participate, such as triggering of Long-

term potentiation. Conversely, pathological conditions that deplete histamine in the brain

might lead to a reduced ability to trigger Long-term potentiation and so to memory loss.

Chapter 1 Introduction

24

In insects, histamine-acivated chloride channels were known for a long time. Native

ionotropic histamine receptors of invertebrates have been characterized in vivo,

particularly in the large monopolar neurons of the visual system of Drosophila (Hardie,

1989), the heart ganglion (Hashemzadeh-Gargari and Freschi, 1992), and the olfactory

receptor neurons of lobster (McClintock and Ache, 1989), where they mediate the pre-

synaptic inhibition of ORNs (Wachowiak et al., 2002). The ionotropic histamine

receptors mediate rapid neurotransmission in the visual system of invertebrates (Burg et

al., 1993; Hardie, 1989). Recently, genes for histamine-gated ion channels were

identified (Zheng et al., 2002; Gisselmann et al., 2002). Two histamine receptor subunits

have been so far cloned from Drosophila: HisCl-α1 (alias hisCl2, ort, hclA and Dm-

HACL1) and HisCl–α2 (alias hisCl1, ort, hclB and Dm-HACL2) (Gengs et al., 2002;

Zheng et al., 2002; Gisselmann et al., 2002). Both form homomultimeric chloride

channels when expressed in Xenopus oocytes, where HisCl-α2 is about an order of

magnitude more sensitive than HisCl-α (Zheng et al., 2002; Gisselmann et al., 2002).

1.2.5: Histamine functions and knockout mice

The histaminergic neurons are involved in many functions such as memory, sleep, and

alertness and feeding. Histaminergic neurons send widespread projections to most

cerebral regions, including those known to be important in sleep-wake control, such as

the cortex, thalamus, and posterior and preoptic / anterior hypothalamus, and to the

forebrain and brainstem aminergic and cholinergic structures (Inagaki et al., 1988; Panula

et al., 1989). In these target areas, histamine modulates neuronal activity-excitability via

H1, H2, and H3 receptors. Moreover, histaminergic neurons firing rate varies across the

sleep-wake cycle, being highest during waking and lowest during rapid-eye movement

sleep.

Administration of various substances impairing histaminergic transmission increases slow

wave sleep, whereas enhancement of transmission promotes wakefulness (Monti et al.,

1991). Muscimol-induced inactivation of the posterior hypothalamus containing

histaminergic cells results in hypersomnia in both normal and experimentally induced

Chapter 1 Introduction

25

insomniac cats (Lin, 2000). Finally, inhibition of histamine synthesis in the same area

increases slow wave sleep, whereas inhibition of histamine degradation elicits long-

lasting arousal (Lin et al., 1986, 1988). In histidine decarboxylase knockout mice,

disruption of histamine synthesis causes permanent changes in the cortical EEG and

sleep-wake cycle and that, at moments when high vigilance is required (lights off,

environmental change etc.), mice lacking brain histamine are unable to remain awake

(Parmentier et al., 2002). Neuronal histamine has been shown to suppress food intake

through activation of histamine H1 receptors in the ventromedial hypothalamus or

inhibition of the H3 receptor in the paraventricular nucleus (Sakata et al., 1988; Ookuma

et al., 1989) each of which is involved in satiety regulation.

Leptin, an ob gene product (Zhang et al., 1994) has been demonstrated to promote

histamine turnover by affecting the post - transcriptional process of histidine

decarboxylase formation or histamine release per se (Yoshimatsu et al., 1999). In

addition, concentration or turnover rate of hypothalamic histamine was lowered in leptin-

deficient ob/ob and leptin receptor-mutated db/db mice, but it was increased in diet-

induced obese animals (Yoshimatsu et al., 1999). In H1 receptor knockout mice it has

been shown that H1 - receptor is a key receptor for downstream signaling of leptin in the

brain that contributes to regulation of feeding, fat deposition, and UCP mRNA expression

(Masaki et al., 2001). Histamine also alters thermoregulation; hypothalamic

histaminergic neurons are activated not only peripherally by high ambient temperature,

but also centrally by Interleukin L-1beta as endogenous pyrogen (Kang et al., 1994). H3

receptor knockout mice display reduced locomotion and body temperature (Toyota et al.,

2002). Histamine neurons stimulate the sympathetic nervous system to increase lipolysis

in the adipose tissue (Bugajski and Janusz, 1981) an effect that depends more on H1

receptor than H2. Also, Stimulation of supraoptic nucleus by histamine causes synthesis

and release of vasopressin which in turn induces antidiuresis (Haas et al., 1975;

Armstrong and Sladek, 1985; Tuomisto et al., 1980).

Chapter 1 Introduction

26

1.2.6: Diseases where histamine is involved

Histamine is assumed to be involved in neurodegenerative disorders like in Alzheimer.

Numerous neurofibrillary tangles were found in the Alzheimer hypothalami, concentrated

in the tuberomammillary area. Most of them were of globular type and extracellular, and

only a minority were histamine immunoreactive. They may represent remnants of

degenerated TM (Nakamura, 1993). Decrease in brain histamine as well as histidine may

contribute to the cognitive decline in Alzheimer's disease directly or through the

cholinergic system (Schneider et al., 1997). The TM neurons seem morphologically

normal in patients with Parkinson disease though, the central histaminergic system

appears to be activated in Parkinson disease, and since the histaminergic innervation is

increased in the substantia nigra. Also, modulation of the histamine H3 receptor occurs in

Parkinson disease at the level of the mRNA expression in the striatum and receptor

density in the substantia nigra. Marked increase occurs in histamine H3 receptors in the

striatum and substantia nigra by tonic dopaminergic inputs (Ryu et al., 1994).

There is growing evidence to suggest the involvement of histaminergic pathways in the

pathophysiology of schizophrenia. In agreement, decreased H1 receptor-mediated

response to histamine is consistently observed among schizophrenic patients (Rauscher et

al., 1980; Nakai et al., 1991). Levels of t-MHA, the major histamine metabolite in brain

(Schwartz et al., 1971) are significantly enhanced in the cerebrospinal fluid of

schizophrenic patients (Prell et al., 1995). Finally, a polymorphism within the H2 receptor

gene was recently reported to be associated with schizophrenia (Orange et al., 1996).

Many patients diagnosed as schizophrenic have either a chronic excess or deficiency of

blood histamine. Nutritional treatment correcting these imbalances has led to great

improvement or recovery for most such patients. Histamine is used to promote alpha

wave activity in the brain, which enables an individual to handle anxiety and stress easier

(McLeod et al., 1998). If the person is deficient in histidine, it leads to a lack of histamine

and creates unbalances in calming alpha-rhythms in the brain allowing the excitatory beta

waves (responsible for the brain activity that leads to anger and tension to promote)

(McLeod et al., 1998).

Chapter 1 Introduction

27

1.2.7: Aims of the work

In Insects, the existence of histamine-gated chloride channels is long known. The

possible occurrence of such channels in vertebrates has been long postulated but no gene

was identified until now. Such channels have a fair chance to belong to the gene-family

of ligand- gated channels.

There were some indications that a so far undiscovered correlation between histamine

and GABA on the level of receptors exist. In insects, GABA and histamine gate the same

channel. In mammals, GABAA receptors are co-localized in close proximity to

histaminergic neurons, but specific interrelationship between GABA and histamine has

not been investigated yet. Therefore the aims of my work were to identify possible

candidates with bioinformatical means for histamine-gated or modulated channels in

vertebrates and to check especially members of the class of ligand-gated ion channels for

possible genes with similarity to insect histamine-gated channels.

The cDNA of found candidates should be cloned and functionally expressed in Xenopus

oocytes. The action of histamine should then be characterized by a two-electrode voltage

clamp measurements.

Chapter 2 Materials

28

Chapter 2

2: Materials

2.1: Chemicals and enzymes

Agarose LE, analytical grade, Biozym

Albumine, bovine, Fraction V, Sigma,

Alkaline Phosphatase, Shrimp, Roche

Ampicillin Disodium Salt, Sigma

ATP, Disodium Salt, Sigma

Collagenase, Worthington Biochemical Corporation

DMSO, Sigma

dNTPs, Invitrogen

DTT, Invitrogen

Diethyl Pyrocarbonate, Sigma

EDTA Disodium Salt, Sigma

Ethidium Bromide solution, Sigma

Fetal Bovine Serum, Invitrogen

Formamide, Sigma

Goat serum, Gibco, Sigma

Herring Sperm DNA, Roche

Levamisole, Sigma

Proteinase K, Roche

Restriction enzymes from: MBI Fermentas, Roche

RNase A, pancreatic, Roche

RNase H, Roche

RNasin, MBI Fermentas

RNase-free DNaseI, Roche, Biozym

Chapter 2 Materials

29

SUPERSCRIPT III RNase H- Reverse Transcriptase, Invitrogen

T4 DNA ligase, MBI Fermentas

T4 DNA polymerase, MBI Fermentas

T4 polynucleotide Kinase, MBI Fermentas

Taq DNA Polymerase, Invitrogen

All other standard chemicals were from Sigma, Fluka, Aldrich Baker, Gerbu, Merck,

Pharmacia, Promega, Riedel de Haen, Roth and Serva and used typically in p.a. quality.

2.2: Drugs used for pharmacological characterizations

Stocks solutions of drugs used for pharmacological characterizations were prepared as

indicated in the following list. If Xenopus Ringer was used as the solvent, care was taken

to check and if necessary adjust the pH to 7.4 after solving of the drugs as especially

histamine acidifies the agonist solutions.

Neurotransmitters:

Acetylcholine 100 mM Xenopus Ringer Sigma

ATP 100 mM Xenopus Ringer Sigma

Dopamine 100 mM Xenopus Ringer Sigma

GABA 1 M Xenopus Ringer Sigma

Glycine 1 M Xenopus Ringer Sigma

Glutamate 1 M Xenopus Ringer Sigma

Histamine 1 M Xenopus Ringer Sigma

Octopamine 100 mM Xenopus Ringer Sigma

Serotonin 10 mM Xenopus Ringer Sigma

GABA-receptor related drugs:

Bemegride 300 mM DMSO Acros

Chapter 2 Materials

30

Diazepam 100 mM DMSO Roche

Pentobarbital 10 mM DMSO Sigma

Propofol 50 mM DMSO Tocris

Flunitrazepam 1 mM Xenopus Ringer Ratiopharm

Histamine-receptor related drugs:

Doxylamine 10 mM DMSO RBI

DM235 100 mM DMSO Sigma

Cimetidine 10 mM Xenopus Ringer Sigma

Famotidine 10 mM Xenopus Ringer Wallgreen's

HTMT 30 mM DMSO Tocris

Histidine 100 mM Xenopus Ringer Sigma

Pyrilamine 100 mM DMSO RBI

R-alpha-Methylhistamine 100 mM Xenopus Ringer Tocris

tele-Methylhistamine 30 mM Xenopus Ringer RBI

Thioperamide 30 mM Xenopus Ringer Tocris

Others:

Harmane 100 mM DMSO Tocris

PTX 3 mM Xenopus Ringer Sigma

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2.3: Primers

All oligonucleotides (primers) were purchased from Invitrogen or MWG Biotech and solved in H2O to a concentration of 100 pmol / µl. Primers S.No. Subunit Primer Sequence

1 θ hGABA-th Not1-Stop GAC TGC GGC CGC TTA ATC GAT ATA CAT ATG GTA TAC CCA

2 θ r GABA-th –fw GCC ATC CAC ATT ACT GAC GAG CTA CAC 3 θ r GABA-th.5-new GCC GAA TTC GCC ATG GGC ATC CGA GGC ATC CGA

GGC ATG CTG 4 θ Gaba-t-Eco.ATG GAC GAA TTC CAC CAT GCT GCG AGC CGC TGA GCT

CCT 5 θ m GABA–th-3 GCA TGG GCC CTA ATA GAC ATG GTA TAA CCA 6 θ r GABA–th-rev TAT CAG GCC ATC CTG CAC ATG TGC TAC 7 θ Gaba-t-Cla1-sp-Xho1 CTC GAG CTA ATC GAT ATA CAT ATG GTA TAA CCA

GTA 8 θ mGABA-th-fw GCA TGC GGC CGC CAT CCA CAT TAC TGA TGG GCT G 9 θ r GABA –th-1000-fw GGC TCG AGC TCC TGG ATA TCA TTT TGG ATG 10 θ mGABA –th-rev GCC ATG TGA ACA CCA AGG ATC CTA GAC 11 θ rGAB-th-3-new ATC CTC GAG CCT GCT GCT GTG GTG ATA CTC 12

θ θ

mGABA-th-5 rGABA–th-3-new

GCA TGC GGC CGC CAT GGG CAT CCG AGG TAT GCT G ATC CTC GAG CCT GCT GCT GTG GTG ATA CTC

13 ε rGe-EV-r ATC ATA CTC TTG GGT CCT CTT AGA ATT CC 14 ε rGAe-up GCT GAG ATG TTG CCT AAA GTT CTC C 15 ε rG e–EV ATA ACC ATA CCC AAC CAG ATG GC 16 ε Mus-rat-gaba-e-up ACC ATG GTG CCT AAA GTT CTC CTG ATG 17 ε Mus-rat-gaba-e-down CCA GCT GGA GCC TAC AGG TTA AGG 18 ε r GA e-down TCC TGG GGA ACT GAG GTG ATT GC 19 β1 H-B1-rev GTG TAC ATA GTA AAG CCA ATA AAC 20 β1 r GABA b1-down GAG TCT AA CCG AAC CAT GAG AC 21 β1 H-B1-fw TGG ACA GTA CAA AAT CGA GAG AG 22 β1 r-B1 –fw TGG ACA GTA CAA AAT CGA GAG AGT TTG 23 β1 r-B1-rev GTG TAC ATA GTA AAG CCA ATA AAC GA 24 β2 RR-GABA – B2-up GCC ACC ATG TGG GGC TTT GCG GGA GGA AGG 25 β2 RR-GABA- B2-Do ATC AAG TGT TAA CAT AGT ACA GCC AG 26 β3 r-GABA-B3-Del Stop GTT AAC ATA GTA CAG CCA GTA AAC TAA 27 γ2 r-GABA –g2-atg-BamH1 CCG GAT CCA CCA TGA GTT CGC CAA ATA CAT GG 28 α1 r-GA-a1-4 CAA GCC CGT GAT GAA GAA AAG TCG Stop codon:

29 β3 H-GABA –b3 Not1-Stop –EV

GAC TGC GGC CGC TAG ATA TCG TTA ACA TAG TAC AGC CAG TA

30 γ2 H-GABA g2-Not 1-Stop-EV GAC TGC GGC CGT TAG ATA TCC AGA TAA AGA TAG GAG ACC CA

31 α1 GABA a1-Not1-Stop-Ev GAC TGC GGC CGC TAG ATA TCT TGA TGG GGT GTG GGG

32

γ2 H-GABAA g2- Not 1-Stop – EV

GAC TGC GGC CGC TAG ATA TCC AGA TAA AGA TAG GAG ACC CA

33 γ2 rGABA–g2-Stop-Xho1 GCC TCG AGT CAC AGA TAA AGA TAG GAG AC 34 ε rGABA-e-Not1-Stop–EV GAC TGC GGC CGC TAG ATA TCC AGG TTA AGG CAA

ATC ACC CAG TA

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32

35 ε HGABA-e-rev-O CAA GTT AAG GCA AAC AAG CCA GTA GAG CAC 36 ε HGABA–e-fw-O TTG TCG AAA GTT CTT CCA GTC TTC CTA GGC 37 ε RGABA-e-fw-O TTG CCT AA GTT CTC CTG ATG CTC CTC 38 ε RGABA-e-rev-O CAG GTT AAG GCA AAT CAC CCA GTA GAC 39 ε Rat-ep-start-H3-r CTG GAA GCT TCT TTC CAC TAG GCT GAG GCT GAG

GCC CAA AG 40 GFP 1\2 EcoRV-GFP-5` ATC GTG AGC AAG GGC GAG GAG CTG TTC ACC 41 GFP GFP down w\o stop ATC CTT GTA CAG CTC GTC CAT GCC

2.4: Standards for DNA

Following pre-made DNA size standards were used:

Gene Ruler 100 bp DNA Ladder, MBI Fermentas

Gene Ruler 1 Kbp DNA Ladder, MBI Fermentas 250 bp DNA Leiter, Diagonal

2.5: Consumption materials

Borosilicate glass capillaries (GC150TF-10) for electrophysiological studies were from

Clark Electronical Instruments. Pipette tips were from Sarstaed. Other plastic ware like

15 ml and 50 ml polypropylene tubes, 50 ml plastic tubes, 0.5-1.5 ml reaction tubes as

well as PCR-tubes, syringes and Petri dishes were bought from Sarstaed, Eppendorf or

Biozym. Cell culture dishes were from Nunc.

Distilled water was prepared in a Quarz-double distilling unit and autoclaved at 121°C

and 20 PSI for 20 minutes. Plastic ware that had to be sterile was either autoclaved under

the same conditions for 15 minutes or was used from unopened bags.

Chapter 2 Materials

33

2.6: Kits

RNA isolation:

TRIzol Reagent, Invitrogen

DNA cleanup and isolation:

QIAquick PCR Purification Kit, Qiagen,

QIAquick Nucleotide Removal Kit, Qiagen,

QIAquick Gel Extraction Kit, QiagenPlasmid DNA isolation

Qiagen plasmid maxi Kits, Qiagen

Others:

Amplicap T7 or T3 Kit, Biozyme

First strand synthesis: SuperScript III, Invitrogen

2.7: RNase free materials and chemicals

When working with RNA only RNase free solutions and vessels are used, to avoid

degradation of RNA.

For the production of RNase free water (Sambrook et al., 1989), ddH2O water was

combined with DEPC to a final concentration of 0.1 %, incubated for 30 minutes at room

temperature and then autoclaved for 15 minutes at 20 PSI and 121° C to expel DEPC.

DEPC destroys RNases, but can also kill RNA. However, when it comes in contact with

moisture, it hydrolyzes to form ethanol and carbon dioxide and is therefore rendered

harmless through appropriate treatment.

Glassware was baked over night at 180° C in an oven and solutions were made with

DEPC-water in those RNase free vessels.

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2.8: Frequently used buffers

Agar-plates: 15 g/l in LB-/NZ-Medium,

autoclaved

Alkaline Phosphatase-buffer, 10 x: 500 mM Tris-HCl; pH 8.5

50 mM MgCl2

Ampicillin stock: 10 mg/ml in H2O

Barth´s solution: 88 mM NaCl

1 mM KCl

0.82 mM MgSO4

0.33 mM Ca(NO3)2

0.41 mM CaCl2

2.4 mM NaHCO3

5 mM Tris-HCl, pH 7.4

Ethidiumbromide stock: 10 mM in H2O

HBS, 10 x: 1.4 M NaCl

0.25 M HEPES

14 mM Na2HPO4

LB (Luria-Bertani) medium; pH 7.4: 10 g/l Tryptone

5 g/l yeast extract

10 g/l NaCl

autoclaved

Ligation buffer, 5 x: 250 mM Tris-HCl; pH 7.6

50 mM MgCl2

5 mM ATP

5 mM DTT

Lysis buffer: 10 mM Tris-HCl; pH 7.9

1 mM EDTA

15 % Sucrose

2 mg/ml Lysozyme

0.2 mg/ml RNase

Chapter 2 Materials

35

0.1 mg/ml BSA

M10-Medium: 500 ml MEM

50 ml FBS

5 ml L-Glutamine, 200 mM

5 ml Pen/Strep-Soln.

ND96 99.6 mM NaCl

2 mM KCl

1 mM MgCl2

5 mM HEPES, pH 7.5

Pen/Strep solution: 10,000 U Penicilline

10 mg Streptomycine

in 1 ml 150 mM NaCl soln.

Probe buffer, 5 x: 20 % Ficoll 400, (w/v)

100 mM EDTA

0,025 % Bromephenolblue

0,025 % Xylenecyanole, (w/v)

PNK buffer: 250 mM Tris-HCl; pH 7.6

50 mM MgCl2

25 mM DTT

0,5 mM Spermidine

0,5 mM EDTA

Buffer P1: 50 mM Tris-HCl; pH 7.9

10 mM EDTA

100 µg/ml RNase A

Buffer P2: 200 mM NaOH

1 % SDS

Buffer P3: 3 M Potassium acetate; pH 7.4

Buffer QBT: 750 mM NaCl

50 mM MOPS; pH 7.0

15 % Ethanol (v/v)

0.15 % Triton X-100, (v/v)

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Buffer QC: 1 M NaCl

0.05 M MOPS; pH 7.0

15 % Ethanol

Buffer QF: 1.25 M NaCl

0.05 M Tris-HCl; pH 8.5

15 % Ethanol (v/v)

Reverse Transcriptase buffer, 5x 250 mM Tris acetate, pH 8.4

375 mM Potassium acetate

40 mM Magnesium acetate

1.-Strand buffer, 5 x: 250 mM Tris-HCl, pH 8.3

375 mM KCl

15 mM MgCl2

Taq-DNA-Polymerase buffer, 10 x: 200 mM Tris-HCl; pH 8.4

500 mM KCl

TBE: 90 mM Tris-HCl; pH 8.3

90 mM Boric acid

2 mM EDTA

TE, 1 x: 10 mM Tris-HCl; pH 7.9

1 mM EDTA

Transcription buffer: 400 mM Tris-HCl; pH 8.0

60 mM MgCl2

100 mM Dithiothreitol

20 mM Spermidine

Xenopus Ringer's soln. 115 mM NaCl

2.5 mM KCl

1.8 mM CaCl2

10 mM Hepes, pH 7.2

Chapter 2 Materials

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2.9: Bacterial strains

XL1-Blue (Stratagene) recA1 endA1 gyrA96 thi-1 hsdR17 supE44 relA1

lac [F´proAB lacIqZ∆M15 Tn10 (Tetr)]c

2.10: Plasmid vectors

General cloning and expression vectors: pSGEM: Oocyte expression vector derived from pGEMHE vector (3022 bp); (Liman et

al., 1992) Kindly provided by Dr. Michael Hollmann, Bochum.

pBluescript II KS (+) (Stratagene)

pRC/CMV (Invitrogen)

pCDNA3 (Invitrogen)

GABA-receptor expression vectors:

Following vectors originate from the German Genome Resource Center Berlin (RZPD):

GABAA-receptors:

Human delta subunit in pCMV-SPORT6 clone IRATp670E0653D6

Human alpha 1 subunit in pBluescriptR clone IRAKp961A1533Q

Human alpha 3 subunit in pBluescriptR clone IRAKp961F0547Q

Human alpha 5 subunit in pT3T7 clone IRATp970H0853D6

Human alpha 6 subunit in pSPORT6-sfi clone DKFZp686D23263Q

Human beta 3 subunit in pCMV-SPORT6 clone IRAKp961K0613Q

Human gamma 1 subunit in pBluesciptR clone IRAKp961J0448Q

Human gamma 2L subunit in pCMV-SPORT6 clone IRAKp961L0931Q

Chapter 2 Materials

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GABAC-receptors:

Human rho1 subunit in pT3T7-PacI clone IMAGp998P2111525Q

The plasmid pCDNA3.1-GABA-myc-theta containing the human GABAA-theta subunit

cDNA was a kindly gift of P. Wingrove, MSD. The plasmid pCDNA-GABA-β2

containing the rat GABAA-β2 subunit cDNA was kindly provided by R. Ruprecht,

München.

The plasmids:

pCDNA3-rGAα1 containing the rat GABA(A) alpha1-subunit cDNA in pCDNA3

pSGEM-rGAα1 containing the rat GABA(A) alpha1-subunit cDNA in pSGEM

pCDNA3-rGAβ1 containing the rat GABA(A) beta1-subunit cDNA in pCDNA3

pSGEM-rGAβ1 containing the rat GABA(A) beta1-subunit cDNA in pSGEM

pCDNA3-rGA ε containing the rat GABA(A) epsilon-subunit cDNA in pCDNA3

pSGEM-rGA ε containing the rat GABA(A) epsilon-subunit cDNA in pSGEM

were from the Lehrstuhl für Zellphysiologie plasmid collection.

2.11: Software

General sequence analysis was done with the DNASTAR program package. Alignments

of DNA sequences were done using the program Megalign. DNA sequences can be

translated into amino acid sequences with Mapdraw, which also can find restriction sites

for restriction enzymes. Protein as well as DNA Sequence comparisons to known and

published sequences were performed in the Internet (NCBI USA) with BLAST (Basic

Local Alignment Search Tool) Search, according to (Altschul et al., 1990).

Chapter 3 Methods

39

Chapter 3

3: Methods

3.1: Characterizing, isolating and concentrating nucleic acids

3.1.1: Determination of concentrations of nucleic acids

The quantification of the amount of nucleic acids in solutions was measured by

adsorption of light with a wavelength of 260 nm in a Thermo Helios Gamma photometer,

Thermo Biotech. One optical density (OD) is equivalent to a concentration of 50 µg / ml

for double stranded DNA molecules, to 40 µg / ml for RNA molecules and to 33 µg / ml

for oligonucleotides.

3.1.2: Gel electrophoresis

Following amplification of DNA by PCR (3.2.2) or linearization by restriction reactions

(3.3.1) there is need to determine, whether the appropriate product has been produced.

This is typically accomplished by agarose gel electrophoresis.

Samples were mixed with loading buffer to a final concentration of 1 x and loaded in the

wells of the gel, which was then run in a gel chamber in 1 x TBE buffer at a voltage of

approximately 10 V / cm.

The gel-running buffer was supplied with EtBr, an intercalating dye, to a final

concentration of 40 ng / ml high enough to make DNA visible under the UV-light at 302

nm, which was done with a ChemiImager 4400 (Biozym). To measure the size of a

fragment, the 1 KB DNA marker ladder was used (gene ruler, MBI-Fermentas).

Chapter 3 Methods

40

3.1.3: Phenol:chloroform extraction of nucleic acids

Phenol:chloroform extraction is used to remove protein and other contaminants from

nucleic acids and is often required to remove enzymes after an enzymatic reaction.

To one volume of DNA or RNA one volume 50:50 phenol:chloroform was added. After a

30 seconds vortexing step, the solution turned milky white. The upper aqueous phase and

the lower organic phase were separated by centrifugation for 5 minutes at 13,000 rpm in a

microcentrifuge. The upper phase was carefully removed, mixed with one volume of

chloroform and again vortexed for 30 seconds. After another centrifugation step the

supernatant was transferred to a clean tube and purification proceeded with ethanol

precipitation (3.1.4).

3.1.4: Ethanol precipitation of nucleic acids

Ethanol precipitation is a standard method used to concentrate nucleic acids. The method

relies on the tendency of nucleic acids to precipitate in a solution of salt and ethanol,

particularly at colder temperatures. Sodium acetate is the most commonly used salt, but

ammonium acetate is often used when co-precipitation of single nucleotides is to be

avoided.

To 1 volume of DNA or RNA either 0.1 volumes of 3 M sodium acetate, pH 5.5 and 2

volumes of 100 % ethanol were added. Tubes were placed at -20° C for at least 30

minutes. After a brief thaw samples were spun in a microfuge for 15 minutes at 13,000

rpm. The supernatant was carefully drawn off and discarded. After adding 70 % ethanol,

the reaction tubes were spun again at the same conditions. The pellet was dried briefly in

a SpeedVac Concentrator (SAVANT) or air-dried. The nucleic acids were then dissolved

in a desired volume of water or TE buffer.

Chapter 3 Methods

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3.1.5: QIAquick Agarose Gel Extraction Kit

DNA can be extracted from agarose gels using the QIAquick method. The DNA bands

were excised from the agarose gel and distributed in centrifuge tubes. Three volumes of

Buffer QG, a chaotropic salt, were added to the gel slice containing DNA. The tubes were

incubated at 50° C until the agarose had completely melted. The molten gel was

centrifuged through a QIAquick column (1 min, 13.000 rpm). The DNA is absorbed to

the DNA-binding resin under these conditions of high ionic strength and low pH.

Subsequently, the column was washed with 650 µl PE buffer. At the wash it was

necessary to remove all traces of PE buffer. Therefore the tubes were spun again to get

rid of all the liquid. To elute the DNA, 50 µl of EB buffer were added and the samples

were incubated for 1 minute at room temperature. The tubes were centrifuged for 30

seconds and the eluate containing the pure DNA was collected into a clean tube. All

centrifugation steps were performed at 13,000 rpm in a microfuge, which corresponds to

approximately 10,000 g.

3.1.6: QIAquick PCR-Purification Kit

This protocol was designed for clean-up PCR-generated DNA fragments or the

purification of templates for in-vitro transcription.

To the foregoing reaction 3 volumes of buffer PB were added per one volume of the

reaction sample and the mixed solution was layered onto a QIAquick column. A flow-

through was produced by centrifugation, which was discarded afterwards. The column

was washed with 750 µl of buffer PE and drained by another centrifugation step. To elute

the DNA 50 µl of buffer EB were applied, let stood for 1 minute and then centrifuged.

All centrifugation steps were performed at 13000 rpm in a microfuge, which corresponds

to approximately 10000 g.

Chapter 3 Methods

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3.1.7: RNA Extraction

RNA was isolated from mouse or rat brain tissues using Trizol (Invitrogen). 100 mg

tissue that RNA was to be isolated from was placed in glass homogenizer. Ten volumes

of Trizol reagent (1 ml) were added, and the tissues were thoroughly homogenized. The

homogenate was incubated at room temperature for 5 min before 200 µl chloroform was

added per 1 ml of homogenized tissue. The samples were shaken by hand for 15 sec and

incubated at room temperature for 3 min. The samples were centrifuged for 10 min at

13000 rpm. The top layer of the three distinct resultant layers was carefully transferred to

a new sterile tube where 500 µl isopropanol was added per ml of sample solution. This

was incubated at room temperature for 10 min and centrifuged for 10 min at 13000 rpm.

Following this centrifugation, the isopropanol was carefully removed, leaving the RNA

pellet intact. One milliliter of 70 % ethanol was then added to the tube and the samples

were centrifuged for 10 min. The ethanol was carefully removed and the RNA pellet was

allowed to air dry. Once dry, the pellet was resuspended in 50 µl DEPC treated water and

then stored at -70° C.

3.1.8: Quick preparation of plasmid DNA

For a quick and convenient preparation of plasmid DNA of multiple cultures for clone

characterization for cloning purpose the Easy-prep method was used (Berghammer and

Auer, 1993). 1.5 ml of an over night culture was centrifuged at 13,000 rpm in a table top

centrifuge, the supernatant discarded and the pellet resuspended in 50 µl lysis buffer. The

tubes were incubated for 1 min at 95° C and subsequently incubated on ice for one more

minute. The tubes were centrifuged for 20 min at 13,0000 rpm and 20 µl of the

supernatant was transferred into a fresh tube. 4 µl of the DNA solution was typically used

for analytical digestions with restriction enzymes (3.3.1). The DNA was stored at -20° C.

Chapter 3 Methods

43

3.1.9: Maxi preparation of plasmid DNA using the QIAGEN Plasmid Maxi

Kit

The QIAGEN plasmid purification is based on a modified alkaline lysis procedure,

followed by binding of plasmid DNA to QIAGEN Anion-Exchange Resin under

appropriate low salt and pH conditions. RNA, proteins, dyes and low molecular weight

impurities are removed by a medium salt wash. Plasmid DNA is eluted in a high-salt

buffer, and then concentrated and desalted by isopropanol precipitation.

Up to 500 µg of high-copy plasmid DNA from 250 ml LB cultures of E. coli can be

recovered. Bacterial cultures were spun down and the pellet resuspended in 10 ml of

buffer P1. After addition of the same amount of buffer P2, the solution was gently, but

thoroughly mixed, by inverting the tube 4-6 times. The same procedure was carried out

with 10 ml prechilled buffer P3 and the mixture incubated on ice for 5 minutes. While

centrifuging the solution at 10,000 g for 10 minutes, the necessary amount of QIAGEN-

tip 500 filter tips was equilibrated with 10 ml of buffer QBT each and the column was

allowed to empty by gravity flow. The supernatant from the centrifugation step was then

applied to the column. The QIAGEN-tip 500 was washed 2 times with 10 ml buffer QC

and the DNA eluted into a clean tube by applying 8 ml of buffer QF. The DNA was

precipitated with 0.7 volumes of room temperature isopropanol and immediately

centrifuged at 10000 g for 30 minutes in a microfuge and the supernatant carefully

decanted. The DNA was washed in 5 ml 70 % ethanol, air-dried for 5 minutes and

redissolved in 500 µl TE buffer.

3.2: PCR Methods

3.2.1: Reverse transcription The synthesis of first strand cDNA was accomplished using SUPERSCRIP III RNase H-

Reverse Transcriptase (RT), which is a DNA polymerase that synthesizes a

complementary DNA strand from single-stranded RNA. 2 µg of mouse or rat brain RNA

Chapter 3 Methods

44

(3.1.6), 100 ng of oligo dT(17-22) primer and RNase free water were mixed and heated

for 5 minutes at 70° C (total volume 10 µl). Immediately after the samples had been

chilled on ice, they were quickspun. 4 ml 5x Reverse Transcriptase buffer (Invitrogen),

10 U RNasin, 4 µl 25 mM MgCl2, 2 µl 0.1 M DTT and 1 µl 10 mM dNTPs were added at

room temperature (water up to 19 µl). The reaction was started by adding 100 U

SUPERSCRIPT III RNase H- Reverse Transcriptase. After 1-½ hours incubation at 42°

C, the reaction was stopped by adding 1 µl 0.5M EDTA, pH 8.0. The tubes were boiled

for 2 minutes at 95° C to stop the reaction.

3.2.2: Polymerase Chain Reaction

PCR (Mullis and Faloona, 1987) is used to amplify a region of DNA. The reaction

generally requires design of exact oligonucleotide primers (at least 17 bp; preferably > 20

bp) at either end of the region of interest, though there are several ways around this

necessity. Through a series of cycles of denaturation of the DNA template, annealing of

the primers and extension of DNA by a DNA polymerase, the region of DNA between

the primers is exponentially amplified. The Taq DNA polymerase is isolated from

Thermus aquaticus and catalyzes the polymerization of nucleotides into duplex DNA in

the 5’→3’ direction in the presence of magnesium. Taq exhibits furthermore a 5’→3’

exonuclease activity. Reactions contained 1x PCR buffer, 1.5 mM MgCl2, 200 µM

dNTPs, 2.5 U Taq Polymerase, combined with 0.125 U Pfu and 20-50 pmol upward

sense and downward antisense primer each. The entire reaction volume was 50-100 µl

and the fluid was overlaid with a drop of mineral oil to prevent condensation.

The polymerase chain reaction was carried out in a thermal cycler according to the

following amplification conditions:

Step 1: Denaturation 94° C, 1 minute

Step 2: hold at 80° C, enzyme added (hot-start)

Step 3: Denaturation 94° C, 1 minute

Chapter 3 Methods

45

Step 4: Annealing 50-65° C, 1 minute

Step 5: Extension 72° C, 1-3 minutes

Step 6: Final extension 72° C, 10 minutes

After 25-40 cycles were performed, temperature was held at 12° C.

The melting temperature TM of a primer molecule can be calculated in accordance to the

following formula:

TM = 2° C x number of nucleotides (A + T) + 4° C x number of nucleotides (G + C)

The annealing temperature was experimental adjusted between 50-65° C in different

applications.

3.2.3: PCR-based generation of chimeric cDNAs and site-directed mutagenesis

In order to generate chimeric cDNA, for example constructs composed of the 5'-end of

the GABACrho1 subunit and the 3' end of the GABAAβ1 subunit, the 'ligate & PCR'

method was used. The corresponding 3'- and 5'-fragments that need to be fused together

were independently amplified by PCR (3.2.2). Primers located at the site of later fusion

were phosphorylated prior to PCR (3.2.4). In this case the primer located at the 3'-end of

the 5'-fragment and the primer located at the 5'-end of the 3'-fragment. After PCR and

gel-purification of both products (3.1.2 and 3.1.5), about 100 ng of each fragment were

combined and ligated (3.3.5). 1 µl of this ligation mix was used as a template for PCR

with primers located at the 5'-end of the 5'-fragment and the 3'-end of the 3'-fragment.

These are the primers flanking the desired chimeric cDNA, in our example the GABAC ρ1

subunit 'ATG' forward primer and the GABAAβ1 subunit 'Stop' reverse primer. The PCR

products generated in this manner were gel-purified (3.1.2) and the product of the right

size isolated from the gel (3.1.5) and cloned blunt end (3.3.3) or after digestion with

suitable restriction enzymes (3.3.1) into the pSGEM vector. Site-directed mutagenesis

was performed essentially by the same method. In this case, the desired mutation was

incorporated in on of the PCR primers.

Chapter 3 Methods

46

3.2.4:Phosphorylation of PCR primers

For the PCR-mediated generation of chimeric cDNAs (3.2.3), 5'-phosphorylated PCR

primers were needed. In order to phosporylate the oligonucleotide, a 20 µl reaction mix

was set up containing 1 nmol of the corresponding oligonucleotide, 1 mM ATP, 1x PNK

buffer and 5 U of PNK enzyme. After 30 min of incubation at 37° C, the reaction was

terninated by heat (20 min 75° C). 1 µl of this reaction mix containing 50 nmol of

phosphorylated primer was directly used for PCR (3.2.2) without further purification.

3.3: Cloning of DNA

3.3.1: Restriction

Restriction enzymes recognize short, specific (often palindromic) DNA sequences. They

cleave double-stranded DNA at specific sites within or adjacent to their recognition

sequences. Each restriction enzyme has specific requirements to achieve optimal activity.

Conditions such as temperature, pH, enzyme cofactors; salt composition and ionic

strength affect enzyme activity and stability. An analytical scale restriction enzyme digest

is usually performed in a volume of 20 µl on 0.1-0.5 µg of substrate DNA (preparative

scale, 5-10 µg DNA, 20 - 50 µl), using a 2- to 10-fold excess of enzyme over DNA. For

calculation of the amount of enzyme it was assumed that app. 1 U of enzyme cuts 1 µg

DNA in a volume of 50 µl at 37° C in one hour. DNA to be restricted was incubated with

the appropriate restriction enzyme. For all restrictions 10x buffer was used.

3.3.2: Dephosphorylation

Alkaline phosphatases from both bacterial and animal sources are widely used in

molecular biology for the dephosphorylation of 5'-phosphorylated ends of DNA.

Dephosphorylation of a restriction enzyme digested plasmid (5-20 pmoles of 5' ends, 0.1-

0.5 U / pmoles 5' ends) reduces re-ligation and transformation to < 0.5 % of control

(undigested vector). Shrimp Alkaline Phosphatase is a high specific activity, heat-labile

⋅

Chapter 3 Methods

47

alkaline phosphatase for molecular biology applications isolated from Arctic shrimp.

Vectors were linearized with the appropriate restriction enzymes (3.3.1). The phosphate

rest at that 5' end of the vector was cleaved off by adding 10x SAP buffer (Roche) and 1-

5 U enzyme (SAP or CIAP) for 1 µg DNA. Incubation was done for 1 hour at 37° C.

1/20th volume of 500 mM EDTA was added and the tubes were heated for 10 minutes to

70° C to terminate the reaction, followed by purification by gel electrophoresis (3.1.2 and

3.1.5).

3.3.3: Polishing of DNA using T4 DNA polymerase and T4 PNK

To clone PCR products into a blunt end vector it is necessary to polish the DNA

molecules and to ensure the presence of a phosphate rest at their end, which can be

performed using T4 DNA polymerase and T4 PNK. T4 DNA Polymerase catalyzes the

synthesis of DNA in the 5'→3' direction. Its applications are 3' overhang removal and 5'

overhang fill-in to form blunt ends. T4 Polynucleotide Kinase catalyzes the transfer and

exchange of Pi from ATP to the 5' hydroxyl terminus of polynucleotides (double- and

single-stranded DNA). PCR samples were incubated typically in a 1x medium salt

restriction buffer (Orange buffer, MBI-Fermentas) with 1 U T4 DNA polymerase, 5 U T4

PNK, ATP and dNTPs at final concentrations of 1000 µM and 500 µM, respectively. The

samples were let stood at room temperature for 1 hour. To stop the reaction 1 µl 0.5 M

EDTA was added and the tubes were heated for 10 minutes at 70º C.

3.3.4: Fill in reaction

In some cases a filling step using the DNA Polymerase I Large (Klenow) fragment was

used to fill 5' protruding ends hence rendering blunt ends. The fill in protocol was

performed according to the product instructions. In brief 1 - 10 µg of the digested DNA

were mixed with 1 µl Klenow (5 U), 10x buffer, 40 µM of each dNTP in a final volume

of 20 µl. The reaction was incubated at room temperature for 10 minutes and stopped by

heating the mixture at 75° C for 10 minutes.

Chapter 3 Methods

48

3.3.5: Ligation of DNA

The enzyme, which is used in this application, T4 DNA ligase, catalyzes the formation of

phosphodiester bonds in the presence of ATP between double-stranded DNAs with 3'

hydroxyl and 5' phosphate termini. For the ligation reaction 100-200 ng vector was used

and 2-15 µl purified fragment (3.1.5, typically a 3 fold molar excess) was added. Both

DNA molecules were incubated in a 20 µl volume containing 2 µl 5x ligation buffer and

1 µl ligase (5 U) over night at 4-20° C.

3.3.6: Culturing of bacteria

Bacteria were grown on LB Agar plates containing 100 µg Ampicillin per ml. The

antibiotic was added, after the Agar was autoclaved at 121° C for 20 minutes. When

inoculating liquid cultures, LB contained 100 µg Ampicillin per ml.

3.3.7: Transformation of plasmid DNA

Following ligation (3.3.5), 50 µl aliquots of competent XL1-Blue bacteria were combined

with 4-20 µl of the ligation reaction in prechilled 1.5 ml plastic tubes and incubated on

ice for 30 minutes. Bacteria were heat shocked in a 42° C water bath for 45-50 seconds

and instantly placed back on ice for 2 minutes. This procedure caused the cells to become

permeable to the plasmid. Seven volumes of LB medium were added to the tube and

bacteria were incubated one hour in a 37° C incubator. The transformation reaction was

plated and the bacteria then incubated over night at 37° C. Until further use plates were

stored upside down at 4° C.

3.3.8: Sequencing of DNA

Sequencing of DNA was done using the sequencing service of the Lehrstuhl of

molekulare Neurobiochemie, Bochum. Sequencing reactions were done by the Sanger

method using fluorescine-labelled primers or oligonucleotides.

Chapter 3 Methods

49

3.4: RNA techniques

3.4.1: In vitro transcription

For the generation of templates for run-off transcripts, plasmids were linearized using

restriction enzymes (3.3.1) cutting downstream of the cDNA insert (for pSGEM-

constructs, PacI was used always) and purified using the QIAquick PCR purification kit

(3.1.6). Capped complementary RNA (cRNA) was transcribed in vitro using the

Amplicap-T7 kit (Biozym) according to the manufacturer's protocol. The resulting cRNA

was purified by ammonium acetate purification. The integrity and the quantity of the

cRNAs were investigated by agarose gel electrophoresis and visualization under UV light

(3.1.2).

Fig. 3.1: Steps of in vitro cRNA transcription.The common RNA polymerases used for in vitro transcription are SP6, T7 and T3 polymerases. Depending on the orientation of the DNA equence relative to the promoter, the template may be designed to generate a sense or antisense strand RNA (adapted from Swanson and Folander, 1992).

Chapter 3 Methods

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3.5: Functional expression of LGICs in Xenopus laevis

Xenopus laevis is one species among 14 in the Xenopus genus. They are clawed aquatic

frogs, which are found in veldt ponds and lakes in arid and semi-arid regions across

southern Africa. Xenopus laevis were purchased from Horst Kehler GmbH, Hamburg or

Nasco (USA). The frogs were housed in the animal facility of the Lehrstuhl für

Zellphysiologie in tanks containing fresh water under standard keeping conditions at

20° C.

3.5.1: Surgery

The animal was anesthetized with a 1 % tricaine solution. The surgery was performed on

ice under semi-sterile conditions. First, the animal’s skin was rinsed with sterile Barth’s

solution. A celiotomy, 0.5 to 1 cm long and approximately 2 cm above to and parallel

with the crease formed by the hind limb of the frog, gave access to the pleuroperitonial

cavity. The oocytes were carefully pulled out of the abdomen and placed into calcium-

free ND-96. Sutures of the abdominal wall and skin were performed using medical silk.

The frog was monitored in a recovery bath for 2 to 4 hours, before it was taken back to

the animal facility.

3.5.2: Oocyte preparation and injection of cRNA

The oocyte sacks were opened and all lobules cut into pieces of approximately 0.5 cm3

for better separation. Several stages of washing were carried out in a 50 ml plastic

Falcone tube, which was placed on a vertical rotator. After rotating for 15 min, the

calcium-free ND-96 solution was poured off and replaced by 30 ml fresh calcium-free

ND-96. This procedure was repeated five times.

In order to defolliculate oocytes, the oocytes were subjected to gently shaking for 2 h in

calcium-free ND-96 containing 2 mg / ml collagenase type Ia. Afterwards, the oocytes

were rinsed with several changes (3-4) of calcium-free ND-96 and rotated for 15 min. As

Chapter 3 Methods

51

a final step, the oocytes were washed with normal (Ca-containing) ND-96 solution.

The oocytes were transferred to a petri dish containing Barth's medium and selected for

size, stage and damage. The oocytes were stored in 24-well dishes in Barth's media at a

temperature of 15-18° C. Injection of cRNA (3.4.1) was performed with an electronically

controlled air pressure injector using glass pipettes. Glass capillaries were pulled using a

Kopf vertical puller. The tip was manually broken under the microscope (diameter of

about 10-20 µm) and loaded with cRNA by suction (usually 1-2 µl). Oocytes were then

placed into a 35 mm petri dish with a polypropylene mesh glued to the bottom to fix the

oocytes and injected with a given volume of cRNA. 50 nl RNA solution (app. 1 µg / µl)

was injected into an individual oocyte judged by the number of oocytes injected with 2 µl

of the RNA solution. After injection oocytes were incubated at 15-18° C in Barth´s

solution supplemented with 100 U ml-1 penicillin, 50 µg ml-1 streptomycin. Oocytes were

tested for functional expression of LGICs typically after 2–7 days.

3.5.3: Electrophysiological recording using two-electrode voltage clamp

Fig. 3.2: The two-electrode voltage clamp set-up.The Xenopus laevis oocyte is impaled with two electrodes. One electrode measures the voltage and the other injects currents.

Chapter 3 Methods

52

Two-electrode voltage-clamp recording was used to obtain current responses to applied

substances. Agonists and antagonists were diluted to the concentrations indicated with

Xenopus Ringer's solution and were applied by means of a multibarrel, single-tip

superfusion device or, for screening of multiple different solutions to one individual

oocyte by manual application of 100 µl solution by an automatic pipette. Electrodes were

pulled from borosilicate glass using a Kopf vertical pipette puller and were backfilled

with 3 M KCl. Both current and voltage electrode had resistances of 0.2-1 MΩ.

Membrane potential was controlled and current signals were recorded with a two-

electrode voltage-clamp amplifier (TURBO TEC-03, npi) and pCLAMP software (Axon

Instruments). For measuring, one oocyte was placed into the perfusion chamber and

superfused with isotonic Xenopus Ringer at a flow rate of 1-2 ml min-1. For voltage

clamp experiments, membrane potential was typically clamped at -50 mV. In case of very

high or low evoked current amplitude, the holding potential was adjusted in a range from

-30 up to -100 mV.

3.6: Functional expression of LGICs in HEK 293 cells

3.6.1: Culture of HEK 293 cells and transfection

Human embryonic kidney cell line HEK 293 were cultured in M10 medium at 37° C and

5 % CO2 in a cell culture incubator (Heareus) under standard growing conditions. One or

two days prior transfection, cells were seeded at an amount of 2-10x 104 cells into a 3.5

cm dish containing 2 ml M10 medium. Transfections were carried out using Calcium

phosphate precipitation. Cell culture medium was exchanged to 1 ml fresh medium

before starting the transfection. The transfection mix was prepared as follows: a total of

18 µg expression plasmids and 2 µg pIRES-eGFP (Clontech) was solved in 225 µl ddH20

and mixed with 25 µl 2.5 M CaCl2. The solution was carefully overlaid with drop-wise

addition of 250 µl 2x HBS, mixed by up and down pipetting and incubated for 15 min.

100 µl of the transfection mix was used for each dish. After 4-6 h, dished were washed

with PBS twice and fresh M10 medium was added. Electrophysiological experiments

Chapter 3 Methods

53

were performed 1-3 day after transfection. Efficiency of transfection, typically 10-50 %,

was checked by GFP-fluorescence detection of the GFP coded by the pIRES-eGFP.

3.6.2: Patch clamp investigation of GABA receptors expressed in HEK293 cells

Patch clamp technique is an important method for studying electrophysiological

properties of biological membranes, which was developed by Neher and Sakmann. It

allows the recording of macroscopic whole-cell currents flowing across biomembranes

through ion channels. One can study membrane-contained ion channels, receptor

activated channels and second-messenger activated channels. The electrical circuit used

in the patch clamp technique is shown in the following figure.

Fig. 3.3: Schematic representation of patch clamp circuit. The diagram illustrates the circuits in the patch clamp technique. The patch pipette is in direct contact with the cellular membrane to form a seal. The current flowing through the ion channels on the membrane enters the patch pipette through an electrode. With pipette potential of Vp, the current through it (Ip) is then fed to a current-voltage converter (head stage amplifier), which has a feedback resistor (R).

Mammalian culture cells such as HEK 293 are frequently employed for studying

recombinant expression of individual receptors. A vast amount of publications exist

where HEK 293 cells have been used for investigation of GABAA receptors. Cultured

mammalian cells have the advantage of providing an environment that is similar to the

Chapter 3 Methods

54

milieu of the natural mammalian cells. To a great degree, channel permeability, post-

translational processing, signaling and coupling to other cellular factors in these cells are

similar to the processes in most mammalian cells. Post-translational processing like

protein glycosylation varies in other expression systems like Xenopus oocytes. This could

probably interfere with the proper assembly of subunits.

Transfected HEK 293 cells (3.6.1) transiently expressing green fluorescent protein and

the recombinant GABA channels were recorded in the whole-cell voltage-clamp

configuration (Hamill et al., 1981) under visual control using an inverted microscope

(Zeiss, Jena, Germany). The cells were kept in an external solution containing: 138 mM

NaCl, 5 mM KCl, 0.5 mM CaCl2, 1.5 mM MgCl2, 10 mM Glucose and 10 mM HEPES.

pH was adjusted to 7.4 with NaOH. Patch electrodes were pulled from borosilicate glass

(Clark Electromedical Instruments) using a horizontal pipette puller (DMZ Universal

Puller, Zeitz-Instruments) to yield pipettes with resistances of 3 to 6 MΩ. Pipettes were

filled with a solution containing: 140 mM KCl, 0.1 mM CaCl2, 1 mM MgCl2, 5 mM

EGTA, 10 mM HEPES adjusted to 7.3 with KOH. After getting the whole cell mode, the

cell was lifted and it was positioned in front of a perfusion system consisting of a theta

tube flow pipe. Voltage protocols were delivered and current signals were recorded with

an EPC-7 amplifier using the pCLAMP software on a 486-IBM compatible PC running

under DOS. Typical holding potential was -60 mV. Data were collected and analyzed

with pCLAMP 7 software (Axon Instruments) in combination with SigmaPlot 7.0 (SPSS,

Inc).

Chapter 4 Results I

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Chapter 4 4.1: Bioinformatical search for histamine-gated channels

For histamine, one of the aminergic neurotransmitters, molecular biology techniques lead

to the identification of various metabotropic histamine receptors (H1 - H4), which are all

members of the G-protein-coupled receptor family (Hill et al., 1997). There are some

indications from electrophysiological studies, that there might be an additional histamine-

gated ion channel in vertebrates (Hatton and Yang, 2001) but no genes were identified so

far. Histamine was pointed out to be the major neurotransmitter in the invertebrate retina

(Hardie, 1989) but only recently the genes HisCl-α1 and HisCl-α2 for histamine-gated ion

channels were identified (Zheng et al., 2002; Gisselmann et al., 2002) in Drosophila

melanogaster. The discovery that invertebrate histamine-gated chloride channels belong

to the super family of ligand-gated ion channels gave raise to speculations about the

nature of potential histamine-gated channels in vertebrates. Identification of histamine-

gated ion channels in vertebrate, may lead to several physiological and pharmacological

important development in histamine research field.

We started using bioinformatics approaches, to find potential candidates of histamine-

gated ion channel genes or cDNAs in vertebrates. We looked for the sequence homology

for HisCl-α1 and HisCl-α2 in various vertebrates genomes (e.g. Zebra fish, rat, mouse,

human etc). For this, we scanned genomes with the complete coding regions of HisCl- α1

and HisCl-α2 receptor subunits as well as separate scanning for all of the predicted

features characteristics of ligand-gated ion channel subunits: signal peptide, two cysteines

separated by 13 amino acids (cysteine-bridge) and four putative transmembrane (TM1 to

TM4) domains. All these parameters did not show any sequence homology for any

unknown ligand-gated ion channel. However, it exhibited some similarities with known

GABAA receptors subunits as well as some extent to the glycine receptors. We also

Chapter 4 Results I

56

looked for any sequence homology in the EST database, even for incomplete vertebrate

genomes (e.g. Chimpanze and Xenopus laevis).

4.2: Construction of expression vectors for GABAA receptors

As our genome-scanning results showed that GABAA receptors have some similarities

with histamine-gated ion channels; we decided to investigate the action of histamine to

the class of ionotropic GABAA receptors. It was planned that the electrophysiological

characterization to be performed in Xenopus oocytes with two electrode-voltage clamping

(3.5.3). To generate the corresponding cRNA, it was necessary to clone the cDNA for the

GABA receptors to be investigated into the pSGEM vector as this vector allows to in

vitro transcribe cRNA suitable for an effective high level expression. Different subunits

of GABAA receptors from rat (α1, β1, θ, ε), human (α3, α5, β3) and human ρ1 GABAC

were cloned into pSGEM. The constructs pSGEM-mouse γ2 as well as pSGEM-rat β2 and

pSGEM-δ were already available from other projects of our department.

For cloning of pSGEM-α1, -α5, -β1, -β3 and -ε the open reading frame of the cDNA insert

present in various plasmids used as template (see table 1) was amplified by PCR (3.2.2)

using the primers indicated in table 1. The PCR products were gel purified (3.1.2), made

blunt-end and phosphorylated for cloning into EcoRV/SmaI cut pSGEM (3.3.1) or were

cut with suitable restriction sites incorporated into the PCR primers (2.3) and cloned into

the corresponding polylinker sites of the dephosphorylated pSGEM vector (3.3.2). After

ligation (3.3.5) and transformation into E. coli XL1-blue (3.3.7), plasmids isolated from

several colonies (3.1.8) were analyzed by restriction mapping (3.3.1) and from clones

with the correct insert, large amounts of plasmid was prepared by the maxi prep method

(3.1.9). Fig. 4.1 shows the schematic representations of the cloning of α1, β1 and α5

subunits of GABAA receptors. The identity and integrity of the cDNA insert was

confirmed by sequencing (3.3.8) typically using primers flanking the polylinker site (T7

and pSGEM-down or SP6). The final plasmids were named according to table 1.

Chapter 4 Results I

57

Table 1: List of plasmids containing GABAA receptors cDNAs used and templates for PCR as well

For construction of an expression vector for the rat θ subunit, no cloned cDNA was

available. Therefore it was decided to amplify the cDNA by RT-PCR. As a template for

this PCR, rat cDNA was used generated by reverse transcription (3.2.1) from total RNA

isolated from rat brain (3.2.2). As it turned out that it was impossible to amplify the

complete open reading frame of the theta subunit for cloning in one piece, the open

reading frame was amplified in two pieces using the primer pairs theta-ATG, theta-5`up

for the 5'-fragment and theta-stop, theta-3`down for the 3'-fragment (3.2.2). Cloning of

the complete open reading frame was achieved by first cloning of 5' fragment into the

NotI and EcoRI site of pSGEM (3.3.1 - 3.3.7). Into the resulting plasmid, the 3'-fragment

was inserted between the EcoRV and ApaI sites. The final plasmid pSGEMrat-θ

contained the complete open reading frame cloned into the NotI and ApaI sites of the

pSGEM polylinker. The singularly site EcoRV in the theta cDNA was used for fusing

together both fragments. The final plasmid was verified by sequencing (3.3.8) with

primers flanking the polylinker (T7, pSGEM-down) and primer theta-seq-1 sequencing

the fusion site of the two fragments.

Chapter 4 Results I

58

Fig. 4.1: Schematic representation: Cloning of α1 subunit of GABAA receptors. For cloning of pSGEM-α1 the open reading frame of the α1-cDNA insert present in pCDNA3 plasmid used as template was amplified by PCR using specific oligonucleotides. A PCR fragment of 1500 bp of α1 subunit was generated. The pSGEM plasmid was digested by restriction Enzyme BamHI and Xho1 and PCR fragment by BamHI and XhoI and cloned into the corresponding polylinker sites of the dephosphorylated pSGEM vector.

Chapter 4 Results I

59

Fig. 4.2: Gel photographs showing molecular cloning of β1 subunit of GABAA receptors. (A) PCR amplification of β1 subunit from cDNA of rat. Columns (1, 2 and 3) show the PCR amplified bands of β1 subunit. (B) Restriction analysis of the β1 subunit insert cloned in pSGEM vector. Columns (1, 2 and 4) show the positive clones whereas (3) is the negative clone.

Chapter 4 Results I

60

Fig. 4.3: Gel photographs showing molecular cloning of α5 subunit of GABAA receptors. (A) PCR amplification of α5 subunit from cDNA of rat. Columns (1, 2, and 3) show the PCR amplified bands of α5 subunit. (B) Restriction analysis of the α5 subunit insert cloned in pSGEM vector. Column (4) shows the positive clone whereas (1, 2, and 3) are negative clones.

Chapter 4 Results I

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4.3: Establishing functional expression GABAA receptors in Xenopus oocytes

In the first electrophysiological part of this work it had to be shown that GABAA

receptors could be successfully expressed in Xenopus oocytes using our expression

vectors. Further it was intended to test, whether the basic properties of the expressed

GABA receptors, like the affinity for GABA match the data known from literature.

To functionally characterize the GABAA subunits, various combinations of their cRNA

generated by in vitro transcription (3.4.1) were injected into Xenopus oocytes (3.5.2) and

measured using two-electrode voltage clamp technique (3.5.3). Application of GABA to

oocytes injected with cRNA of various subunit combinations elicited inward membrane

currents (typically measured at -60 mV), which are due to the opening of membrane

channels permeable mainly to chloride ions. Dose-response curves were generated by

application of various concentrations of GABA.

Of all the β subunit-containing GABAA receptors in the brain, only 19-25 % contains the

β3 subunit, with 55-60 % containing the β2 subunit and 16-18 % containing the β1 subunit

(Benke et al., 1994). We checked the effect of various β subunits for GABA affinity

when combined with the α1 subunit. Fig. 4.5A shows the dose-response curves for α1β1,

α1β2 and α1β3. GABA activated these heteromultimeric receptors with an EC50 of 8.14 ±

0.03 µM, (n = 9), 3.9 ± 0.08 µM, (n = 5) and 1.03 ± 0.15 µM, (n = 5) respectively.

Incorporation of β3 subunit in to the combination leads to a higher reduction in the EC50

of GABA (Fig. 4.4A).

There are 6 different subtypes of α subunit of GABAA receptors. α and β are described

as having binding site for GABA and in conjunction with γ2 subunit α forms binding site

for benzodiazepines (Minier and Siegel, 2004; Bohme et al., 2004; Gunther et al., 1995;

Walters et al., 2000). Fig. 4.4C shows dose-response curve and bar-diagram for α3β1 and

α5β1. GABA evoked EC50 for these receptors are 1.28 ± 0.07 µM, (n = 3) and 3.7µM ±

0.11, (n = 5); respectively. α3β1 seems to be most sensitive GABA combination when

keeping β1 as a constant subunit.

Chapter 4 Results I

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Fig. 4.4: Concentration-response curves and bar diagrams of GABA for different GABAA receptors. (A and C) Concentration-response and bar-diagram curves for α1β1, α1β2, α1β3. Incorporation of β subunit in to the combination leads to a higher reduction in the EC50 of GABA the order of sensitivity for GABA is α1β3 > α5β1 > α1β1. (B and D) Concentration-response and bar-diagram for α3β1and α5β1 respectively. α3β1 seems to be more sensitive GABA combination than α5β1, when keeping β1 as a constant subunit. (E and F) Concentration-response and bar-diagram for α1β2γ2 and GABAC ρ respectively. ρ1 receptors are much more sensitive for GABA than α1β2γ2.

Chapter 4 Results I

63

The major receptor subtype of the GABAA receptor in the brain most probably consists of

α1, β2 and γ2 subunits. The most likely stoichiometry is two α subunits, two β subunits

and one γ subunit. α1β2γ2 is the most prominent GABAA receptor combination in the

CNS, contributing 70 % of the total GABAA receptors (Mehta and Ticku 1999). The ρ1 is

the GABAC receptors and is insensitive for commonly known modulators like

neurosteroids and barbiturates. Fig. 4.4E shows the dose response curve for α1β2γ2 and ρ1

where GABA activates these receptors with an EC50 of 17.2 ± 0.1 µM, (n = 5) and 0.57 ±

0.01 µM, (n = 5), respectively.

Table 2 shows the EC50 of various GABAA receptor subunit combinations, checked in

this study. α1β3 is the most sensitive subunit combination we studied. The order of

sensitivity for GABA is α1β3 > α3β1 > α1β2 > α5β1 > α1β1> α1β2γ2.

S.No. Subunit Combinations

EC50 (µM)

1. α1β3 1.03 ± 0.15 2. α3β1 1.28 ± 0.07 3. α1β2 3.19 ± 0.08 4. α5β1 3.70 ± 0.11 5. α1β1 8.14 ± 0.03 6. α1β2γ2 17.2 ± 0.10

Table 2: EC50 of various tested GABAA receptor subunit combinations.

For the combination α1β1, an even more detailed characterization was performed. The

reversal potential -25 mV of the GABA evoked current determined by I/V curve fits the

fact that GABA evoked a chloride current (data not shown). Also the basic

pharmacological properties of GABAA receptors like block by PTX and bicuculline were

experimentally confirmed (data not shown). To characterize the potentiation properties of

α1β1, we co-applied 2µM GABA with various concentrations of pentobarbital and

Chapter 4 Results I

64

propofol and found that GABA current was potentiated in a similar manner as described

in previous studies (Parker. et al., 1986), (Fig. 4.5).

Fig. 4.5: Concentration-response curves of propofol (closed circles) and pentobarbital (open circles) potentitated GABA-current on α1β1 receptors. Data represent the mean ± S.E.M. and curve is derived from the fit of the Hill equation to the data taken from 3 oocytes.

Our determined data match essentially the results of other groups (Parker et al., 1986;

Harris et al., 1995; Longoni et al., 1992; Williams et al., 2002) proving that our

expression system is well suited to functionally express GABAA and GABAC receptors.

Chapter 4 Results I

65

4.4: Direct effects of histamine on heteromultimeric GABAA receptors

Next we examined the effect of histamine on GABAA receptors. We checked α1β1, α1β2,

α1β3 and α5β1 GABAA receptor subunit combinations. In most tested oocytes, application

of histamine up to 10 mM concentration did not evoke any response or responses below 1

% of the maximal current evoked by saturating concentrations of GABAA (100 µM - 1

mM). These experiments show that under normal conditions, GABAA receptors are not

activated by histamine. In few tested GABAA expressing oocytes, histamine evoked

small currents especially in the subunit combination α1β2γ2. These few special unusual

cases are described later (Chapter 7) in the results section.

4.5: Modulation of heteromultimeric GABAA receptors by histamine GABAA receptors are modulated by various chemical agents like barbiturates (Sigel,

2002; Boileau and Czajkowski, 1999), neurosteroids (Rick et al., 1998) and anesthetic.

(Krasowski et al., 1998) At lower concentration they show no activity on GABAA

receptor, though co-application of them with GABA modulates GABA-evoked current

(Yamakura et al., 2001; Belelli et al., 1999). Therefore, we decided to check the effect of

co-application of histamine with GABA on GABAA receptors.

4.5.1: Potentiation of α1β1 GABAA receptors by histamine Application of GABA to oocytes injected with rat α1β1 subunits cRNA elicited inward

membrane currents (at -60 mV), which are due to the opening of membrane channels

permeable mainly to chloride ions. The amplitude of the responses evoked by 2 µM

GABA in injected oocytes was greatly potentiated when GABA was applied together

with histamine. Control oocytes showed no response to histamine. Fig. 4.6A shows

potentiation of GABA responses by different concentration of histamine. Measurements

were made by exposing oocyte to increasing concentration of histamine (1 µM - 30 mM)

at EC10 GABA (2 µM). The onset of this potentiating effect was rapid. For example, the

oocyte illustrated in Fig. 4.6A showed a maximal potentiation with in 10 sec of adding

Chapter 4 Results I

66

histamine and recovered to the control value within a similar time after washing out the

histamine. Histamine potentiates α1β1 with an EC50 of 0.49 mM, (n = 7).

(Huang and Dillon, 1999) indicated that GABAergic signaling in the CNS may be

significantly altered during conditions that increase or decrease pH. Histamine is a

polyamine so there is a possibility that prolong standing at room temperature may alter its

pH, too. To avoid such situation we measured the pH (7.4) of histamine solution before

and after the experiment and did not find any alteration in the pH of histamine solution

that ruled out the possibility that potentiation by histamine might be just because of

alteration in pH.

To further characterize the nature of potentiation with GABA we checked potentiation

mediated by histamine at various concentration of GABA. Fig. 4.6B and C shows the

histamine potentiated GABA-evoked current at EC50 (10 µM GABA) and EC90 GABA (1

mM), respectively. The variation in the GABA concentration affects the degree of

potentiation. At EC50 of GABA, the histamine potentiated GABA-evoked current with an

EC50 of 710 ± 60 µM, (n = 7). In case of EC90 of GABA, we found that potentiation

mediated by histamine decreased remarkably. α1β1 receptors were potentiated with much

higher EC50 of histamine than the EC50 calculated for it at EC10 and EC50 of GABA. The

EC50 of histamine at EC90 of GABA is 1.9 ± 0.2 mM, (n = 4).

We next inquired whether evoked potentiation was histamine specific or any other

neurotransmitter or substances can modulate GABAA receptors too. We checked large

variety of known neurotransmitters as well as other substances at concentration up to 1

mM and never found any potentiation of the current evoked by 2 µM GABA (Table 3).

Finally, to check that potentiation was not due to the polyamine nature of histamine, we

checked spermidine, a biogenic polyamine and found no change in GABA evoked

currents. Therefore, we confirmed that GABAA receptors are specifically modulated by

histamine.

Chapter 4 Results I

67

Fig. 4.6: Potentiation of α1β1 receptors by histamine. (A, B and C) The current traces to different concentration of histamine (0.1-10 mM) in the presence of EC10 (2 µM), EC50 (10 µM) and EC90 (1 mM) of GABA respectively. (D) Concentration-response curves of histamine potentiated GABA-current for EC10 (triangle), EC50 (open circles), EC90 (closed circles) of GABA. Data represent the mean ± S.E.M. of at least three independent experiments and curve was fitted using Hill equation.

Chapter 4 Results I

68

Table 3: Various chemicals checked on GABAA receptors.

We investigated if the co-application of histamine with GABA alters the permeability of

the receptor to ions. Therefore, we injected α1β1 subunits cRNAs into Xenopus oocytes.

The membrane current activated by GABA decreased in amplitude as the oocyte was

depolarized and inverted direction at about –25 mV (Fig. 4.7), which corresponds to the

chloride equilibrium potential in Xenopus oocytes under experimental conditions (Barish

Chapter 4 Results I

69

et al., 1983; Kusano et al., 1982). As the potential was made more positive, the peak

GABA current increased to a maximum up to -30 mV. (This rectification at positive

potential is similar to that seen with chloride currents activated by GABA (Gundersen et

al., 1984). It was similar to reversal potential for homo and hetromultimeric GABAA

receptors. Various groups have showed that reversal potential for homomultimeric β2 and

β3 is close to the chloride equilibrium potential (-23 mV) when expressed in Xenopus

oocytes (Ymer et al., 1989; Dascal et al., 1984).

Fig. 4.7: Current-voltage relationship of the GABA-evoked current potentiated by histamine in oocyte injected with α1β1 subunits cRNAs. Square = 2 µM GABA Circle = 2 µM GABA + 3 mM Histamine 4.6.1: Potentiation of α1β1 GABAA receptors by histidine We investigated that if the other metabolite of histamine can potentiate the GABAA

receptor, for this histidine was the ideal candidate. Histamine is synthesized from

Chapter 4 Results I

70

histidine, which is transported in to neurons by L-amino acid transporter. Histidine

decarboxylase converts histidine to histamine (Haas and Panula, 2003). (Fig. 4.8) shows

potentiation of GABA-evoked membrane currents by histidine. Records are from oocyte

injected with α1β1 subunit cRNA. Oocytes were voltage clamped at -60 mV and the

traces show clamp currents. Inward membrane currents correspond to downward

deflection in this figure. Fig. 4.8A shows response evoked by GABA EC10 (2 µM)

combined with the increasing concentration of histidine (1 µM - 1 mM). Fig. 4.8D shows

dose-response curve for histidine potentiated GABA-evoked currents. Histidine

potentiated GABA-evoked current with an EC50 of 168 ± 14 µM. Data were obtained

from 7 oocytes and are normalized as a fraction of the response in each oocyte to 2 µM

GABA alone. The curve was fitted by Hill equation. In the 3 oocytes, where the

maximum response to both GABA (2 µM) and histidine (1 mM) were determined, the

maximal current evoked by histidine plus GABA was 2-2.5 folds of that evoked by

GABA.

4.6.2: Characterization of histidine potentiation

To characterize the nature of potentiation with GABA we checked potentiation mediated

by histidine at various concentration of GABA. Fig. 4.8B and C shows the histidine

potentiated GABA-evoked current at EC50 and EC90 GABA concentrations, respectively.

We found that the variation in the GABA concentration affects the degree of potentiation.

At 10 µM GABA (EC50), histidine potentiated GABA-evoked current with an EC50 of

78.7 µM, (n = 5). In case of higher concentrations of GABA (1 mM, EC90) we found

quite it interesting that histidine potentited α1β1 receptors with much lower EC50 for

histidine than calculated for it at lower EC10 and EC50 GABA concentrations. The EC50

for histidine at 1 mM of GABA (EC90) is 32.4 ± 5 µM, (n = 5) in all cases, in contrast to

histamine, histidine potentiation induced higher robust inward currents.

Fig. 4.8D shows the dose-response relationship of histidine potentiated GABA-evoked

currents. Here, at EC90 GABA, histidine shows much lower EC50 for potentiation than

EC10. Moreover, at EC90 α1β1 receptors were nearly 10 folds more sensitive to the

histidine.

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Fig. 4.8: Potentiation of α1β1 receptors by histidine. (A, B and C) The current traces to different concentration of histidine (1-3000 µM) in the presence of EC10 (2 µM), EC50 (10 µM) and EC90 (1 mM) of GABA respectively. (D) Concentration-response curves of histidine potentiated GABA-current for EC10 (closed circles), EC50 (open circles), EC90 (triangles) of GABA. Data represent the mean ± S.E.M. of at least three independent experiments and curve was fitted using Hill equation.

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4.6.3: Dependence of average histamine and histidine potentiation on GABA concentration Fig. 4.9B shows, the fractional potentiation mediated by histidine. Here the fractional

potentiation was obtained by dividing the effective potentiation evoked by histidine (1

mM) with GABA-evoked current (EC10, EC50, EC90 GABA respectively). At EC90

GABA, the average fractional potentiation of histidine was 10 ± 0.2, (n = 13) folds which

decreases to 6 ± 0.8, (n = 7) & 2.5 ± 0.1, (n = 5) folds at EC50 and EC10 GABA,

respectively. The investigation was performed on different populations of oocytes for the

three distinct GABA concentrations. The action of histamine at different GABA

concentrations on the same individual oocyte is described in Fig. 4.9A.The fractional

potentiation decreased from 5 to 2.5 folds on increasing GABA concentration from 2 µM

-1 mM.

Fig. 4.9: Average fractional potentiation of histamine and histidine depends up on GABA concentration. (A) Average fractional potentiation of GABA currents by histamine at EC10, EC50, and EC90 GABA for α1β1 receptors. The fractional potential is nearly 5 folds at EC10 GABA, which decreases to 2-2.5 folds at EC90 GABA. (B) Average fractional potentiation of GABA currents by histidine at EC10, EC50, and EC90 GABA for α1β1 receptors. In case of histidine at higher concentration of GABA (EC90

GABA), fractional potentiation mediated by α1β1 receptors is nearly 10 folds, which decrease to nearly 2 folds at EC10 GABA. Here the fractional potentiation was obtained by dividing the effective potentiation evoked by histamine \ histidine with GABA-evoked current.

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4.6.4: Histidine does not alter the I/V curve and retains selectivity for the permeability of the channel We also checked the permeability of ions on α1β1 receptors injected in Xenopus oocytes

by co-application of histidine with GABA at various holding potentials. 2 µM GABA

was co-applied with constant 1mM histidine. Like typical chloride channel reversal

potential in oocytes, here histidine potentiated GABA-evoked current reversed at nearly -

20 mV. Fig. 4.10 shows the current-voltage relationship for histidine potentiated GABA-

evoked current. Like GABA, histidine shows rectification at positive potentials. The

rectification properties were not altered by histidine potentiation so that this potentiation

cannot be explained by influencing the current-voltage relationship.

Fig. 4.10: The current-voltage relationship comparison of current evoked by GABA or GABA with histidine in an oocyte injectd with α1β1 subunit cRNA. Circle = 2 µM GABA Square = 2 µM GABA + 1 mM histidine

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4.7: Potentiation of GABAA receptors in HEK 293 cells It is a common fact that the choice of the recombinant expression system can severely

influence the properties of the expressed channels. Therefore, to check whether the

potentiation observed in Xenopus oocytes can be repeated in other cell culture system,

various combinations of GABAA receptors were expressed in HEK 293 cells (3.6.1) and

investigated by patch clamp measurements (3.6.2). To achieve a functional expression of

the α5β1 receptor, the plasmids pCDNA3-rat-G-α5 and pCDNA3-rat-G-β1 were co-

transfected in a 1:1 ratio in HEK 293 cells (3.6.1). The potentiation of histamine–

potentiated currents was determined using brief application of 2 µM GABA alone and 1

mM histamine in combination with 2 µM GABA in whole cell patch-clamp mode.

Potentiation of α5β1 GABAA receptors with histamine produced a substantial increase of

2.5 ± 0.2, (n = 8) folds in the amplitude of currents activated with 2 µM GABA.

Experiments performed with α1β1 receptors in cooperation with Angela Vogt-Eisele and

Katja Erlkamp of our department confirmed these findings. They found that also in α1β1

subunit combinations 1 mM histamine potentiated the current evoked by 2 µM GABA up

to 5 folds (Fig. 4.11). In later experiments with GABAA receptors in a primary culture of

native neurons from the olfactory bulb, they also confirmed that histamine and histidine

potentiated the GABA evoked currents. A proof that the potentiation also occurs in native

neurons and is not limited to artificial recombinant expression systems.

Therefore, my own patch-clamp measurements and those performed in cooperation with

other groups of our department confirm that the potentiation of GABAA receptors

observed in Xenopus oocytes can be repeated in other cell systems, too.

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Fig. 4.11: Potentiation of GABAA receptors by histamine in HEK 293 cells. (A and B) Show the histamine-mediated potentiation in α5β1 (A) and α1β2γ2 (B) subunit combinations, respectively. α5β1 exhibits higher potentiation than α1β2γ2 receptors.

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4.8: Dependence of histamine potentiation on GABAA receptor subunit combinations

We started to investigate two major questions, (1) if substitution of various subunits

affects the affinity for the potentiation mediated by histamine and (2) which subunit has

the binding site for histamine. For this we first of all replaced β subunit keeping the α1

subunits constant. Fig. 4.12A and B shows the current traces for α1β2 and α1β3 subunit

combinations expressed in oocytes respectively. At a concentration of 2 µM GABA,

histamine potentiates α1β2 and α1β3 subunit combinations with an EC50 of 2.15 ± 0.16

mM and 335 ± 0.02 µM, (n = 3) respectively. α1β3 was more sensitive for histamine and

in this subunit combination; the potentiation starts at 100µM histamine whereas α1β2 was

significantly less sensitive though it is potentiated by histamine. The average amplitude

of histamine-mediated potentiation in α1β2 is 2 folds, whereas α1β3 shows more

pronounced potentiation with amplitude of 5 folds. Based on the EC50 values of various

beta subunit combinations, the order of sensitivity for histamine is α1β3 > α1β1 > α1β2.

Therefore, from these experiments we were able to show that affinity for potentiation by

histamine depends up on the subunit combination. Moreover, we started to assume that at

least β subunit should have the binding site for the histamine, as substitution of β

subunits did not abolish potentiation mediated by histamine.

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Fig. 4.12: Dependence of histamine potentiation on GABAA receptor subunit combinations. (A and B) The current traces to different concentrations of histamine (0.1 -10 mM) at EC10 GABA for α1β2 and α1β3 receptor channels respectively. Affinity for potentiation by histamine depends up on the subunit combination as incorporation of β3 subunit with α1 reduces the EC50 for GABA nearly 6 times in comparison to α1β2 receptors. (C) Concentration-response curves of histamine potentiated GABA-current for α1β2 (open circles) and α1β3 (triangles) receptor channels. Data represent the mean ± S.E.M. and curve is derived from the fit of the Hill equation to the data taken from 5 oocytes. Half maximal concentration response is changed for different β subunit combinations.

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In the second series of subunit shuffling experiment, we replaced α1 with α5 keeping the

β1 subunit constant. α1 and α5 have very less sequence homology, so we presumed that if

α subunit has the binding site for histamine than shuffling of α1 with α5 should vanish or

decrease the potentiation meditated by histamine. Fig. 4.13A shows potentiation of

GABA (2 µM) evoked current by histamine on oocyte injected with α5β1 subunit cRNA.

The marked difference between α1β1 and α5β1 is that the onset of potentiation starts at

10-30 µM in α1β1 but in α5β1 it starts at 100 µM histamine concentrations. Also,

comparing with α1β1, the average fractional potentiation mediated by α5β1 is 2.5 lesser

than that of α1β1.

α1β2γ2 is the most prominent GABA subunit combination in the CNS; it accounts nearly

70 % of the total GABAA receptors. We checked α1β2γ2 subunit combinations and indeed

it is potentiatited by histamine Fig. 4.13B. Also like α1β1, the onset of potentiation starts

at 30 µM histamine concentration. Fig. 4.13C shows the dose response curves for all

these different combinations. The EC50 for α5β1 and α1β2γ2 are 1.8 and 1.2 mM, (n = 4)

respectively.

The table 4 shows the EC50 for histamine potentiated GABA-current. The most sensitive

combination for histamine was α1β3 whereas α1β2 was found to be the least sensitive for

histamine comparing to all the subunit combinations checked in our experiments.

Fig. 4.14A and B depicts the bar-diagram of EC50 and fractional potentiation for

histamine on various subunit combinations of GABAA receptors respectively. It is

evident from the data that subunit combinations, which showed less EC50 for histamine,

enhanced maximum histamine-mediated potentiation also.

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Fig. 4.13: Dependence of histamine potentiation on GABAA receptor subunit combination. (A and B) The current traces to different concentration of histamine (0.1-10 mM) at EC10 GABA for α5β1 and α1β2γ2 receptor channels. Affinity for potentiation by histamine depends up on the subunit combinations. (C) Concentration-response curves of histamine potentiated GABA-evoked current for α5β1 and α1β2γ2 receptor channels. Data represent the mean ± S.E.M. and curve is derived from the fit of the Hill equation to the data taken from 5 oocytes. Note, synaptic GABAA receptors (α1β2γ2) are also potentiated by the histamine.

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Fig. 4.14: Comparison of potentation mediated by histamine on different GABA subunit combinations. (A) Bar-diagram showing comparison by EC50 for histamine potentiated GABA current. Note that α1β3 is the most sensitive subunit combination. (B) Bar-diagram illustrating fractional potentiation on GABAA receptors.

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Table 4: EC50 to histamine potentiated GABA current of various GABAA receptors. EC50 values for peak currents were determined with histamine concentrations ranging from 0.01 µM to 1 mM and the data were fitted to the Hill equation.

4.9: Homomultimeric channels of β1 subunit and the effect of histamine and histidine As our substitution experiments gave an idea that at least beta subunit should have the

binding site for histamine, we decided to check the expression of homomultimeric

channels formed by β1 subunits. It is reported that when β1 is injected or transfected in

oocytes or HEK 293 cells, respectively, it forms homomultimeric channels (Miko et al.,

2004). The channels formed by β1 subunit show high leakage currents i.e., spontaneous

channel activity. The spontaneous channel activity can be blocked by Picrotoxin (PTX)

which is a Cl- -channel blocker.

Oocytes were injected with β1 subunit cRNA alone and expression of the channel was

checked after 3 days by voltage-clamp technique. We first ensured that indeed there is an

expression of β1 subunit homomultimeric channels in the injected oocytes. We employed

two criteria, to check the expression, first 100 µM PTX was applied which blocks the

spontaneously opened channels formed by β1 subunit indicated by an apparent 'outward'

current evoked by PTX at oocytes clamped to -60 mV. Second, we applied 3 mM GABA

to record even the very small currents evoked by GABA and detected small evoked

inward currents. Interestingly, when we applied 3 mM histamine (~ EC50 value for α1β1

receptor) alone, we found that it directly activates the β1 channel with much higher

amplitude than GABA (Fig. 4.15A). We also checked 1 mM histidine on β1 receptors and

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like histamine, histidine directly gated β1 homomultimeric channels (Fig. 4.15B). Co-

application of 3 mM histamine with 3 mM GABA evoked almost same amplitude of

currents compared to the current induced by 3 mM histamine alone. Moreover, co-

application of 1 mM histidine with 3 mM GABA showed the same amplitude of current

like current mediated by 1 mM histidine alone. In homomultimeric β1 channels, the

amplitude of histidine-gated current was higher than the histamine gated currents up to

3/4. We tried 1 mM and 10 mM histamine concentration with 3 mM GABA to check if

the lower or higher concentration of histamine induces any potentiation. For both

concentrations, we found the same amplitude of current compared to histamine alone.

Therefore, at least at the concentrations of histamine and GABA tested, there was no

potentiation effect of histamine but rather a direct opening of the channel. The high

spontaneous activity of β1 channels caused a general instability of oocytes during

measurement of injected oocytes. Therefore, we could not check the lower concentration

of histamine on β1 receptors and extended the characterization of the receptor. A more

detailed characterization was done with the other homomultimeric β2 and β3 receptors as

described later (Chapter 5 and 6).

We investigated if is there any possibility of contribution of homomultimeric channels

formed by β1 in α1β1 subunit combination for histamine direct activation. Does

expression of β1 alone in α1β1 can cause artefacts or misinterpretation of our results? We

ruled out this possibility for two reasons: firstly, in α1β1 2 µM GABA induces a huge

GABA-evoked current whereas in case of β1 there is no response in our experimental

conditions and second, there is no spontaneous channel activity in case of α1β1 receptors

which can be confirmed by application of PTX, which does not show any blocking

activity when applied alone on α1β1 receptors.

These experiments gave us the clear idea that indeed the β1 subunit has a binding site for

histamine. The explanation for not direct activation by histamine on α1β1 receptors is

discussed in the discussion section.

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Fig. 4.15: Activation of homomultimeric β1 receptors by histamine and histidine. (A) Current traces elicited by GABA and GABA plus histamine. Expression of homomultimeric β1 receptors is confirmed by the application of 100 µM PTX, which blocks spontaneously opened channels formed by homomultimeric β1 receptors. (B) Current-traces elicited by GABA and GABA plus histidine. Both histamine and histidine with or without GABA elicited same amplitude of current.

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4.9.1: Homomultimeric channels of β3 subunit and effect of histamine There are subtle differences in the expression of β1 and β3 homomultimeric channels.

GABAA receptors that contain β3 subunits are not activated in the absence of GABA.

They do not have any spontaneously opened channel activity and on the other hand, 100

µM bicuculline nearly completely inhibits current activated by an EC50 concentration of

GABA in cells expressing homomultimeric β3 subunits but had a very small inhibitory

effect in cells expressing homomultimeric β1 subunits (Miko et al., 2004). As in our

previous experiments we found that α1β3 is the most sensitive subunit combination for

histamine potentiation, we looked for the expression of homomultimeric β3 subunit and

effect of histamine on it. Like β1 we found that β3 is also activated by 3 mM histamine

alone. The co-application of 3 mM histamine with 3 mM GABA had the same amplitude

of current mediated by 3 mM histamine alone. The characterization of β3 is presented in

more detail in Chapter 5.

4.10: Molecular cloning of ρ1 subunit of GABAC receptors We asked could GABAC receptor like GABAA receptors be potentiated by histamine. For

construction of an expression plasmid, rat GABAC receptor ρ1 subunit cDNA was cloned

in to pSGEM vector. The plasmid containing ρ1 cDNA as a template was amplified by

PCR (2.10) using oligonucleotides rho1-ATG and rho1-stop (3.2.2). A PCR fragment of

1500 bp of ρ1 subunit was generated, having BamH1 site at 5'-terminal and Xho1

restriction site at 3'-terminal. The pSGEM plasmid was digested by restriction Enzyme

BamHI and Xho1 and PCR fragment by BamHI and XhoI (3.3.1). Fragments were

separated using gel electrophoresis (3.1.2) and purified using a gel extraction kit (3.1.5).

After ligation (3.3.5) and transformation into E. coli XL1-blue (3.3.7), plasmids isolated

from several colonies (3.1.8) were analyzed by restriction mapping (3.3.1) and from

clones with the correct insert, large amounts of plasmid was prepared by the maxi prep

method (3.1.9). The identity and integrity of the cDNA insert was confirmed by

sequencing (3.3.8) using primers flanking the polylinker site (T7 and pSGEM-down) (Fig.

4.16).

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Fig. 4.16: Schematic representation: Cloning of ρ1 subunit of GABAC receptors. For cloning of pSGEM-ρ1 the open reading frame of the ρ1 cDNA insert present in pCDNA3 plasmid used as template was amplified by PCR using specific oligonucleotides. A PCR fragment of 1500 bp of ρ1 subunit was generated. The pSGEM plasmid was digested by restriction Enzyme BamHI and Xho1 and PCR fragment by BamHI and XhoI and cloned into the corresponding polylinker sites of the dephosphorylated pSGEM vector.

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4.10.1: GABAC receptors: No potentiation by histamine and histidine We analysed GABAC receptors to test if GABAC receptors can also be potentiated by

histamine. Interestingly, histamine up to 10 mM did not modulate the GABA-elicited

currents at 200 nM GABA (EC10 ρ1 receptor channels). Fig. 4.17 shows insensitivity of

ρ1 receptor channels for modulation by histamine. In contrast to GABAA receptors, ρ1

was not potentiated by histamine at any GABA concentration tested. Therefore, we

conclude here that potentiation by histamine is specific only for GABAA receptors.

Fig. 4.17: No potentiation by histamine and histidine on GABAC receptors. The current traces for the EC50 GABA alone and in the presence of histamine and histidine. Both histamine and histidine do not show any potentiation effect on GABA-evoked current.

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4.11: Possible mechanisms of the histamine action In the next set of experiments, the nature of the histamine action on GABAA receptors

was examined in greater detail. In the first set of experiments, possible binding sites were

narrowed down by a comparison of the histamine action with other potentiating agents

(4.11.3), then the action of histamine on the GABA affinity of α1β1 receptors was

characterized (4.11.4) and finally, the binding site for histamine on the β1 subunit was

identified by the investigation of mutated GABA receptors (4.11.5, 4.11.8).

4.11.1: Histamine binding site is different from pentobarbital binding site on β3 subunit of GABAA receptors β subunits of GABAA receptors have binding site for various modulators like

pentobarbital and propofol (Bali and Akabas 2004; Karsowksi et al., 1998; Sanna et al.,

1995). The following experiments were performed with the β3 subunit because of better

expression success compared to β1 homomultimeric channels. We first checked any

similarity of histamine binding site with pentobarbital binding site. Pentobarbital

activated homomultimeric β3 subunit when applied alone (30-100 µM). Fig. 4.18A shows

the traces for current evoked by 100 µM pentobarbital. Bemegride is the potent

antagonist for pentobaribital. When 3 µM bemegride was applied with 100 µM

pentobarbital it inhibited pentobarbital-evoked current more than 50 % (Fig. 4.18A).

Bemegride acts as a reversible antagonist because the GABA-evoked current returned to

its normal level after washing out the bemegride within 1 minute. Fig. 4.18B shows the

traces of histamine-evoked currents on β3 subunit. 500 µM histamine-evoked current

(~EC50 of histamine on α1β1 receptors) is not blocked by 3 µM bemegride. To exclude

the possibility that 3 µM bemegride was a very small concentration for higher histamine

(500 µM) concentration, we either decreased histamine concentration and increased

bemegride concentration and vice-versa. Even in these conditions we did not find any

inhibition for histamine-evoked current by bemegride. The next question was if

bemegride was a specific blocker for pentobarbital. Therefore, bemegride action on

propofol activation was investigated. Application of 50 µM propofol, on homomultimeric

β3 subunit activated the channel. Fig. 4.18C shows the current traces of 50 µM propofol

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and effect of 3 µM bemegride on propofol evoked current. 3 µM bemegride did not block

the propofol-evoked current, we also applied higher concentration of bemegride (10 µM)

but found no inhibition by bemegride. All these experiments suggested that pentobarbital

should act on a different binding site compared to histamine and propofol.

4.11.2: Histamine binding site is similar to propofol binding site on β3 subunit of GABAA receptors - Experiments on homomultimeric β3 subunit We decided to check the possibility of any similarity between propofol and histamine

binding site. There is no specific propofol antagonist existing in our knowledge so, we

could not proceed in previous experimental way. But for the histamine action on GABA

receptors there are several candidates existing, namely the histamine analogs acting on

metabotropic histamine receptors. We first checked those antagonists for histamine-

evoked current. Screening of various metabotropic histamine receptor antagonists pointed

thioperamide (H3 antagonist) as the potent antagonist for histamine-evoked current on

homomultimeric β3 subunits. Fig. 4.18E shows the inhibition of histamine-evoked current

by thioperamide. 30 µM thioperamide inhibits more than 50 % of the current evoked by

500 µM histamine. Remarkably, when 30 µM thioperamide was co-applied with 300 µM

propofol it blocked the propofol-evoked current also more than 50 % (Fig. 14.8F).

Furthermore, we checked the activity of thioperamide on pentobarbital-evoked current.

Application of 30 µM thioperamide did not show any inhibition on pentobarbital-evoked

current Fig. 4.18D. All these experimental data revealed three important things. First time

we showed that thioperamide can act as an antagonist for propofol and hence we identify

the new antagonist for propofol action. Second, these experiments give further arguments

that pentobarbital has a different binding site than histamine. Third, as both propofol and

histamine are blocked by thioperamide with same efficacy, which lead to

pharmacological confirmation that propofol and histamine may have the same binding

site. In the later described pharmacological characterization of β3 homomultimeric

channels (5.4.1) it was found that even the IC50 for thioperamide was comparable for the

block of propofol as well as histamine-evoked currents.

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Fig. 4.18: Histamine binding site is similar to propofol binding site. (A) Inhibition of pentobarbital- evoked current by bemegride, which acts as a reversible antagonist. (B and C) Both histamine and propofol-evoked currents are not inhibited by bemegride. (D) Pentobarbital-evoked current is not inhibited by thioperamide (H3 antagonist). (E and F) Thioperamide blocks histamine and propofol-evoked currents.

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4.11.3: Histamine binding site is similar to propofol binding site on β3 subunit of GABAA receptors - Experiments on heteromultimeric α1β1 receptors The preliminary electrophysiological experiments gave us an idea that histamine may

have the similar binding site like propofol. We asked the question if this is the case then,

if propofol and histamine are mixed together at their EC50 they should potentiate the

GABA-evoked current like if one of them is applied at nearly to its EC80-90. As having the

same binding site means, at EC50 this binding site will be half occupied by one of the

modulators, so addition of another agonist should lead in the enhancement of GABA-

evoked current. Second, if both of them are mixed together at their EC90 and co-applied

with 2 µM GABA, then there should not be any additional potentiation. The reason

behind it was that at saturation concentration, the modulator should almost occupy the

modulator’s binding site so addition of a modulator having the similar binding site should

not enhance any more potentiation. If it is not the case then in EC90 mixing experiment,

addition of another modulator should lead more enhancement of GABA-evoked current

as presence of two modulators with different binding sites should potentiates GABA-

evoked current more efficiently.

Fig. 4.19A shows the current-response relationship for propofol and histamine

potentiation on α1β1 receptor at EC10 GABA. When 3 µM propofol (~ EC50 of propofol

for α1β1 receptor) was co-applied with 2µM GABA, it enhanced the GABA current up to

170 %. With 2 µM GABA, 1 mM histamine (~ EC50 value of histamine potentiation on

α1β1 receptors) evoked almost same amplitude (155 %) of potentiation, similar to

propofol. When 3µM propofol and 1 mM histamine were co-applied with 2 µM GABA,

they increased the GABA-evoked current amplitude up to 250 %; that means simply both

of them showed an additive effect, which was an indication that they somehow may act

on the similar binding site.

Fig. 4.19B depicts the similar experimental strategy but here the parameters were

modified. We checked the effect of EC90 of both modulators alone and in combination of

each other with 2 µM GABA. 10 µM propofol (~ EC90 of propofol for α1β1 receptor)

increased the amplitude of GABA-evoked current at EC10 GABA nearly 300 %. 10 mM

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histamine (~ EC90 of histamine for α1β1 receptor) induced nearly 200 % increase in

amplitude for GABA-evoked current. As expected, the combination of both 10 µM

propofol and 10 mM histamine at EC10 GABA did not enhance the GABA-evoked

current more than 300 %. So, co-application of both modulators at their EC90 did not

enhance more amplitude of GABA-evoked current w.r.t when they applied alone. From

these electrophysiological experiments we were able to get the notion that both

modulators are acting on the same binding site.

In the next series of experiments we investigated the different GABAA receptor subunit

combination. Oocytes injected with α5β1 subunit cRNA were measured after 2-3 days.

We employed the same strategy like previous α1β1 experiments. First we checked the

effect of propofol, histamine alone and in combination with the co-application of 2 µM

GABA at their EC50 concentration. The combination of histamine and propofol showed

the additive effect and increased the GABA-current nearly two fold comparing to current

evoked by histamine or propofol with 2 µM GABA (Fig. 4.20A). Next, we checked the

co-application of histamine and propofol alone and histamine plus propofol with 2 µM

GABA at their EC90 concentration. Interestingly, there was no additive effect. The

amplitude of the current evoked by histamine or propofol with GABA alone or histamine

plus propofol with GABA remained almost same (Fig. 4.20B). This experiment also gave

us the further proof that histamine and propofol have same binding sites.

4.11.4: Effect of histamine on EC50 of GABA on GABAA receptors Fig. 4.21B shows dose-response curves, plotted using hill equation, for GABA alone

(filled symbols) and GABA plus 3 mM histamine (open symbols) for α1β1 receptors. The

data for GABA alone at concentration below about 0.1 µM are on a straight line. At

higher concentration, the increments in response with increase in concentration declined,

and 1 mM GABA gave a nearly maximal response. A further increase in the

concentration from 1 mM to 3 mM caused a slight decrease in the response. However,

this may have arisen because of desensitization during relatively slow drug application.

When histamine (3 mM) was applied together with GABA, the dose-response curve for

GABA was remained same.

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Fig. 4.19: Histamine binding site is similar to propofol binding site. (A) Traces show current recorded from an oocyte expressing α1β1 GABAA receptors. GABA-evoked current is potentiated by both histamine and propofol, applied at their EC50. The co-application of histamine and propofol at their EC50 concentration with 2 µM GABA, elicited almost double amplitude of current compared with histamine or propofol. (B) Co-application of histamine and propofol at their EC90 with 2 µM GABA did not show additive effect.

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Fig. 4.20: Histamine binding site is similar to propofol binding site. (A) Traces show current recorded from an oocyte expressing α5β1 GABAA receptors. GABA evoked-current is potentiated by both histamine and propofol, applied at their EC50. The co-application of histamine and propofol at their EC50 concentration with 2 µM GABA, elicited almost double amplitude of current compared with histamine or propofol. (B) Co-application of histamine and propofol at their EC90 with 2 µM GABA did not show additive effect.

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We wanted to check, no GABA affinity shift for other receptors also, and so oocytes

were injected with α1β2 subunits cRNA. Fig. 4.21C shows the current-response

relationship for α1β2 receptors with GABA alone (filled symbol) and GABA plus 10 mM

histamine (open symbols). Histamine did not shift the EC50 of GABA significantly also.

GABA alone has EC50 of 3.6 µM whereas in presence of histamine the EC50 is 3.9 µM, (n

= 5). Therefore, data from α1β1 and α1β2 clearly indicated that histamine does not shift

the EC50 of GABA.

Fig. 4.21: Concentration-response relationship of GABA in the presence of histamine. (A) Dose-response relationship of the currents activated by GABA and GABA plus 3 mM histamine on α1β1 GABAA receptors respectively. (B and C) Concentration-curves plotted using Hill equation for GABA alone (filled circle) and GABA plus histamine (open circle) for α1β1 and α1β2 receptors respectively. Note that co-application of histamine with GABA does not change the EC50 of GABA.

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Whereas most of the GABAA expression oocytes showed no histamine-induced shift in

the GABA affinity, there were some few exceptions. The amount of potentiation by

histamine is considerable variable. In some rare oocytes expressing α1β1 and α1β2, 10

mM histamine showed no effect at 10 µM GABA but in some of these oocytes, histamine

potentiated the action of low concentrations of GABA (2 µM) consistent with the idea

that under certain circumstances, histamine might influence the GABA affinity (Fig.

4.22).

Fig. 4.22: In few oocytes expressing α1β1 or α1β2, 10 mM histamine showed no potentiation of the current evoked by 10 µM GABA but potentiated currents evoked by lower concentrations of GABA. Example for α1β1 expression oocytes (A) that showed an EC50 shift from 1.25 to 0.68 µM GABA induced by 10 mM histamine. GABA dose-response curves with and without histamine were determined on the same individual oocytes (n = 3). Also rare α1β1 expressing oocytes showed a similar behaviour (C).

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4.11.5: Molecular cloning of point mutation in β1 subunits Previous reports have suggested that Methionine at 286 position (M286) of the GABAA

receptors β1 subunit is involved in propofol potentiation (Krasowski et al., 1998). To

determine whether this domain in the GABAA β1 subunit plays a similar role in histamine

mediated potentiation, we decided to create a point mutation (M286W) at position

corresponding to the mutation in the GABAA receptor β1 subunit that is important for the

propofol binding. Rat GABAA receptor β1 subunit cDNA was cloned into pSGEM vector

(4.2). To mutate the methionine to tryptophan in the GABAA receptor β1 subunit at 286

position, site-directed mutagenesis (3.2.3) was performed by using mutated

oligonucleotides in conjunction with a primer-directed polymerase chain reaction (3.2.2).

A mutated DNA fragment of the β1 subunit coding TM3 to C-terminal end including the

mutation M286W was PCR amplified using primers having a BamHI site at the 3´end. In

the amplified PCR fragment at 5' end the EcoRV site was kept preserved. The plasmid

containing cDNA of β1 subunit was digested by restriction enzymes BamHI and EcoRV

fragments were separated using gel electrophoresis (3.1.2) and purified using a gel

extraction kit (3.1.5). The mutated, BamHI-EcoRV restricted PCR fragment was ligated

into the pSGEM vector. After ligation (3.3.5) and transformation into E. coli XL1-blue

(3.3.7), plasmids isolated from several colonies (3.1.8) were analyzed by restriction

mapping (3.3.1) and from clones with the correct insert, large amounts of plasmid was

prepared by the maxi prep method (3.1.9). The identity and integrity of the cDNA insert

was confirmed by sequencing (3.3.8) using primers flanking the polylinker site (T7 and

pSGEM-down) (Fig. 4.23).

4.11.6: β1 (M286W) mutation completely abolishes potentiation mediated by histamine Previous reports have suggested that Methionine at 286 position (M286) of the GABAA

receptors β1 subunit is involved in propofol potentiation (Krasowski et al., 1998) To

determine whether this domain in the GABAA β1 subunit plays a similar role in

histamine-medited potentiation, we created point mutation (M286W) at position 286

(4.11.6) corresponding to the mutation in the GABAA receptor β1 subunit that is

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Fig. 4.23: Schematic representation: Cloning of β1 (M286W) subunit of GABAA receptors. To mutate the methionine to tryptophan in the GABAA receptor β1 subunit at 286 position, site directed mutagenesis was performed using mutated oligonucleotides in conjunction with a primer-directed polymerase chain reaction. A mutated DNA fragment of β1 subunit coding TM-3 to C-terminal end including the mutation M286W was PCR amplified using primers having a BamH1 site at the 3´ end. In the amplified PCR fragment at 5´end the EcoRV site was kept preserved. The mutated, BamH1, EcoRV restricted PCR fragment was ligated in to the pSGEM vector.

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important for the propofol binding. The mutant receptors were then characterized by

determining the potentiation by histamine on GABA-evoked current. Fig. 4.24A

represents a schematic representation for the mutation done on β1 subunit. This mutation

lies in between the TM2 and TM3 domains and specifically to the exterior or outer most

portion of TM2 domain.

Fig. 4.24B shows the electrophysiological characterization of the mutation. Oocytes were

injected with α1β1 (M286W) subunit combinations. Firstly, we checked the potentiation

mediated by propofol on mutant receptor. At EC10 GABA (2 µM), propofol was checked

with increasing concentration (1-10 µM). No potentiation of GABA-evoked current by

propofol was in accordance with previous mutational study done by other groups

(Karsowski et al., 1998). Interestingly when histamine was checked at EC10 GABA, it did

also not induce any potentiation. Fig. 4.24B describes the current traces for histamine. In

α1β1 (M286W), three different concentration dependent effects were checked. We tested

various concentration of histamine (0.1 - 30 mM) at EC50 (~10 µM) and EC90 (~1 mM) of

GABA to completely rule out any potentiation that could be attributed by these

concentrations. Therefore, from these experiments we confirmed that histamine binds to

the similar binding site of propofol.

4.11.7 Histamine and propofol have similar binding sites Fig. 4.25 depicts the effect of β1 subunit mutation on propofol and histamine mediated

potentiation in α1β1 (M286W) expression oocytes. At EC10 GABA, 30 mM histamine did

not show any potentiation that indicates even at that high concentration mutation is so

pronounced that it abolishes all the activity of histamine. It is of special interest also, as in

some studies it has been shown that at very higher concentration of agonist, there is some

degree of activation or potentiation like in pentobarbital mutational study. Induction of no

potentiation by histamine on α1β1 (M286W) shows clearly that M286 is the vital amino

acid for the histamine binding on β1 subunit. Moreover, we also found that 10 µM

propofol with 2 µM GABA does not induce any potentiation. We also did some mixing

experiments where 30 mM histamine was co-applied in the presence of 2 µM GABA and

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Fig. 4.24: Complete abolition of potentiation by histamine on α1β1 (M286W) mutant GABAA receptors. (A) Schematic diagram of β1 subunit of GABAA receptor. Topology of a single subunit is highlighted showing the four transmembrane domains (TM1-TM4) and the location of the mutated amino acid in TM-3 domain and (B) GABA−α1β1 M286W-expressing oocytes were voltage-clamped at -40 mV and a series of GABA (2 µM) plus histamine mixes were bath-applied for 10 s. Application of histamine up to 10 mM with EC10 GABA did not induce any potentiation. Current traces from wild type α1β1 receptor for histamine are shown at right side for comparison.

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Fig. 4.25: Mutation at 286 position on β1 subunit of GABAA receptors abolishes the potentiation mediated by propofol as well as histamine. (A) Current traces from the wild type α1β1 receptors showing dose-dependent potentition evoked by propofol at EC10 (2 µM) GABA. (B) α1β1 (M286W) mutant receptors are not potentiated by the propofol. (C) Co-application of saturating concentration of histamine with or without propofol at EC10 GABA, does not potentiate α1β1 (M286) mutant receptors.

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10 µM propofol. Even at that high concentrations, we did not observe any potentiation,

whereas in our previous results we have shown that mixing experiment in α1β1

potentiated GABA-evoked current in the presence of 30 mM histamine and 10 µM

propofol with 2 µM GABA. These experimental data also give further proof that

histamine and propofol has same binding site.

4.11.8: β1 (M286W) mutation completely abolishes potentiation mediated by histidine We also investigated the effect of β1 subunit mutation on the potentiation of histidine.

Fig. 4.26A shows the current-traces where oocytes were injected with α1β1 (M286W)

subunit combination. Application of 1 mM histidine (~ EC99 in α1β1 receptors) with 2 µM

GABA, did not show any potentiation. We checked various concentration of histidine (1-

1000 µM) to clearly characterise the abolition of histidine-mediated potentiation by

mutation. Also we did experiments with different GABA concentration like EC50 and

EC90 GABA, there also keeping all the parameters similar but we could not detect any

potentiation (Fig. 4.26B and C). From these data we demonstrate here that like histamine,

histidine has the same binding site on β1 subunit at the same location similar to propofol

binding site.

4.11.9: Sequence alignment with GABAC receptors depicts that histamine has similar binding site as propofol Moreover, sequence alignment of ρ1 subunit with various beta subunits of GABAA

receptors showed that at position 286 in ρ1 subunit there is tryptophan (W) instead of

methionine (M), which makes ρ1 subunit insensitive to propofol. From this analysis two

important conclusions were derived. First, GABAC receptors are not potentiated by

histamine and histidine; second, having tryptophan instead of methionine at position 286

makes ρ1-receptor insensitive for propofol as well as histamine that give also another

proof for same binding site for propofol and histamine.

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4.11.10: β1 (M286W) mutation does not interfere with the potentiaion mediated by other modulators We also examined mutant receptor sensitivity to other modulators like neurosteroids and

pentobarbital, which have the binding site on β1 subunit. Fig.4.27A shows the current

traces from the oocytes injected with α1β1 subunits cRNA. Application of 50 µM

pentobarbital and 50 µM of the 5α-Pregnane-3α-ol-11, enhanced GABA evoked currents

up to 2–2.5 and 3–3.5 respectively. Fig. 4.27B, shows the current-traces for the response

of the Neurosteroide and pentobarbital on α1β1 (M286W) receptor. Both modulators

potentiated the mutant receptor at same concentration, checked with wild type α1β1 with

almost same efficacy. Therefore, mutation is specific only for the propofol and histamine.

It does not interfere in the activity of other modulators at least for neurosteroids and

pentobarbital.

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Fig. 4.26: Complete abolition of potentiation by histidine on α1β1 (M286W) mutant GABAA receptors. (A, B and C) Current traces for dose-response curve for histidine at EC10, EC50. EC90 of GABA respectively. GABA− α1β1 (M286W) - expressing oocytes were voltage-clamped at -40 mV and a series of GABA (2 µM) plus histidine mixes were bath-applied for 10 s. Co-application of hisitidine with GABA did not induce any potentiation. Current-traces from wild type α1β1 receptor for histidine are shown at right side for comparison.

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Fig. 4.27: M286W mutation on β1 subunit does not affect other modulators activity. (A) Wild type α1β1 receptors were exposed to propofol, pentobarbital and neurosteroide in the presence of EC10 GABA. The response elicited by GABA was potentiated in the presnece of mentioned modulators. (B) On α1β1 (M286W) mutated receptor GABA-evoked current was potentiated by both pentobarbital and neurosteroide whereas mutation abolished the propofol potentiation.

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Summary of major findings of the first part of my thesis:

1. Identification of two new modulators for GABAA receptors. 2. GABAA receptors are potentiated by both histamine and histidine. 3. Potentiation of GABAA receptors by histamine does not arise from any shift in

equilibrium potential or change in form of the current-voltage relationship. 4. Histamine-mediated potentiation is not restricted to the oocyte recombinant

system but also shown in HEK 293 cells.

5. Affinity for potentiation by histamine depends up on the subunit combinations. 6. Our experiments show that histamine-binding site is located on β subunits of

GABAA receptors. 7. Our pharmacological and mutational studies show that indeed histamine binding

site is located at M286 on β1 subunit of GABAA receptors, which is the binding site for propofol also.

8. GABAC receptors are not modulated by histamine.

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Chapter 5 5: Characterization of homomultimeric β3 channels

In our previous experiments, we found that both β1 and β3 are activated by histamine

alone. We asked the question, do β3 subunits of GABAA receptors behave like possible

histamine-gated chloride channels? For this reason, we decided to characterize β3

subunits in detail. Homomultimeric β1 subunits form spontaneously opened channel,

which have very high leakage current, because of which we could not fully characterize

β1 subunit. Homomultimeric β3 receptors were more stable and showed very less leakage

current, so we extended our experiments on β3 subunit of GABAA receptors.

5.1: Homomultimeric β3 receptors behave like histamine-gated ion channels Injection of β3 cRNA in to Xenopus oocytes resulted in expression of substantial

histamine-activated currents not seen in uninjected oocytes. At a holding potential of -50

mV, micro molar concentrations of histamine rapidly activated inward currents that

deactivated quickly after washout of histamine, (Fig. 5.1A). As the histamine

concentration was raised from 10 µM to 10 mM, there was a progressive increase in

histamine-evoked current that seemed to reach at plateau at about 10 mM histamine.

Representative histamine dose response curves are shown in Fig. 5.1C. Fitting the data to

the Hill equation yielded an EC50 of 597 ± 5 µM, (n = 7) for the β3 subunit injected

oocytes.

The effect of histidine, the histamine metabolite, was also checked on β3 homomultimeric

channels. Results from five oocytes, exposed to a range of histidine concentrations are

shown in Fig. 5.1B. As the histidine concentration was raised from 10 µM to 10 mM,

histidine-evoked currents increased in amplitude in a dose-dependent manner. The

histidine-evoked current has an EC50 of 1.135 ± 0.15 mM, (n = 5). There are subtle

differences in the activation by histamine and histidine. First, histamine activates β3

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homomultimeric channels with less than the half maximal concentration than histidine.

Second, histidine induces higher amplitude current (2-4 folds larger) than histamine at

saturating concentrations determined in the same individual oocytes.

We also checked another metabolite in histamine pathway. Telemethyhistamine (t-MHA)

is produced in synapses by the methylation of histamine by histaminemethyltransferase

(Haas and Panula, 2003). t-MHA-currents were first detectable at about 30 µM

concentration, increased steeply in size as the t-MHA concentration was raised and

reached a maximum with about 30 mM t-MHA (Fig. 5.2). The EC50 derived from fitting

data to Hill equation was 1.1 ± 0.13 mM, (n = 3).

5.2: Histamine-gated homomultimeric β3 receptors behave like typical ligand-gated chloride channels To identify the ion permeating through the receptor channel, the current reversal potential

was determined while activating the β3 receptor with histamine (Fig. 5.3). The membrane

current activated by GABA decreased in size as the oocyte was depolarized and inverted

direction at about -20 mV, which corresponds to the chloride equilibrium potential in

Xenoups oocytes (Barish et al., 1983; Kusano et al., 1982). Moreover our current-voltage

relationship fits with β2 and β3 reversal potential reported by various groups where they

showed that reversal potential for β2 and β3 was close to the chloride equilibrium

potential (-23 mV) when expressed in Xenopus oocytes (Ymer et al., 1989; Dascal et al.,

1984).

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Fig. 5.1: Homomultimeric channels of β3 subunit and effects of histamine and histidine. (A and B) The dose-response relationship for histamine and histidine from an oocyte expressing homomultimeric β3 receptors. Bars over current traces indicate the duration of histamine or histidine application with the concentration of applied agonist in µM. The traces in A and B were obtained from the same oocytes. The holding potential was -50 mV. (C) Concentration-response curves for histamine (open circle) and histidine (closed circle) for homomultimeric β3 receptors. Data points shown are the means of multiple normalized experiments and curve was derived from the fit of Hill equation (n = 5).

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Fig. 5.2: Dose-response curve of t-MHA for homomultimeric β3 receptors. Oocytes were voltage-clamped (-50 mV) 3-6 days after injection of in vitro transcribed cRNA encoding β3 subunit of GABAA receptors. Various concentration of t-MHA applied for 10 seconds produced inward currents. Bars indicate the duration of application of agonist.

Fig. 5.3: Current-voltage relationship of the current activated by histamine in an oocyte injected with β3 subunit cRNA of GABAA receptors. Points indicate peak currents elicited by 500 µM histamine applied with membrane potential clamped at different levels.

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5.3: Relative comparison of various agonist of histamine with GABA - Relative agonists efficacy compared to GABA To find the relative comparison between maximal current induced by GABA or

histamine, we expressed β3 subunit cRNA in Xenopus oocytes. We could not detect any

current of GABA up to 1 mM GABA concentration but from 3 mM GABA we could

detect a very small GABA-evoked current. Oocytes exhibiting inward current in response

to 3 mM GABA greater than 25 nA (at -50 mV) in amplitude were selected for this

experiment. So β3 homomultimeric channels are also activated by GABA but with very

small amplitude. Next, we determined the sensitivity of these homomultimeric receptors

to histamine, histidine and t-MHA. Fig. 5.4A shows the current traces for GABA,

histamine, histidine and t-MHA applied to the same individual oocyte. The bar graphs in

Fig. 5.4B shows the average percentage activation of β3 homomultimeric channel by

histamine and its precursors. We compared 3 mM GABA with saturating concentration

(10 mM) of histamine, histidine and t-MHA. 10 mM t-MHA induces 3 folds higher

amplitude of current than 3 mM GABA. 10 mM histidine showed the highest amplitude

(9 folds) of current w.r.t. 3 mM GABA whereas, 10 mM histamine showed moderate

current enhancement up to 6 folds.

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Fig. 5.4: Relative comparison of current evoked by histamine and its metabolites with GABA. (A) The current traces for GABA, histamine, histidine and t-MHA applied on the oocyte expressing homomultimeric β3 receptors. Saturation concentration was taken for each agonist. (B) Average percentage activation of homomultimeric β3 receptors by histamine and its metabolites (n = 3).

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5.4: Pharmacological characterization of histamine-gated homomultimeric β3 receptors After establishing that β3 is indeed behaves like histamine-gated ion channels, we asked

whether the histamine-evoked current on homomultimeric β3 receptors, can be blocked

by known antagonists of metabotropic histamine receptors. Fig. 5.5A is a plot of sub

maximal histamine-activated currents for β3 homomultimeric channels and effect of

thioperamide (H3 receptor antagonist) on it. Thioperamide significantly reduced the half

maximum histamine-current amplitude evoked by 500 µM histamine in a dose-dependent

manner. The IC50 value for thioperamide is 9.7 ± 0.8 µM, (n = 5). Also, at high

concentration 300 µM thioperamide blocks the spontaneously open channels formed by

β3 subunit. Homomultimeric β3 currents evoked by 500 µM histamine were inhibited in

concentration-dependent manner by famotidine (Fig. 5.5B). Famotidine (H2 receptor

antagonist) maximally inhibited the histamine-activated current, with an IC50 of 77.1 ±

8.9 µM, (n = 5). The affinity of thioperamide was higher than the famotidine. We also

checked cimetidine (another H2 receptor antagonist) up to 500 µM concentration but did

not find any inhibition of the current evoked by 500 µM histamine. Also pyrilamine (H1

receptor antagonist) did not block any histamine-evoked current on homomultimeric

β3 receptors up to 500 µM concentration. Fig. 5.5C shows current-dose inhibition by

thoperamide and famotidine dose-response curve was fitted by Hill equation.

We observed that β3 homomultimeric channels were strongly picrotoxin sensitive.

Picrotoxin inhibition was fully recovered. Picrotoxin inhibited histamine-activated

current with an IC50 of 300 nM, (n = 3). HTMT, an agonist for homomultimeric receptors

also reduced the histamine activated current with an IC50 48.7 ± 2.5 µM, (n = 3) (Fig.

5.6A). Harmane and related alkaloids, such as harmine and harmaline, are agents with a

β-carboline structure. These compounds are also found endogenously in mammalian

tissues, including central nervous system. These harmala alkaloids have some

pharmacologic effects involved in convulsive or anticonvulsive actions. Harmane and

other related β-carbolines are putative endogenous ligands of the benzodiazepine

receptor. Harmane is a competitive inhibitor of benzodiazepine receptor binding in vitro.

We checked the activity of harmane on histamine-evoked currents on β3 receptors. Fig.

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5.6B shows the dose-response current inhibiton by harmane. The amplitude of histamine

evoked current decreased with increasing concentration of harmane. Harmane blocks

histamine current with an IC50 of 82.7 ± 6.3 µM, (n = 4). Like also thioperamide and

famotidine at higher concentration it shows outward current which indicates the

inhibition of spontaneously activated open channels.

5.4.1: Thioperamide acts as a competitive blocker for histamine-evoked current The tested antagonists like thioperamide, famotidine and pyrilamine are competitive

antagonists for the metabotropic histamine receptors. In the next set of experiments it was

tested if thioperamide acted as a competitive antagonist for the histamine action on

homomultimeric GABA β3 receptors also. The current evoked by different concentration

of histamine was effectively blocked when thioperamide was given in 30 µM

concentration. At rising concentrations of histamine the thiopermide block was less

pronounced and at 10 mM histamine, 30 µM thioperamide didn't cause any block of the

histamine induced current at all (Fig. 5.7A). A dose-response curve constructed from

three of such measurements showed that thioperamide shifted the dose-response curve to

the rightward direction and the EC50 of histamine shifted from 597 µM to 2.72 mM (Fig.

5.7B), a clear indication that the blocking mechanism is competitive. Competitive

blockers can act in two different ways. First, competitive blockers acting also in the

absence of the agonist by blocking for example population of spontaneously active

receptors or channels are also called 'inverse agonists'. Second, blockers showing no

action in the absence of the agonist are called 'neutral antagonist'. It was found that

thioperamide also blocked the population of spontaneously open channels in β2 and β3

homomultimeric channels (data not shown). Such action of competitive antagonist

classifies thioperamide as an 'inverse agonist'.

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Fig. 5.5: Dose-dependent inhibition by thioperamide and famotidine of histamine-activated current in homomultimeric β3 receptors. (A and B) The dose-response inhibition of histamine-evoked current by thioperamide and famotidine from oocyte expressing homomultimeric β3 receptors. Bars over current traces indicate the duration of application of 500 µM histamine and histamine plus thioperamide \ famotidine in µM concentration. The holding potential was -50 mV. (C) Average dose-inhibition curve for thioperamide (open circle) and famotidine (closed circle) on histamine-activated current respectively (n = 5). Data points shown are means of normalized experiments and curve was fitted using Hill equation.

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Fig.5.6: Dose-dependent inhibition by HTMT and harmane of histamine-activated current in homomultimeric β3 receptors. (A, B) The dose-response inhibition of histamine-activated currents by HTMT and harmane from oocyte expressing homomultimeric β3 receptors. Bars over current traces indicate the duration of application of 500 µM histamine and histamine plus HTMT \ harmane in µM concentration. The holding potential was -50 mV. (C) Average dose-inhibition curve for HTMT (closed circle) and harmane (open circle) on histamine-activated current respectively (n = 3). Data points shown are means of normalized experiments and curve was fitted using Hill equation.

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Fig. 5.7: Thioperamide acts as a competitive antagonist. (A) Current traces showing inhibition of histamine-evoked current with 30 µM thioperamide. (B) Concentration-response curve for histamine alone (closed circle) and histamine with 30µM thioperamide (open circle). Note thioperamide shifts the EC50 of histamine-evoked current to right side.

5.5: Comparison of L- and D–histidine action on β3 homomultimeric receptors Optical isomers are two compounds which contain the same number and kinds of atoms,

and bonds (i.e., the connectivity between atoms is the same), and different spatial

arrangements of the atoms, but which have non-superimposable mirror images. Each

non-superimposable mirror image structure is called an enantiomer. The two enantiomers

of histidine are L- & D-histidine.

L-Histidine D-Histidine

We investigated, can the optical activity alter the behavior of histidine and does both

compounds equally potent in activating β3 receptors. Fig. 5.8A and B shows current-

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traces of L- & D-histidine. Induction of D-histidine current started from 100 µM and had

EC50 of 1.1 ± 0.13 mM, (n = 3). The major difference in the activities of L- & D-

histidine was that current evoked by D-histidine had smaller amplitude than L-histidine

(at all concentrations, we checked).

Fig. 5.8: Comparison of current evoked by L- and D-histidine on β3 receptors. (A, B) The dose-response relationship for L- and D-histidine respectively. L-histidine induces higher magnitude of current than D-histidine though both have the same EC50 values.

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5.6: Inhibition of propofol-induced current by thioperamide on homomultimeric β3 receptors In our previous experiments on α1β1 receptors, we found that histamine and propofol

have similar binding site. After characterization of β3 receptors pharmacology for

histamine, we looked for the activity of propofol on β3 receptors. We hypothesized that

like histamine, propofol should behave like histamine and propofol alone should induce

propofol-evoked current. Indeed, when propofol alone was applied on β3 receptors, it

started to induce a propofol-evoked current at 30 µM propofol concentration. We could

not make the dose-response curve for propofol. However, gradual increase of propofol

from 30 µM - 10 mM mediated propofol-evoked current in a dose dependent manner but

did not lead to the maximal saturation of the receptor. This can be described as that in

hetromultimeric recombinant receptors; higher concentration of propofol starts the direct

activation of the channel. In our experiments for dose-response curve, this can be the

case. Therefore we decided to use equivalent concentration of propofol w.r.t. histamine.

300 µM propofol (~ EC50 of histamine = 597 µM on β3 receptors) was taken for further

experiment, which produced current > 1 µA. In β3 receptors, thioperamide acts as a

competitive antagonist and inhibits 500 µM histamine-evoked current at IC50 of 9.7 µM.

Therefore, we checked the pharmacology of propofol by thioperamide. Fig. 5.9 shows,

dose-dependent inhibition of propofol-evoked current by thioperamide. Thioperamide

inhibits the propofol-evoked current with an IC50 of 9.5 ± 0.05 µM, (n = 5). Remarkably,

this IC50 value for propofol-evoked current is almost similar to histamine-evoked current

on β3 receptor for thioperamide. This pharmacological experiment gives another strong

support for similar binding site for histamine and propofol.

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Fig. 5.9: Dose-dependent inhibition by thioperamide of propofol activated current in homomultimeric β3 receptors. (A) Current traces showing thioperamide mediated inhibition of propofol-activated currents. (B) Average dose-inhibition relation of thioperamide on propofol-activated current respectively. Data represent the mean ± S.E.M. and curve is derived from the fit of Hill equation to the data taken from 5 oocytes.

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Summary of major findings:

1. Homomultimeric β3 subunit forms histamine-gated ion channel with an EC50 of 597 µM.

2. Homomultimeric β3 receptors are also activated by histidine and t-MHA with an

EC50 of 1.1 and 1.1 mM respectively.

3. Metabotropic histamine H3 receptor antagonist thioperamide acts as a competitive inhibitor for histamine-evoked current with an IC50 of 9.7 µM.

4. Histamine evoked current can be inhibited by famotidine (H2 receptor antagonist),

harmane and HTMT with a relative order of HTMT > famotidine > harmane.

5. Homomultimeric β3 receptors are also activated by propofol alone. Propofol-evoked current can be blocked by thioperamide.

6. Inhibition of both histamine and propofol-evoked current by thioperamide gives

also evidence that both histamine and propofol have same binding site.

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Chapter 6

6: Characterization of homomultimeric β2 channels

Our previous results have shown that β1 and β3 behaved like histamine-gated chloride

channels. In the literature, until now to our knowledge there is no report about the

successful expression of β2 homomeric channels. We decided to re-examine first the

result of injecting β2 alone into Xenopus oocytes. Unexpectedly, the human β2 cRNA

expressed histamine-gated chloride channels, suggesting the formation of

oligohomomeric β2 receptors.

6.1: Homomeric β2 receptors behave like histamine-gated ion channels Fig. 6.1A shows the histamine generated membrane currents in oocytes injected with β2

cRNA. Histamine generated substantial membrane currents that resembled closely the

currents generated by oocytes injected with β3 subunit. The currents generated by low

concentration of histamine applied to β2 injected oocytes were fairly well maintained; but

the current elicited by high concentrations desensitized rapidly and recovered well after

washing. The histamine currents of β2 injected oocytes were fast desenzitising and

occasionally had amplitude of > 1.5 µA with 3 mM histamine. The histamine dose-

current amplitude relationships were fitted well by Hill equation with a resulting EC50 of

280 ± 2.1 µM, (n = 7), (Fig. 6.1C). The effect of histidine, the histamine metabolite, was

also checked on β2 homomultimeric channels. β2 homomultimeric channels were

activated by histidine in a dose dependent manner. Also histidine-evoked currents had

amplitude > 2.5 µA with 3 mM histidine. Fig. 6.1B shows the dose-response curve where

the induction of histidine currents start from 10 µM and reached to saturation at 10 mM

histidine. The histidine current had EC50 of 480 ± 5.8 µM, (n = 7). Thus, our results show

that β2 can form homomultimeric channels when injected into oocytes, which are

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activated, by histamine and histidine. t-MHA directly activated inward currents in a

concentrationdependent manner over a concentration range of 0.1 - 30 mM (Fig. 6.2).

The efficacy of t-MHA at these homomultimeric receptors was quite less compared to

histamine, as t-MHA induced currents of very less amplitude. 6.2: Histamine-gated homomultimeric β2 receptors behave like typical ligand-gated chloride channels

To identify the ion permeability through the receptor channel, the current reversal

potential was determined while activating the receptors with histamine. The voltage-

current relation indicates a rectifying channel with a reversal potential around -20 mV

(Fig. 6.3), as expected for chloride ions in Xenopus oocytes using Ringer as external

solution.

6.3: Relative comparison of various agonist of histamine with GABA

It has been controversial whether or not β2 homomultimers can form functional GABA-

gated ion channels. In one report it has been shown that both β1 and β3 can form

homomultimeric channels (Miko et al., 2004; Sigel et al., 1989; Krishek et al., 1997). To

address the same question for the β2 subunit, we further examined the β2 homomers. Fig.

6.4A and B shows the relative comparison of histamine and its metabolites with GABA.

At 10 mM saturating concentration histidine-evoked current was 8 folds higher than 3

mM GABA.

6.4: Pharmacological characterization of histamine-gated homomultimeric β2 receptors Thioperamide, H3 receptor antagonist, inhibited histamine-gated ion channels formed by

β2 receptors in a concentration-dependent manner over a range of 1-60 µM (Fig. 6.5A)

The IC50 value of thioperamide concentration-response curve for homomultimeric β2

subunit were 28.4 ± 1.2 µM, (n = 5). At 100 µM thioperamide showed outward current,

indicating the blockage of spontaneously activated channel-current. Next, H2 receptor

antagonist, famotidine was investigated for its activity on β2 subunit receptors.

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Fig.6.1: Homomultimeric channels of β2 subunit and effect of histamine and histidine. (A, B) The dose-response relationship for histamine and histidine from oocyte expressing homomultimeric β2 receptors. Bars over current traces indicate the duration of Histamine or histidine application with the concentration of applied agonist in µM. The traces in A and B were obtained from the same oocytes. The holding potential was -40 mV. (C) Concentration-response curve for histamine (closed circle) and histidine (open circle) for homomeric β3 receptors (n = 7). Data points shown are the means of multiple normalized experiments.

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Fig. 6.2: Dose-response curve of t-MHA for homomultimeric β2 receptors. Oocytes were voltage-clamped (-50 mV) 3-6 days after injection of in vitro transcribed cRNA encoding β2 subunit of GABAA receptors. Various concentration of t-MHA applied for 10 seconds produced inward currents. Bars indicate the duration of application of agonist.

Fig. 6.3: Current-voltage relationship of the current activated by histamine in an oocyte injected with β2 subunit of GABAA receptors. Points indicate peak currents elicited by 500 µM histamine applied with membrane potential clamped at different levels.

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Fig. 6.4: Relative comparison of current evoked by histamine and its metabolites with GABA. (A) The current-traces for GABA, histamine, histidine and t-MHA applied on the oocyte expressing homomultimeric β3 receptors. Saturation concentrations were taken for each agonist. (B) Average percentage activation of homomultimeric β2 receptors by histamine and its metabolites (n = 3).

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Famotidine inhibited histamine current with an IC50 of 70.5 ± 1.8 µM, (n = 3). At 300µM

concentration it also blocked the spontaneously active currents (Fig. 6.5B). The dose-

response curve was fitted with hill equation (Fig. 6.5C). Thioperamide is a more potent

antagonist than famotidine on β2 receptors. Pyrilamine, H1 receptor antagonist, at a

concentration up to 300 µM did not affect current evoked by 500 µM Histamine. The

affinity and efficacy of thioperamide block were much greater than the famotidine.

Control β2 currents evoked by 500 µM histamine were inhibited in concentration-

dependent manner by HTMT (Fig. 6.6A). HTMT maximally inhibited the histamine-

activated current, with an IC50 of 100 ± 1.2 µM, (n = 3).

Fig. 6.6B is a plot of sub maximal histamine-activated currents for β2 homomultimeric

channels and effect of harmane on it. Harmane significantly reduced the half maximum

histamine-current amplitude in a dose-dependent manner. The IC50 value for harmane is

36 ± 1.6 µM, (n = 3). Also, at high concentration 300 µM harmane blocks the

spontaneously open channels formed by β2 subunit.

6.5: Relative comparison of homomultimeric β2 and β3 receptors

Fig. 6.7 shows the relative comparison of β2 and β3 homomultimeric channels by their

pharmacology. β3 is 3 times more sensitive to thioperamide for histamine-evoked current

than β2. Also for HTMT, β2 was 2 folds more sensitive than β3. Famotidine, H2

antagonist, however, blocked histamine-evoked current with same efficacy on β2 and β3

homomultimeric receptors. Only harmane, was found to be more sensitive on β2 than β3

ranging up to 2-3 folds higher sensitivity.

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Fig. 6.5: Dose-dependent inhibition by thioperamide and famotidine of histamine-activated current in homomeric β2 receptors. (A, B) The dose-response inhibition of histamine-activated currents by thioperamide and famotidine from oocyte expressing homomultimeric β2 receptors. Bars over current traces indicate the duration of application of 500 µM histamine and histamine plus thioperamide \ famotidine in µM concentration. The holding potential was -40 mV. (C) Average dose-inhibition curve for famotidine (open circle) and thioperamide (closed circle) on histamine-activated current respectively (n = 3). Data points shown are means of normalized experiments and curve was fitted using Hill equation.

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Fig. 6.6: Dose-dependent inhibition by HTMT and harmane of histamine-activated current in homomultimeric β2 receptors. (A, B) The dose-response inhibition of histamine-activated currents by HTMT and harmane from oocyte expressing homomultimeric β2 receptors. Bars over current traces indicate the duration of application of 500 µM histamine and histamine plus HTMT \ harmane in µM concentration. The holding potential was -40 mV. (C) Average dose-inhibition curve for HTMT (open circle) and harmane (closed circle) on histamine-activated current respectively (n = 3). Data points shown are means of normalized experiments and curve was fitted using Hill equation

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Agonist β2 β3

Histamine 280 ± 2.1 µM 597 ± 5.1µM

Histidine 480 ± 5.8 µM 1135 ± 0.15 µM

Table 4: EC50 to Histamine and its metabolites on homomultimeric β2 and β3 receptors

Antagonist β2 β3

Famotidine 70.5 ± 1.8 µM 77.1 ± 8.9 µM

Thioperamide 28.4 ± 1.2 µM 9.7 ± 0.8 µM

HTMT 100 ± 1 µM 48.7 ± 2.5 µM

Harmane 36 ± 1.6 µM 82.7 ± 6.3 µM

Table 5: EC50 of various antagonists on homomultimeric β2 and β3 receptors.

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Fig. 6.7: Relative comparison of homomultimeric β2 and β3 receptors. (A and B) Bar diagram showing relative comparison based on pharmacology of various antagonists and EC50, respectively (β2 blue bar, β3 purple bar).

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Summary of major findings:

1. Homomultimeric β2 subunit forms histamine-gated ion channel with an EC50 of 280 µM.

2. Homomultimeric β2 receptors are also activated by histidine (EC50 0.48 mM) and t-

MHA. 3. Metabotropic histamine H3 receptor antagonist thioperamide inhibits histamine-

evoked current with an IC50 of 28.4 µM.

4. Histamine evoked current can be inhibited by famotidine (H2 receptor antagonist), harmane and HTMT with a relative order of harmane > famotidine > HTMT.

5. Homomultimeric β2 receptors are more sensitive to histamine than β3 receptors.

6. Pharmacologically homomultimeric β2 and β3 receptors are quite distinct to each

other.

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Chapter 7 7.1: Characterization of ε-subunit containing receptors

The ε-subunit of the GABAA receptor was independently cloned and functionally

characterized in recombinant expression systems by two groups (Davies et al., 1997;

Whiting et al., 1997). Both groups showed that co-expression of alpha beta epsilon

subunits produced functional receptors; however the sensitivity of these receptors to the

potentiating effects of general anesthetic agents differed. In human and rat tissues,

complex patterns of transcripts are derived from the genes that encode the gamma-

aminobutyric acid GABAA receptor epsilon subunit. A ε subunit transcript

(approximately 3.6 kb) is expressed at relatively high levels in regions of the human brain

and heart, but is not detected in most other major tissues. The ε subunit has greater

amino-acid sequence similarity to the γ subunits than to any other classes of GABAA

receptor polypeptides. Furthermore, the ε and γ subunits both require the presence of α

and β subunits to incorporate within functional receptors (Davies et al., 1997). Therefore

it was hypothesized that ε and γ subunits compete for the common site in the GABA

receptor complex. It has demonstrated that expression of ε subunit does indeed modify

the functional properties of α1β3γ2 receptors stably expressed in WSS-1 cells (Davies et

al., 1997). Presence of ε subunit in GABAA receptor complex causes the spontaneous

current activity. Moreover, our initial experiments in bioinformatics revealed some

distinctive sequence homology characteristic for histamine-gated chloride channels. We

decided to examine the properties of ε subunits in Xenopus oocytes.

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7.2: ε-containing receptors behave like histamine-gated ion channels It was noted that oocytes injected with ε subunit cRNAs required an unusually large

holding current to maintain a potential of -50 mV and the resulting baseline value was

relatively unstable. We injected different subunit combinations to check the effect of ε

subunit on receptor complex.

Oocytes co-expressing α1β1ε subtypes were activated by histamine (30 - 10,000 µM) in a

concentration-dependent manner (Fig. 7.1A). The α1β1ε had histamine EC50 of 3 mM,

(n = 9). We also observed that α1β1ε subunit combination was highly sensitive to GABA.

The comparison of GABA-evoked current is also shown in Fig. 7.1A.

Oocytes injected with β1ε subunits cRNA did not give any response even at the 1 mM

GABA concentration. It is very different from the α1β1ε receptor where even 1 µM

GABA concentration gives large current response. As α subunit is responsible for the

binding of GABA to GABAA receptor complex we concluded it as logical for no

response to GABA. But what was quite interesting also here that β1ε subunit combination

formed highly spontaneously active channel which were activated by histamine in a dose

dependent manner Fig. 7.1B.

In case of oocyte injected with ε subunit cRNA showed spontaneous current in the

absence of any agonist. Oocyte clamped at -50 mV usually had leakage current up to 1

µA. Interestingly; also application of histamine activated the ε subunit in a dose-response

manner (Fig. 7.1C). When GABA was applied on homomeric ε receptors it blocked the

spontaneously opened channel. From all our experiments, we concluded that ε was also a

similar histamine-gated ion channel.

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Fig 7.1: Dose-response relationship for histamine and histidine on homo and heteromultimeric channels formed by ε subunit of GABAA receptors. (A, B and C) Dose-response curve to different concentrations of histamine (100 µM – 10 mM) for α1β1ε, β1ε, ε. Left side in (A) activation of α1β1ε by GABA. Excluding α1β1ε, β1ε and ε subunits combination are not activated by GABA. All of the oocytes were voltage-clamped at -50 mV and each trace is representative of observation made from four to five determinations.

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7.3: Pharmacology of ε subunit To characterize the channel pharmacologically various antagonists for metabotropic

histamine receptors were checked. As the EC50 of ε is nearly 3 mM, so in our further

experiments we always took 3 mM histamine. We found that cimetidine - H2 receptor

antagonist - inhibited histamine-evoked current potently. It inhibits the histamine-evoked

current in a dose dependent manner with an IC50 of 3-5 µM (Fig 7.2A). We checked the

H1 receptor antagonist pyrilamine and observed that it was less sensitive antagonist.

Pyrilamine inhibited histamine-evoked current at IC50 value of 300-400 µM.

Thioperamide blocked the histamine-evoked current with an IC50 of 3 µM. The

histamine-evoked current was blocked in highly selective manner by PTX, a chloride

channel blocker, at nM concentration.

Fig. 7.2: Pharmacological characterization of homomeric ε subunit receptors. (A and B) Dose-response inhibition by cimetidine and pyrilamine. Cimetidine (H2-receptor antagonist) inhibited histamine-evoked current more effectively than pyrilamine (H1-receptor antagonist). All of the oocytes were voltage clamped at -50 mV and each trace is representative of observation made from four to five determinations.

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7.4: Expression of α1β1ε, β1ε and ε in HEK 293 cells

To check that the high EC50 value of histamine for homomultimeric ε subunit can be

reduced in different cell culture system, we decided to functionally express and

characterize ε subunit in HEK 293 cells. As we found other two subunit combinations

α1β1ε, β1ε were also sensitive to histamine, we transfected HEK 293 cells with α1β1ε, β1ε

as well as ε subunit cDNAs. First of all, we checked α1β1ε subunit combinations, which

like in oocytes were highly sensitive for GABA. Interestingly, we found histamine did

not show any response even up to 3 mM concentration. What was evident that we did not

observe any spontaneous channel activity and channels were highly stable with almost no

leakage current, which was in contradiction with other ε subunit combination (for e.g.

α1β3ε) by other research group. We also checked β1ε and ε receptors in HEK 293 cells

but never found any response for histamine or GABA. Interestingly, there was also no

leakage current in any of the combination tested. We were curious for this anomalous

behaviour of the receptor and asked if ε subunit is really transported to the cell membrane

of the HEK 293 cells as it is documented in several reports that the single subunits are

retained in the endoplasmic reticulum of the cell and are not transported to the cell

membrane (Bollan et al., 2003; Gorrie et al., 1997). The reason and explanation is

already being discussed in the introduction section.

7.5: Molecular cloning of GFP-tagged subunits of GABAA receptors

The negative results obtained in the HEK 293 cells transfected with ε subunit alone, lead

us to check first the expression of ε subunit at the cell membrane. We addressed the

question, does ε able to be transported to the cell membrane or not. For this we tagged

ε subunit with GFP as well as to further get more information for the transportation of the

subunits we decided to tag α1 and β1 subunits of GABAA receptor with GFP.

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7.5.1: Molecular cloning of GFP- tagged α1 and β1 subunits of GABAA receptors

In the next set of cloning steps pCDNA3 vectors were cloned that allowed the expression

of GABA receptors fused with a GFP on the C-terminal end. In the first step, the open

reading frame of the rat GABA α1 and β1 cDNA was PCR amplified using the plasmid

pSGEM-rat-α1 / β1 as templates (3.2.2). Primer pairs used were rGA-α1-4-stop codon,

GABA α1-Not1-Stop-Ev and RR-GABA-β2-up, RR-GABA-β2-do for α1 or β2,

respectively (2.3). In the corresponding 'down' primer, an EcoRV site was inserted

upstream of the stop codon that served for later insertion of the GFP fragment. After gel-

purification (3.1.2 and 3.1.5) the PCR fragments were cut by EcoRI and NotI (flanking

sites present in the used PCR primers for cloning) and ligated into the EcoRI and NotI cut

pCDNA3 vector (3.3.5). From clones with the correct insert, plasmid was prepared

(3.1.8) and the insert verified by sequencing (3.3.8). The resulting plasmids were named

pCDNA3-α1-EcoRV-stop and pCDNA3-β1-EcoRV-stop. pCDNA3-GFP was digested by

enzymes EcoRV and HpaI to take out the GFP open reading frame. The insert isolated

from this plasmid consists of the entire GFP open reading frame without start and stop

codon. The plasmids pCDNA3-α1-EcoRV-stop and pCDNA3-β1-EcoRV-stop were cut

by restriction enzyme EcoRV (3.3.1) and dephosphorylated (3.3.2). Restrction by EcoRV

caused the linearization of plasmids with blunt ends. These blunt-end plasmids could be

ligated with the EcoRV- HpaI-GFP fragment as HpaI generates blunt ends, too. From

clones with the correct insert, plasmid was prepared (3.1.8) and the sequence and

orientation of the fused GFP verified by sequencing (3.3.8). The resulting plasmids were

named pCDNA3-α1-N-GFP and pCDNA3-β1-N-GFP. Fig. 7.3 shows the schematic

representation of GFP tagged α1 subunit of GABAA receptor. For the GFP tagged β1

subunit the same procedure was employed.

7.5.2: Molecular cloning of GFP-tagged ε subunit of GABAA receptor

To tag the ε subunit C-terminal with GFP we employed different cloning strategy which

required 3 sub cloning steps. As ε subunit contains EcoRV, restriction site cloning could

not be done like in α1 and β1-GFP tagged experiment.

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Fig. 7.3: Cloning of GFP-tagged α1 subunit of GABAA receptors. The open reading frame of the rat GABA α1 was PCR amplified using the plasmid pSGEM-rat-α1 as templates. Primer pairs used were rGA-α1-4-stop codon, GABA α1-Not1-Stop-Ev. In the corresponding 'down' primer, an EcoRV site was inserted upstream of the stop codon that served for later insertion of the GFP fragment. After gel-purification the PCR fragments were cut by EcoRI and NotI and ligated into the EcoRI and NotI cut pCDNA3 vector. The resulting plasmid was named pCDNA3-α1-EcoRV. To take out the GFP open reading frame from pcDNA3-GFP, it was digested by enzymes EcoRV and HpaI. The plasmids pCDNA3-α1-EcoRV-stop was cut by restriction enzyme EcoRV (3.3.1) and dephosphorylated.These blunt-end plasmids were ligated with the EcoRV- HpaI-GFP fragment. The resulting plasmids were named pCDNA3-α1-GFP.

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In first step, by rGABA ε-Not1-Stop–EV in conjuction with a primer-directed polymerase

chain reaction, restriction sites for EcoRV and NotI were introduced, before and after the

stop codon of ε subunit. The elongation conditions were as follows: 94o C for 2 min,

followed by 20 cycles at 94° C for 1min, 59º C for 1 min, and 72º C for 1 min, and

ending with an incubation at 72º C for 10 minutes.

The pCDNA3 vector was digested by restriction enzymes EcoRV whereas SalI restricted

PCR fragment. Fragments were separated using gel electrophoresis and purified using a

gel extraction kit (3.1.5). Chimeric fragments were ligated using the rapid ligation kit.

Purified plasmid DNA for transfection was made using the plasmid-maxi kit (3.1.8) and

the entire coding region was sequenced.

In the second step of cloning, to take out the GFP open reading frame from pCDNA3 /

GFP, the vector was digested by enzymes EcoRV and HpaI. The plasmids containing ε-

EcoRV-stop-NotI was cut by restriction enzyme EcoRV. This restriction of plasmid

produced two fragments large and small. This large fragment was ligated with GFP

fragment, cloned and sequenced.

In the third step of cloning, plasmid containing the large fragment of ε subunit with GFP

was digested by EcoRV which linearised the plasmid with blunt ends. In this linearised

plasmid, the small ε fragment was ligated as a result after the ligation we obtained the

complete reading frame of ε subunit tagged with GFP. Purified plasmid DNA for

transfection were made using the plasmid-maxi kit (3.1.8) and the entire coding region of

all mutant was sequenced using Big dye ready reaction mix and an ABI automated DNA

sequencer. Fig. 7.3A and B shows the schematic representation of cloning strategy for

GFP tagged ε subunit. Fig. 7.4A and B shows the schematic representation of the GFP-

tagged ε subunit of GABAA receptors.

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Fig. 7.4 (A): Cloning of GFP-tagged ε subunit of GABAA receptors. In rGABA ε-Not1-Stop–EV in conjunction with a primer-directed polymerase chain reaction, restriction sites for EcoRV and NotI were introduced, before and after the stop codon of ε subunit. The pCDNA3 vector was digested by restriction enzymes EcoRV whereas PCR fragment was restricted by SalI. Fragments were separated using gel electrophoresis. The plasmids containing ε-EcoRV-stop-NotI was cut by restriction enzyme EcoRV.

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Fig. 7.4 (B): Cloning of GFP-tagged ε subunit of GABAA receptors. In the second step of cloning, to take out the GFP open reading frame from pCDNA3 / GFP, the vector was digested by enzymes EcoRV and HpaI. The plasmid containing ε-EcoRV-stop-NotI was cut by restriction enzyme EcoRV. This restriction of plasmid produced two fragments, large and small. This large fragment was ligated with GFP fragment, cloned and sequenced plasmid containing the large fragment of ε subunit with GFP was digested by EcoRV which linearized the plasmid with blunt ends. In this linearized plasmid, the small ε fragment was ligated as a result after the ligation we obtained the complete reading frame of ε subunit tagged with GFP.

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7.6: Expression of α1, β1 and ε in HEK 293 cells HEK 293 cells were transfected with GFP-tagged ε subunit and GFP-tagged α1β1

combinations localization of the ε subunit was observed under the fluorescence

microscope. Fig. 7.5 shows the localization of the α1β1 subunit in the HEK 293 cells.

The uniform distribution of fluorescence on the cell membrane shows the transportation

of the α1β1 subunit combinations. However, HEK cells transfected with GFP-tagged ε

subunit showed no fluorescence on the cell membrane, Fig. 7.6. It is evident from

microscopic study that ε subunit is retained in the cytoplasm and not transported to the

plasma membrane. However, we also observed small fluorescence near the cell

membrane in a vesicular form but did not find the expression of ε on cell membrane.

HEK 293 cells transfected with GFP-tagged ε subunit were also characterized by whole-

cell Patch clamp technique. Again, in this case we can not measure any response to

GABA and histamine which confirms our result from microscopy that ε is retained in the

cytoplasm of HEK 293 cells and do not form any homomultimeric channels in HEK 293

cells when transfected alone.

Fig. 7.5: Surface expression of heteromultimeric α1β1 receptor of GABAA receptors in HEK 293 cell. Expression was determined using fluorescence of GFP-tagged α1 and β1 subunits of GABAA receptors. Images were collected by fluorescence microscopy. The uniform distribution of green fluorescence indicates the co-localization of the α1β1 receptors at cell membrane of HEK 293 cells.

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Fig. 7.6: Surface expression of homomultimeric ε receptor of GABAA receptors in HEK 293 cells. Expression was determined using fluorescence of GFP-tagged ε subunit of GABAA receptors. Images were collected by fluorescence microscopy. The lack of any fluorescence on the cell membrane shows that the ε subunit in not transported to the cell membrane. Most of the fluorescence is visible in the cytoplasm of the HEK 293 cells.

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Summary of Major Findings

1. Homomultimeric ε subunit forms histamine-gated ion channel with an EC50 of 3 mM histamine when injected in Xenopus oocytes.

2. Incorporation of different subunits with ε reduces the spontaneously opened channel frequency of the receptors. The order of decreasing spontaneous channel activity in our subunit combinations checked was ε > β1ε > α1β1ε.

3. Histamine evoked current can be inhibited by pyrilamine, cimetidine,

thioperamide. The increasing order of sensitivity of antagonist was pyrilamine < thioperamide < cimetidine.

4. In HEK 293 cells the ε subunit is not transported to the cell membrane.

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Chapter 8 8.1: Properties of α1β2γ2 receptors and direct activation by histamine As described in (4.8) α1β2γ2 composed receptors the GABA response was potentiated by

histamine. In contrast to the findings at α1β1 receptors, histamine alone (without GABA)

was activating small currents in α1β2γ2 receptors when expressed in Xenopus oocytes. In

most tested oocytes, these current evoked by high concentrations of histamine (10 - 30

mM) were rather small reaching typically 5-10 % of the maximally current evoked by

saturating concentrations of GABA (Fig. 8.1A). In some extreme instances, they were

reaching up to 35 % of the maximal GABA-induced currents (Fig. 8.1B). In few cases,

histamine-induced currents were detected in other subunit combinations like α1β1 also,

but in these few cases such currents were detectable at all, they had typical currents as

low as 1-3 % of the GABA induced currents only.

It was tested, whether such histamine-activated currents were detectable in other

expression systems also. Therefore, α1β2γ2 receptors were expressed in HEK 293 cells

and transfected cells were exposed to GABA and histamine (Fig. 8.1C). At a holding

potential of -50 mV, GABA induced a desensitizing inward current that was not blocked

by 20 µM Zn2+, a proof, that the cell expressed α1β2γ2 receptors because α1β2 receptors

would have been blocked under these conditions. Application of 3 mM histamine to the

same individual cell didn't induce any detectable current (Fig. 8.1C). My findings that

histamine didn't directly activate heteromultimeric GABA receptors expressed in HEK

293 cells was supported by the experiments performed by Angela Vogt-Eisele and Katja

Erlkamp, who found that histamine also evoked no detectable current in HEK 293 cells

expressing α1β1 (personal communication).

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What could be the reason for the pronounced histamine activated current found in the

oocyte expression system? As it is described later, homomultimeric channels composed

of β subunits behave like histamine-activated channels (5.1.7). Though the formation of

heteromultimeric channels composed of α-β-γ subunits is strongly favoured, there is a

high chance of getting also homomultimeric β or γ channels if the pool of available

channel subunits is extremely unbalanced. Recently it was shown that GABA-gated γ2

homumultimeric channel could also be functionally expressed in Xenopus oocytes

(Ataulfo et al., 2004). Therefore, it was tested whether histamine might activate those γ2

channels also and indeed it was found, that 10 mM histamine activated currents in

Xenopus oocytes expressing γ2 subunits. So, in oocytes injected with a combination of α,

β, and γ subunits we have at least two possible channel populations, namely β and γ

homomultimeric channels that are known to be activated by histamine. The variation in

the amount of histamine-induced current might reflect different amounts of

'contaminating' homomultimeric β or γ channels. The reason that there are no histamine-

evoked currents detectable in α1β2γ2 expressing HEK 293 cells lies in the fact that

homomultimeric β2 channels are not targeted to the cell membrane, also the expression of

homomultimeric γ2 receptors was never reported in HEK 293 cells but only in Xenopus

oocytes. Therefore, possibly, that the level of potential homomultimeric channels is much

lower or not even existing in HEK 293 cells and, as consequence, there is also no current

evocable by histamine in contrast to Xenopus oocytes.

For the next experiment it was aimed to check whether the nature of action of histamine

varies with its concentration. At low histamine concentrations of 0.3 mM histamine

activated nearly no currents but had a clear potentiating effect on the current evoked by 2

µM GABA (Fig. 8.1E). At higher concentrations of 10 mM histamine, the effect of the

direct activation predominates and the current evoked by the mix of (10 mM histamine +

2 µM GABA) seems to be the simple summation of the current evoked by 10 mM

histamine and 2 µM GABA alone (Fig. 8.1E). In this oocyte it seems that the potentiating

effect of histamine at low concentration is replaced by a direct activation of the receptor

at high concentrations of histamine. This fits to the finding that potentiating agents like

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propofol also show potentiating effects at low and direct activating effects at high

concentrations.

The last experiment shows that we can not rule out that at least a part of the histamine-

evoked current in Xenopus oocytes is mediated by α1β2γ2 receptors oocytes directly and

not by possible heteromultimeric β and γ channels.

In the next set of experiments, the influence of the GABA concentration was investigated

in greater detail. For the potentiation experiments, we used rather low concentrations of 1

mM histamine to avoid the direct activation of currents, further; oocytes were chosen that

had a small fraction histamine activated currents (> 5 % of currents evoked by 2 µM

GABA) only. In these oocytes (n = 3), histamine potentiated the current evoked by 2 µM

GABA up to 2.5 folds but the potentiation decreased with higher GABA concentrations

and no potentiation was detectable at 100 µM GABA. This behaviour fits to the idea that

1 mM histamine lowers the EC50 for GABA in these oocytes. All investigated oocytes

described in this chapter were verified to express α1β2γ2 receptors due to the absence of

the block with 10 - 30 µM Zn2+ (Fig. 8.1G).

8.2: Homomultimeric γ2 receptors: Activation by histamine and its metabolites We also checked the expression of homomeric γ2 subunit receptors. Oocytes injected with

γ2 showed activation by histamine. Comparing to saturating concentration of 3 mM

GABA, current evoked by 10 mM histamine was 1/2 fold smaller. We also found that

like homomeric β subunit receptor histidine and t-MHA activated the homomeric

γ2 subunit receptors. The relative order of current amplitude is histidine > histamine > t-

MHA.

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Fig.8.1: Anomalous behaviour of α1β2γ2 receptors. (A) In some oocytes histamine alone activated current in α1β2γ2 receptors. These current evoked by histamine (10-30 mM) were 5- 10 % of the maximally current evoked by saturation concentration of GABA. (B) In some extreme cases, histamine-evoked current reached to 35 % of the maximal GABA-evoked current. (C) In HEK 293 cells, GABA-induced current was not blocked by 20 µM Zn2+, confirming that the cell expressed the α1β2γ2 receptors. Application of 3mM histamine did not induce any detectable current. (D) In α1β1 receptors histamine alone did not induce any current (E) The nature of histamine activity varies with its concentration. At EC10 GABA, low concentration of histamine did not induce any current and show only potentiating effect but higher concentrations shows the direct activation. (F) Like α1β1 at saturating concentration of GABA histamine-evoked potentiation is maximum. (G) All investigated oocytes were verified to express α1β2γ2 receptors due to the absence of the block with Zn2+.

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Fig. 8.2: Activation of homomultimeric γ2 receptors by GABA, histamine and its metabolites. Oocytes injected with γ2 subunit alone showed activation by histamine and its metabolites. Compared to saturating concentration of 3 mM GABA, current evoked by 10 mM histamine was nearly 1 \ 2 fold smaller. The relative order of amplitude of histamine and its metabolites was histidine > histamine > t-MHA. Bars over current traces indicate the duration of applied agonist in mM concentration.

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Chapter 9 Discussion We were interested to find the histamine-gated ion channel in vertebrates. Two major

documentations, first the identification of histamine-gated ion channel in Drosophila

melanogaster (Gisselmann et al., 2002) and second electrophysiological evidence for a

possible histamine-gated ion channel in the SON nucleus (Hatton and Yang, 2001), lead

us to search for the gene of histamine-gated ion channels in mammals. A systematic

scanning of various vertebrate genome and EST databases from zebra fish to human

using the sequence information of newly identified Drosophila HisClα1 and HisClα2

genes and looking for various characteristic features of ligand-gated ion channels like

signal peptides, binding and transmembrane domains did not identify any novel gene for

ligand-gated ion channels having similarity with known histamine-gated ion channels.

However, histamine-gated channels from insects showed some homology with known

GABAA receptors subunits as well as some homology to glycine receptors also.

We found the homology to GABAA receptors interesting because of the various reasons.

The histamine containing neurons are found exclusively in the TM nucleus of the

hypothalamus in the brain (Bekkers, 1993). GABA containing neurons in the

ventrolateral preoptic nucleus innervates the TM nucleus and form symmetrical synapses

with histaminergic neurons. The GABA synthesizing enzyme, glutamate acid

decarboxylase (GAD) and GABA are also found in most TM neurons (Vincet et al.,

1983; Takeda et al., 1984; Senba et al., 1987). Interestingly, many neurons in TM

nucleus contain both GABA and histamine. In invertebrates, GABA and histamine can

act at the same ionotropic histamine receptor (Gisselmann et al., 2004). Therefore, we

supposed that histamine was candidate for acting on vertebrate GABAA receptor also. To

prove our hypothesis, we investigated the action of histamine on recombinant and native

GABAA receptors. Various chemical agents like benzodiazepines, barbiturates, steroids,

convulsants and anesthetics modulate GABAA receptors (Sigel et al., 2002; Boileau et

al., 1999; Rick et al., 1998; Olsen et al., 1997; Kraswoski et al., 1998). Modulation of

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151

GABAA receptors influences several functions such as epilepsy; sleep disorders, anxiety

and pain (Kash et al., 1997; Delory et al., 1988; McKernan et al., 2000; Nelson et al.,

2002).

Histamine potentiates GABA-evoked currents in GABAA receptors We found that GABAA receptors are potentiated by histamine. The currents elicited by 2

µM GABA (a concentration near EC10 for α1β1 receptors) was potentiated by histamine

up to 3-4 folds. Potentiation by histamine does not affect the permeability of the ion

across the channel pore. This suggests that general property of the channel is not altered

in the presence of the histamine. The M2 pore-forming domain determines ion selectivity

of ligand-gated ion channels. The M2 domain possesses three rings of charge called the

cytoplasmic, intermediate and extracellular charged domains (Imoto et al., 1988). The

cytoplasmic ring is negative in all ligand-gated ion channels, whereas the intermediate

and extracellular rings are negative in cation-selective and positive in anion-selective

channels (Karlin and Akabas, 1995). The intermediate ring of charge is located near the

intracellular mouth of the channel, where the pore is most constricted, and is thought to

provide the electrostatic interaction for cation or anion selectivity (Imoto et al., 1988;

Lester, 1992; Wilson and Karlin, 1998; Xu and Akabas, 1996; Keramidas et al., 2002).

As the channel formed by the α1β1 subunit combinations has the intermediate and

extracellular ring which is positive charged so no alteration in the permeability is logical

during the potentiation mediated by the histamine.

On the level of a single GABAA channel, the increase in the amplitude of GABA-evoked

current by histamine may arise by several possibilities, like increase in the frequency of

open probability of the channel, increase in open time period and alteration of the single

channel conductance. In oocytes expressing α1β1 GABAA receptors the potentiation of

high concentrations of histamine persisted even at high, saturating GABA concentrations

and histamine potentiated the GABA-evoked current beyond the maximal current evoked

by GABA. In those oocytes showing a robust potentiation by histamine at 10 µM GABA,

no affinity change for GABA in the presence of histamine occurs, pointing out that the

overall the tendency of the receptor to open at various concentrations of GABA is the

Chapter 9 Discussion

152

same which suggests that the potentiation by histamine does not change the open

probability of the receptor. It may alter the open frequency, the open time duration or the

conductance of the channel, but future work has to be done on single channels to find out

the nature of histamine action. In virtually all oocytes expressing αβ subunit

combinations where the effect was systematically tested, GABA-potentiation was

independent on the GABA concentration (except for some few rare oocytes where

histamine didn't potentiate well. In these oocytes, histamine had a detectable effect at low

GABA concentrations only). This was different in oocytes expressing α1β2γ2 receptors.

Here, the potentiating effect was higher at lower GABA-concentrations and the nearly

vanished at high, saturating GABA concentrations. This is consistent with the idea that

histamine affects the GABA affinity and shifts the dose response curve to the left site.

The direct histamine-activated current component prevented a reliable quantitative

analysis of the of the EC50 shift so far that therefore remains to be determine in future but

the fact itself could be clearly demonstrated. Our findings suggest that γ2 subunits

influence the potentiation by histamine in heteromultimeric channels. This is not

unexpected as γ2 subunits also alter the effect of other potentiating agents like

benzodiazepines, too. In addition, we demonstrated that γ2 subunits have a histamine-

binding site itself as indicated by the fact that the corresponding γ2 homomultimeric

channels are gated by histamine.

Different subunit combinations of GABAA receptors are potentiated by histamine;

however they exhibit different sensitivity to histamine potentiation. For example

α1β3 subunit combinations were found to be very sensitive to histamine. α1β2, though

potentiated by histamine, showed higher concentration dependability for histamine. This

is also consistent with other reports, (Walters et al., 2000) showed the biphasic

potentiation of α1β2γ2 receptors by diazepam. At nM concentrations, diazepam

potentiated α1β2γ2 receptors, which leads to saturation at 1 µM. On the other hand, µM

concentration of diazepam (20 µM and above) evoked a second component of

potentiation, further increasing GABA-elicited current from 3 fold (the nM component)

to approximately eight folds. In contrast, α1β2 receptors show monophasic potentiation.

Chapter 9 Discussion

153

Diazepam elicited only a single component of potentiation with in the µM concentration

range for α1β2-receptor channel. Also the lack of α1β2 receptor channel sensitivity to

diazepam with in the nM concentration range has been documented previously, as the

presence of γ subunit is essential in conferring diazepam nM action (Pritchett et al.,

1989).

Experiments done in HEK 293 cells clearly show that the observed effect was not an

artifact of the Xenopus oocyte system and can also be observed in HEK 293 cells.

Potentiation mediated by histamine shows the same degree of potentiation in different

cell culture system. Our experiments in HEK 293 cells reveal that the degree of

potentiation mediated by histamine remains unchanged. Histamine showed similar

potentiating effects as in oocytes for e.g. at 2 µM GABA, 1 mM histamine evoked an up

to 5-folds potentiation of the GABA current (2.9 folds potentiation on an average) which

is in accordance with the potentiation in oocyte where we observed an up to 5 folds

potentiation. Also histamine-mediated potentiation of GABAA receptors is applicable to

bulbous neurons, which indicate that not in artificial recombinant system but in also

native neurons this effect can be reproduced.

We observed on α1β2γ2 receptors a considerable direct activation by high concentrations

of histamine in some batches of oocytes. The relative amplitude of the histamine-evoked

current to the maximum-evoked current of GABA or histamine potentiated GABA-

evoked current was typically up to 10 %. We found in HEK 293 cells that application of

histamine up to 3 mM does not show any activity. The direct activation by histamine can

be described like that in the α1β2γ2 receptors there is strong chance that a strong

expression of β or γ subunits can cause a considerable population of histamine-activated

homomultimeric receptors in addition to the expression of heteromeric receptors. As we

have clearly shown that both homomultimeric β and γ channels can be activated by

histamine alone, so the occurrence of histamine-activated currents in oocytes injected

with α1β2γ2 subunits should not be a surprise. But we cannot rule out the possibility that

at least a fraction of the observed histamine-activated current is carried by directly

activated heteromeric α1β2γ2 receptors.

Chapter 9 Discussion

154

We found that the metabolites in histamine pathways also potentiate GABA-evoked

current. L-histidine is a water-soluble amino acid. Histidine potentiated 2 µM GABA

current up to two folds. The in vivo actions of supplemental L-histidine are entirely

unclear. Our experiments give an idea that one of the possible mechanisms of histidine in

the body could be the interaction of histidine with GABAA receptors. The potentiation

mediated by histamine and histidine has intrinsic differences. First, histidine potentiates

GABAA receptors with lower EC50 than histamine in α1β1 receptors. The difference of

EC50 in between them is 1/10. Second, the amplitude of potentiation evoked by histamine

at 2 µM GABA concentration is higher (4-5 folds) than histidine (2 folds). Third, the

average fractional potential decreases for histamine from lower to higher GABA

concentration whereas in case of histidine it increases.

Histamine acts on GABAA β-subunits on the propofol binding site The subunit responsible for the histamine-binding site was determined by our subunit

substitution experiments and functional expression of homomultimeric channels of

GABAA receptors. Substitution experiments where we shuffled subtypes of one subunit

keeping another subunit constant, gave us an idea that β subunit is the possible candidate

for the histamine binding site. To get the clear indication for the subunit responsible for

histamine binding, we checked the homomultimeric channel expression of β subunit. It

has been well documented that homomultimeric β subunits can form functional channels

that either can open spontaneously or can be directly activated by some general anesthetic

(Sigel et al., 1990; Cestari et al., 1996; Krishek et al., 1996; Davies et al., 1997; Sanna et

al., 1995). However, whether or not homomeric β subunits can form functional GABA-

gated ion channels is still controversial and appears to depend on different species. For

instance, while human and bovine β receptors were found to form channels that can be

gated by GABA (Sanna et al., 1995; Blair et al., 1988; Pritchett et al., 1988; Krishek et

al., 1994) rat and mouse β1 and β3 subunits were insensitive to GABA (Sigel et al., 1990;

Krishek et al., 1996; Davies et al., 1997). We confirm by our experiments that rat β1

forms homomultimeric receptors, which are spontaneously active and can be blocked by

PTX. Our results are further extension for the view that β1 subunit can form

homomultimeric channel, which has already shown by (Miko et al., 2004). Although the

Chapter 9 Discussion

155

precise reason behind discrepancy in the results from different group is not clearly

understood but there are a number of possibilities that could attribute, in this discrepancy.

First, it can be because of different expression level of β1 subunit. Second, it may depend

on the level of posttranslational modulation of the subunits, which can vary pronouncedly

among different batches of Xenopus oocytes. Another possibility to reconcile this

discrepancy can be due to different levels of spontaneous activity of the β1 subunits

expressed in oocytes in different experimental conditions.

β subunit has the binding site for various modulators like pentobarbital and propofol. By

our pharmacological and electrophysiological results first we discriminated any

possibility of similarity between the pentobarbital and the histamine-binding site. That

was achieved by using antagonists for pentobarbital and histamine, respectively. First,

bemegride an antagonist of pentobarbital, inhibited pentobarbital-evoked current strongly

but did not have any influence on histamine-evoked currents. Second, thioperamide the

H3 antagonist competitively blocked the histamine-evoked current but was insensitive for

pentobarbital. Interestingly, in the same approach we found similarity in between

propofol and histamine binding site. We hypothesized that histamine should have the

binding site similar to propofol because of various reasons. Thioperamide inhibited

propofol-evoked current with same efficiency like its activity on histamine-evoked

current. Bemegride didn’t block propofol-evoked current, which was consistent with

earlier findings that pentobarbital and propofol have different binding sites. The

bemegride experiment was a clear indication to hypothesize that the different binding site

of different modulators need different antagonists. Having the same efficiency for the

same antagonist indicates that two different modulators can have the same binding site.

Moreover, in experiments where we mixed propofol and histamine to their EC50 and EC90

concentrations and co-applied them with 2 µM GABA on α1β1 and α5β1 receptors gave

another support for our hypothesis.

Propofol binding alters the GABAA receptor structure in the M3 membrane-spanning

segment region (Williams and Akabas, 2002). In several pioneering studies using

mutated recombinant receptors, several groups identified amino acid residue M286 the

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156

binding site for propofol on β subunit. (Krasowski et al., 1998) has shown that on α2β1

(M286W) mutant receptor, propofol concentration greater than 10 µM failed to enhance

sub maximal GABA-currents. In contrast to lack of the potentiation, propofol still

directly activates this mutant receptor, with a concentration-response relationship that

overlaps with that for the wild-type α2β receptors. Moreover, mutation of a GABA

binding site residue, β2W157, reduced direct activation but did not affect propofol´s

modulatory actions (Fukumi et al., 1999). These results suggest for propofol, potentiation

and direct activation may involve binding to distinct sites. Sub maximal GABA currents

at wild type α1β1 receptors were potentiated by propofol. Thus, in agreement with the

other published studies, the presence of γ subunit is not required for the potentiation by

propofol. Our mutagenesis result also strongly proves hypothesis that (M286W) residue

on β subunit is indeed the binding site for propofol. In α1β1 (M286) mutant receptors,

propofol did not enhance the propofol-evoked currents at the same anesthetic

concentration tested in wild type α1β receptors. However, this mutation leads to the

spontaneous activity in the α1β1 (M286W) mutant receptor, which can be illustrated by

large leakage current at -60 mV holding potentials.

Remarkably the mutation in α1β (M286W) mutant receptor abolished the potentiation

mediated by histamine and histidine. No potentiation of mutant receptor by histamine up

to 10 mM and histidine up to 1 mM concentration was detected which strongly suggest

that both have the same binding site like propofol. We checked the effect of mutation at

different GABA concentration from 2 µM to 1 mM GABA with various concentrations

of histamine and histidine and found that even varying the concentration so much does

not lead still any potentiation. It is interesting to note also that this mutation does not

affect the activity of other modulators on β subunits. Modulation by pentobarbital and

neurosteroid remained same in α1β (M286W) mutant receptors. These data, first, provide

functional correlates for the receptor binding data, demonstrating vanished efficacies in

anesthetic–induced enhancement of ligand binding with mutant α1β1 (M286W) receptors;

and second strongly support the hypothesis that this methionine residue at the TM3

domain is important for the allosteric actions of propofol, histamine as well as histidine.

Chapter 9 Discussion

157

It is somewhat perplexing that methionine substitution at the aligned β3 M2 position,

β3N265, virtually eliminates propofol´s anesthetic efficacy in a knock-in mouse (Jurd et

al., 2003). Methionine occupies a volume 43 Ao greater than aspargine and is more

hydrophobic. Thus, steric bulk at position 265 can alter propofol binding, perhaps by

inducing a conformational change at the propofol binding site ~ 10 Ao away. Moreover,

we show that GABAC receptors are not potentiated by histamine. This is in accordance

with the previous findings for the action of other modulators on GABAC receptors

(Bormann et al., 1995; Feigenspan et al., 1998). Ionotropic GABAC receptors composed

of ρ subunits are insensitive to benzodiazepines, anesthetic and barbiturates (Polenzani

et al., 1991; Shimada et al., 1992). So, histamine behaves somehow like other modulators

already characterized. Sequence alignment show that the amino acid present at 286

location on ρ1 subunit of GABAC receptor is tryptophan instead of methionine which

gives the reason for no potentiation by histamine on GABAC receptors.

Homomultimeric receptors composed of β subunits are histamine-gated channels Our experiments on the homomultimeric β1 and β3 receptors support further the idea that

homomultimeric β and β3 receptors can be functionally expressed at least in Xenopus

oocytes forming histamine-gated channels. We contradict that β3 subunits do not form

spontaneously active channels shown by (Miko et al., 2004). We used human and they

used rat β3 subunits, this can be a plausible reason for having spontaneous channel

activity we observed in our experiments. Human homomultimeric β3 receptor is highly

sensitive for PTX, which blocks the open channel with in nM concentration.

Homomultimeric β3 receptors exhibit profound differences in the activity of histamine

and histidine. First, histamine activates homomultimeric β3 receptors with less half

maximal concentration than histidine. Second, histidine induces higher amplitude of

current (2-4 folds larger) than histamine at same concentration. Another metabolite of

histamine, t-MHA activates homomultimeric β3 receptors but with higher EC50 and lower

amplitude of currents. Histamine retained the ion permeability of homomultimeric β3

receptors. Homomultimeric β3 receptors show the pharmacology like H2 and H3

metabotropic histamine receptors. The histamine antagonists thioperamide and

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158

famotidine inhibit histamine-evoked current. In case of homomultimeric β3 receptors

thioperamide acts as a competitive antagonist and is more active than famotidine. This is

the first report, which shows that homomultimeric β3 receptors behave like histamine-

gated ion channels.

We also investigated if β2 receptors are functionally expressed. Until now, nobody has

reported that β2 subunit of GABAA receptors can form homomultimeric functional

channels even though there are some reports which point out the spontaneous activity

without any ligand activation by homomultimeric β2 receptors. We stated that possible

homomultimeric β2 receptors may act like histamine-gated ion channels rather than

GABA. In this study we have demonstrated the functional expression of homomultimeric

β2 receptors in Xenopus oocytes. GABA as well as histamine and its metabolites activate

homomultimeric β2 receptors. The effect of activation by GABA is nearly 1\8 fold less

than histamine and its metabolites. In addition, we found that the magnitude of

spontaneous opening of homomultimeric β2 receptor channel was predominant and the

spontaneous channel activity accounts for open probability.

The inability of other groups to access the functional expression of homomultimeric

β2 receptors could be due to the different level of spontaneous activity of

homomultimeric β2 receptors in oocytes under different experimental conditions. We

have found that the magnitude of the spontaneous activity appeared to be so predominant

that it nearly overshadowed the magnitude of GABA-activated current in cells expressing

homomultimeric β2 subunits. Overall the magnitude of the PTX-sensitive current was 5-9

folds larger than that of the GABA-activated current. This indicates that the tendency of

homomultimeric β2 subunits to open spontaneously increase with a decrease in the

sensitivity of these receptors to GABA. It is therefore plausible to predict that when

homomultimeric β2 receptors channel open spontaneously at the maximal probability,

these receptors might no longer respond to activation by GABA.

Chapter 9 Discussion

159

The most important finding for the homomultimeric β2 receptors is that they are activated

by histamine and its metabolites. The amplitude of current activated by them is much

higher than the GABA-evoked current. Comparing it with PTX-evoked 'outward' current,

histamine-evoked current showed higher amplitude of the current also. Our different

approach to look for functional expression of homomultimeric β2 receptors lead us to be

able to measure the activity of these homomultimeric receptors. Also pharmacologically,

we were able to block the histamine-evoked current by various antagonists like

thioperamide, famotidine, HTMT and harmane.

We confirmed our finding that β subunits of GABAA receptors can form functional

homomultimeric channels by our electrophysiological, pharmacological studies. We ruled

out the possibility of any artifact or contamination of any other subunit as all 3 different β

subunits exhibited the histamine-evoked current and could be studied by further by using

histamine antagonists.

β2 and β3 are pharmacologically different. We found that β2 is more active than β3

subunit of GABAA receptors w.r.t. histamine. Also we have already shown that though

all these β subunits were activated by histamine they exhibited different pharmacology.

This may have various important aspects. Although there is evidence suggesting that

other type of GABAA receptors subunits apart from β subunits also could be involved in

spontaneous channel activity of the wild type of GABAA receptors, it may be at partly

rely on the presence of distinct β subunits (Sigel et al., 1990; Krishek et al., 1996). There

is evidence showing that spontaneous opening of GABAA receptor channels can be

detected in-vivo in spinal cord neurons (Mathers, 1985) and in pituitary cells (Taleb et al.,

1987; Hamann et al., 1990).

Different actions of histamine on homo- and heteromultimeric GABAA receptors We were also interested to further characterize the homomultimeric β receptors. Why

does histamine activate homomultimeric β receptors directly but does not directly act on

heteromultimeric α1β1 receptors? Here it is only potentiating the GABA response but not

Chapter 9 Discussion

160

opening the channel directly. This can be explained by looking into the structural

arrangement of subunits in heteromultimeric receptor complexes of GABAA receptors.

The minimal structural requirement for GABAA receptors gated by GABA is a

heteropentamer built from two different subunits with one peptide derived from the

α class and the other from the β class of variants (Schofield et al., 1987). Thus it was

expected that both subunit classes contribute to the formation of the binding pocket.

Indeed, a number of amino acids are members of both classes and most of them,

conserved within a subunit class, have been identified as being involved in high-affinity

agonist binding. The first one in this series was recognized by the F to L mutation at

position 64 in the rat α1 subunit in an electrophysiological assay (Sigel et al., 1990), later

confirmed by direct photolabelling of the site with (3H muscimol) (Smith and Olsen,

1994) whereas the homologous residue in the α5 variant has shown to be involved in the

formation of the GABA binding pocket. The equivalent residues in the β2 and γ2 subunits

do not have GABA binding properties (Sigel et al., 1992). The two neighboring amino

acids R66, corresponding to R70 in α5, and S68 (T in α6) in the α variant have been

reported to contribute to the GABA binding domain (Boileau et al., 1999) as well as

R120 in α (Hartvig et al., 2000). We suggest that the in the α1β1 receptor the histamine /

propofol binding domain defined by M286, may not be accessible to the histamine in the

same manner as in homomultimeric β receptors, as in the α1β1 heteropentamer there are

lot of chances for different subunit combinations to occur like 3 α and 2 β, 2 α and 3 β or

4 α and 1 β so it seems that the presence of α subunits in the α1β receptors hinders the

accessibility of histamine to the domain of β responsible for histamine gating in

homomultimeric β1.

The finding that histamine does not activate the α1β1 receptors directly can be also

explained by the earlier studies, which indicate that the pentameric GABAA receptors

undergo dynamic structural arrangement during their assembly in the endoplasmic

reticulum, transportation to the cell membrane and activation in the presence of a ligand.

Addition of the ligand triggers a small rotation of the extracellular domains of the

receptor subunits (Unwin, 1995), which then opens the channel pore formed by the

Chapter 9 Discussion

161

adjoining TM2 regions of the five subunits (predicted from the data obtained with

nicotinic acetylcholine receptors (Unwin, 1993)). Using disulphide bond mapping in

recombinant GABAA α1β1 mutant receptors, (Horenstein et al., 2001) could demonstrate

that the extracellular portion of the TM2–lined pore is more flexible than the intracellular

portion and that these domains of the α1 and β1 subunits may rotate asymmetrically, since

homologous residues (α1T261C and β1T261C form disulphide bonds only when the

receptors are activated by GABA. The resulting covalent modification keeps the channel

open. In the pentameric structural arrangement of the α1β1 receptors there may be the

possibility that presence of histamine doesn't change the alteration in the dynamic of the

receptor as a result the TM2 domain is not exposed to the ligand and hence the receptor

in the presence of histamine is not activated. In the case of homomultimeric β receptors

the pentameric arrangement of the β1 subunits is not hindered and only the TM2 domain

of the β1 subunit forms the pore, so there is more accessibility of this region for the

histamine, which leads to its structural rearrangement and eventually leads to the

activation by the histamine.

Also in the case of direct propofol activation it is reported that the direct activation is

absent in α4β1γ2 receptors (Wafford et al., 1996), and greater in α6β3γ2 than in α1β3γ2

receptors though propofol shows a higher efficiency in the potentiation of the GABA

effect (Krasowski et al., 1997). It is clearly evident that subunit structural arrangement

changes the accessibility of propofol to the GABAA receptor. As histamine and propofol

share the same binding site, the different actions of histamine on homo-and

heteromultimeric receptors can be explained in a parallel manner. Histamine action on further GABAA subunits

Our idea that spontaneous activity can be mediated by other subunits of GABAA

receptors lead us to further check some of the newly identified GABAA receptors

subunits. We extended our search for the histamine-activated homomultimeric channels

for GABAA receptors. The ε subunit indeed was the ideal candidate to go further using

this approach as it is well established that ε subunit causes the spontaneous activity when

Chapter 9 Discussion

162

it is incorporated with different subunits of GABAA receptors to form heteromultimeric

receptors (Davies et al., 1997; Whiting et al., 1997). The ε subunit was also important in

our study as we found that this subunit behaves somewhat differently from other GABAA

receptors subunits. For instance, it is not potentiated by barbiturates and benzodiazepines

and its current-voltage relationship does not show outward rectification at the positive

holding potential that are two important characteristics features of other GABAA subunits

(Davies et al., 1997; Whiting et al., 1997).

There are several reports about the heteromultimeric receptor containing ε subunit but till

now nobody has shown the homomultimeric expression of ε subunit. Here we show also

that ε subunit can form functional homomultimeric ε receptors when injected in oocytes.

These homomultimeric receptors show high spontaneous activity and from our

experience, are difficult to measure as the histamine-evoked current has very small

amplitude and because of high spontaneous activity the histamine-evoked currents are

difficult to measure.

We also report here some of the new combinations active at least in recombinant system

which can also be activated by histamine. α1β1ε containing receptors have become the

ideal choice for the characterization of ε subunit further. It is documented that ε competes

for the γ subunit and there are several reports indicating that βγ can form functional

heteromultimeric channels. Our result also indicate that replacing the γ subunit with ε

facilitate the functional expression of heteromultimeric β1ε receptors. Comparing them

with α1β1ε receptor, we found that they are highly spontaneously active and GABA does

not activate them up to 1 mM concentration. Heteromultimeric α1β1ε receptors have very

high affinity for GABA application of GABA on β1ε receptors blocks the spontaneously

active channels. The homomultimeric ε receptors are highly sensitive for histamine

receptor antagonist as most of them have roughly the IC50 of 3-5 µM excluding

pyrilamine which requires very high concentration (~ 300-500 µM) to block the

histamine-evoked currents. Quite interestingly comparing the pharmacology of ε subunit

with various homomultimeric β subunits receptors show marked differences. Cimetidine

Chapter 9 Discussion

163

was not able to inhibit histamine-evoked current in various homomultimeric β subunits

receptors whereas it is highly active antagonist for homomultimeric ε receptors.

Possible functional consequences of histamine potentiation Various subunit combinations of GABAA receptors exhibit GABA-evoked current

potentiation by histamine. Potentiation by histamine on various subunit combinations

gives indication that histamine potentiation is not restricted to a specific location in brain

area. As both GABAergic and histaminergic systems are widely distributed, so our

current finding indicates the global interaction between histamine and GABA in brain.

This can have several pivotal effects. As synaptic GABAA receptors are also potentiated

with histamine to same extent (for example α1β2γ2 were potentiated by histamine up to 4

folds), it proves that synaptically GABA receptor can be targeted by histamine. This

gives an idea of the importance of the localization of the GABAA receptors in the TM

nucleus of the hypothalamus. There can be several possible sources of the histamine to

the GABAA receptors. As it is documented that one possible source of histamine

targeting GABAA receptors are TM neurons. As some of these neurons are containing

both GABA and histamine, one cannot exclude that both neurotransmitters are even co-

released at the same synapse. In addition, histamine can diffuse out of a histaminergic

synapse by a `spill-out` effect as described for GABA-synapses (Rossi and Hamann,

1998) and thus may act on neighboring synaptic or extra synaptic GABAA receptors.

Further, mast cells in the brain are an additional source for histamine. Mast cells occur in

the CNS of many species and up to 50 % of the brain histamine is attributable to the

presence of these cells. By selectively enhancing the GABAA receptors, histamine should

enhance process in which GABA currents participate, especially those related to

histaminergic neurons in the TM nucleus like sleep and wakefulness (Lin et al., 1986 and

1988; Paramentier et al., 2002).

Modulation of GABAA receptors at synapses can have profound effects. For example in

epilepsy, compounds that increase or reduce GABAergic inhibition are used, respectively

as anti-epileptic or pro-convulsive drugs for many forms of human or experimental

epilepsy (Prince, 1978). Impaired inhibition is thought to be important in temporal lobe

Chapter 9 Discussion

164

epilepsy, the most common feature of epilepsy in human. Enhancement in GABA current

by histamine can possibly be used for epilepsy treatment.

Dependency of potentiation by histamine on various subunits combination may lead to

alteration in the function differently in different location of the brain, which means the

histamine-evoked potentiation should enhance more the brain’s area activity where α1β3

receptors are predominantly expressed. This is also an important aspect as GABAA

receptors exhibit different activity for GABA in the various brain area depending on the

subunit combinations. This is also consistent with other modulators like propofol, which

show at lower concentrations a sedative effect in the frontal cortex while at higher

concentration it causes hypnosis in thalamus and at even more higher concentration leads

to immobility due to the action of propofol on spinal cord (Rudolph and Antkowiak,

2004).

Our current finding indicate a possible resolution for a quite interesting but vague

hypothesis which depicts that hypothalamic networks that are involved in sleep

regulation might have a key role in mediating anesthetic induced hypnosis. Although not

identical, sleep and general anesthesia share common EEG patterns. General anesthesia

produces a variety of useful effects, including sedation (defined as a decrease in activity)

and hypnosis (defined as promoting the onset of sleep). (Nelson et al., 2002) showed that

the same brain region may be important for both sleep and anesthetic. The authors report

that TM nucleus is also a key target for at least two anesthetics that are known to act at

GABAA receptors. During deep anesthesia (so called stage III), EEG recordings resemble

those associated with non-REM sleep, suggesting that they are generated by similar

patterns of cortical and thalamocortical activity. In their study the authors discovered the

anatomical sites responsible for the hypnotic action of propofol and found that it is

located in TM nucleus. Our findings lead into the same approach though we did not

perform experiments in TM nucleus but it is quite evident that physiological localization

of propofol and histamine at least in one brain region show their common mode of

activity.

Chapter 9 Discussion

165

Our results report for the first time that antagonist defined for histamine can have the

possible side effects in the body via interaction with GABAA receptors. So, it should be

taken in care when these antagonists are administrated in the body. Our result may give a

clue about some of the side effects observed during the therapy in which histamine

antagonist are used. It is also of importance that thioperamide acts on propofol and

inhibits propofol-evoked current in the range of histamine IC50. As far as from our

knowledge, this is the first time report for the thioperamide acting as a potent antagonist

for propofol. So anesthesia mediated by propofol can be overcome by using antagonists

on the basis of thioperamide in the future.

Histamine and neurological diseases We assume that the current findings may lead to a new approach about interaction

between GABA and histamine in certain brain disorders like schizophrenia.

Abnormalities in GABA activity in schizophrenia have been consistently shown in the

last ten years. Schizophrenia is associated with both decreased numbers and

abnormalities in the distribution of GABAergic neurons in the cortex, particularly in the

cortical laminae (Kaplen & Saock, 1995). GAD1, the GABA precursor, has also been

shown to decrease activity in schizophrenia. This decreased activity has been found in

nucleus accumbens, putaman, amaygdala and hippocampus. This suggests that loss of

neuroinhibitory control of GABA, in specific regions of the brain, may be responsible for

some of schizophrenia’s symptoms. There is growing evidence to suggest the

involvement of histaminergic pathways in the pathophysiology of schizophrenia.

Overactive histamine activity is thought to contribute significantly to deficit symptoms

such as apathy and social withdrawal associated with the disorder, while the efficacy of

certain atypical antipsychotic may include their action at histamine receptors (Rauscher et

al., 1980; Nakai et al., 1991). Many patients diagnosed as schizophrenic have either a

chronic excess or deficiency of blood histamine and its end product t-MHA (Schwartz et

al., 1971). Nutritional treatment correcting these imbalances has led to great

improvement or recovery for most such patients. One of histamine’s many roles in the

body is to act as on inhibitory neurotransmitter. It is used to promote alpha wave activity

in the brain which enables an individual to handle anxiety and stress easier (McLeod et

Chapter 9 Discussion

166

al., 1998). If the person is deficient in histidine, it leads to a lack of histamine and creates

unbalances in calming alpha-rhythms in the brain allowing the excitatory beta waves

(responsible for the brain activity that leads to anger and tension) to promote (McLeod et

al., 1998). We speculate, during schizophrenia, when the GABA level in brain is altered,

brain tries to compensate the loss of inhibitory neurotransmission. Increase in histamine

level in schizophrenia supports our idea. In schizophrenia, GABAA receptors at the soma

and axon initial segments of pyramidal neurons might be locally unregulated in response

to a reduction in perisomatic inhibitory input from chandlier and wide arbor neurons.

Brain tries to overcome the reduced level of GABA, by promoting higher expression of

GABAA receptors. There should be the enhancement for the affinity of GABA for

GABAA receptors. As our results show that histidine increases affinity of GABAA

receptors for GABA and hence leads to hyperpolarisation in neurons to minimize the

reduction of GABA in brain. Our result shows that even in the presence of low GABA,

histamine potentiates GABAA receptor and may show anxiolytic effects.

Chapter 10 Summary

167

Chapter 10 Summary

In this thesis we demonstrate for the first time that histamine effectively potentiates

GABA-evoked current in heteromultimeric GABAA receptors. Further it directly opens

homomultimeric GABAA receptors that thus function as histamine-gated channels. A

typical 2-4 folds potentiation was found for various combinations of GABAA receptors

expressed in Xenopus oocytes and HEK 293 cells, an effect not restricted to recombinant

receptors but present in native neurons of the olfactory bulb, too. In homomultimeric β

channels, histamine opened the channel with a far better efficacy than GABA. A

pharmacological characterization in combination with the investigation of mutated

β1 subunits (M286W) indicates that histamine acts on the propofol-binding site. We

found that the histamine metabolite histidine had similar potentiating effects. Histamine

and histidine are good candidates for the endogenous ligand acting at the allosteric

modulatory propofol binding site and thus may modulate neuronal events by acting

directly on GABAA receptors in addition to the known histamine action on metabotropic

H1-H3 receptors. We identified further GABAA β subunits as potential components of

mammalian histamine-gated channels, whose existence has been proposed for many

years.

Direct activation by histamine on homomultimeric GABAA receptor could be a missing

link in the various physiological processes like sleep and wake. It is tempting to check the

β subunit expression in the SON nucleus where reports indicate the presence of the

possible ionotropic histamine-gated channel.

The direct activation of histamine on GABAA receptors points the possible answer for

one of the evolutionary questions yet to be answered. Why during the evolution, the

nature just produced metabotropic histamine receptors and not a distinct class of

ionotropic histamine channels in vertebrates? Other neurotransmitters like GABA and

Chapter 10 Summary

168

ACh have both metabotropic as well as ionotropic receptors, then what was the need not

go in the same trend for the nature to produce ionotropic histamine receptors? Maybe our

bioinformatics approaches are still far from identifying the 'real' histamine-gated ion

channel of vertebrates as it might belong to a yet unidentified class of ion channels

distinct from the superfamily of ligand-gated ion channels. Nevertheless, various GABAA

receptors behave like histamine-gated ion channels in recombinant expression systems

and are thus good candidates for being components of the long sought-after histamine

gated-channels in vertebrates.

Chapter 11 References

169

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Appendix

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Appendix I J. Biol. Chem., Vol. 280, Issue 16, 16254-16262, April 22, 2005

A Novel Chloride Channel in Drosophila melanogaster Is Inhibited by Protons*

Katrin Schnizler , Beate Saeger ¶, Carsten Pfeffer , Alexander Gerbaulet , Ulrich Ebbinghaus-Kintscher¶, Christoph Methfessel , Eva-Maria Franken¶, Klaus Raming¶, Christian H. Wetzel||, Arunesh Saras||, Hermann Pusch||, Hanns Hatt||, and Günter Gisselmann||** Summary

A systematic analysis of the Drosophila genome data reveals the existence of pHCl, a

novel member of ligand-gated ion channel subunits. pHCl shows nearly identical

similarity to glutamate-, glycine- and histamine-gated ion channels, does however not

belong to any of these ion channel types. We identified three different sites, where

splicing generates multiple transcripts of the pHCl mRNA. The pHCl is expressed in

Drosophila embryo, larvae, pupae and the adult fly. In embryos, in situ hybridization

detected pHCl in the neural cord and the hindgut. Functional expression of the three

different splice variants of pHCl in oocytes of Xenopus laevis and Sf9 cells induces a

chloride current with a linear current-voltage relationship that is inhibited by extracellular

protons and activated by ivermectin in a pH-dependent manner. Further, currents through

pHCl channels were induced by a raise in temperature. Our data give genetic and

electrophysiological evidence that pHCl is a member of a new branch of ligand-gated ion

channels in invertebrates with however a hitherto unique combination of pharmacological

and biophysical properties.

Appendix

190

Appendix II Identification of a novel branch of invertebrate ionotropic

acetylcholine receptors in C. elegans Günter Gisselmann, Christian H. Wetzel, Michael Kuczkowiak, Arunesh Saras, Hermann Pusch & Hanns Hatt Lehrstuhl für Zellphysiologie, Ruhr-Universität Bochum, Universitätsstraße 150, 44780 Bochum, Germany

Abstract The wealth of the complete C. elegans genome sequence has given us the possibility to

describe all members of the superfamily of ligand gated ion channels present in an

individual species. Homology analysis of candidate genes for ligand gated cation

channels revealed that the previously described gene ACR-22 defines a new subfamily

consisting of 14 putative gene coding for ligand-gated ion channel subunits with weak

sequence homology to nicotinic acetylcholine receptors. By an functional screening in

Xenopus oocytes we found that ACR-22 and R13A5.4, a close homologe to ACR-22,

were able to build functional homomultimeric ion channels activated by acetylcholine

(ACR-22A: EC50 = 15 µM, R13A5.4: EC50 = 95 µM). proteins. Whereas all general

characteristic sequence features of ligand gated ion channels were found in ACR-22 and

R13A5.4 there were some pronounced sequence differences to typical acetylcholine

gated cation channels. Especially the two adjacent cysteins were absent that are present in

nearly all ligand binding acetylcholine receptor α-subunits. Further, the pore forming

M2-region critical for the ion selectivity of the ion channel showed an untypical charge

distribution. Investigation of ACR-22 channels expressed in Xenopus oocytes reveals that

they were permeable for monovalent cations and calcium. At ACR-22, carbachole and

choline were agonists but nicotine failed to activate ACR-22. d-Tubocurarine, levamisole

and strychnine blocked acetycholine induced currents. The electrophysiological

properties and the sequence analysis suggest that ACR-22 and R13A5.4 define a novel

branch of acetylcholine receptors in C. elegans.

Acknowledgments

191

Acknowledgements This work would have not been conceivable without enthusiastic help and support of many people. Prof. Dr. Hans Hatt, Dr. Günter Gisselmann, Dr. Hermann Pusch, and Mary Grace Lucero, participated directly in the experiments. I would like to give my sincere thanks to Prof. Dr. Hanns Hatt for a very kind, motivating and guidance during my whole study, for support, valuable advice and comments. Words are less to express my gratitude to Dr. Gisselmann for helping me in every aspect of my doctoral work. Especially during our project meetings, his criticisms, the appreciation, and the noble approach given by him are highly appreciated. I show my gratitude to Dr. Pusch for teaching me the basic electrophysiology and giving me always friendly and warmth encouragement. Thomas Lichleitner for help and advice on computer programming and for providing software and H. Bartel and W. Garbowski for technical assistance during the experiments. I ought to give my special thanks to the post docs in our lab specially Christian Wetzel for giving me in depth knowledge for patch clamp techniques; Angela for our collaborative work; Ulrike Thomas for an exceptionally kind parental care and always giving me support and help. On this moment my heartiest thanks to my coach Slavko for help, support and a congenial atmosphere. I would like to mention specially the name of Mary Grace Lucero for her dedication and endless patience during the proof reading of my thesis and for making this thesis to finish on time. I show all my respect, gratitude to my parents, brother and sisters for their benediction, inspiration, for letting me to finish my doctoral study. I want to thank my friends namely, Nils, Jennet, Christian, who made my stay in Germany as one of the memorable days of my life.

1

Curriculum Vitae Name Arunesh Saras Title PhD Date of Birth 20.10.1978 Place of Birth New Delhi, India Nationality Indian Education 1998-2000 BSc (H), Zoology, Kirori Mal College, (New Delhi, India)

2000- 2002 MSc (H), Biotechnology Madurai Kamraj University

(Madurai, India)

2002-2003 Junior Research fellow (JRF) All India Institute of Medical Science (AIIMS) (New Delhi, India).

2003 onwards Graduate Student at the International Graduate School of Neuroscience Ruhr University Bochum, (Bochum, Germany). Scholarships

1991 5th standard : Merit Scholarship given by Govt of India 1992-1997 6th-12th standard : Merit Scholarship given by Govt of India 1998-2000 BSc (1st-3rd year) Merit Scholarship given by Govt of India 2001-2002 MSc (1st-2nd year) Merit Scholarship given by Department of

Biotechnology, India.

2002-2003 Council of Scientific & Industrial Reasearch (CSIR), Joint CSIR-UGC Junior Research Fellowship (JRF) , New Delhi, India.

2

2003 onwards International Graduate School of Neuroscience (IGSN) International Graduate School of Neuroscience Scholarship.

Academic Awards

1983 Gold Medal given by the former President of India, Mr. Gyani Zail Singh 1989 Delhi (Zone Level) Science Fair

First Prize for the Model : Different Surfaces have different friction The model was selected for the State Level Science Fair

Model was telecasted on Indian National Television channel, Delhi Doordarshan,

Programme name – New Scientist

1994 Science Open Merit Test

First prize (among particitpating candiates from school), Organised by Delhi State Science Teacher´s Forum, New Delhi, India.

1994 Lion Pratibha ( General Knowledge Contest) Merit Certificate

1995 Lion Pratibha ( General Knowledge Contest) Merit Certificate

1998 India Quiz Competition First Prize in All India Radio (AIR) Quiz Competition, the program was telecasted on international channel of AIR

1999 All India Biomedical Entrace Examination All India Second Rank Conducted by the Delhi University

2000 Summer Training Award The Second the best Summer Trainee from the MKU since 1990 under the supervision of Vice - Chancellor of Delhi Prof. Deepak Pental

Research Training May-June 2000 Transformation and Regenaration in Arabibopsis by Agrobacterium and Cloning work Mentor : Prof. Deepak Pental Vice Chancellor Delhi University, Delhi

3

January-July 2001 Identification of Superior Biotypes of Casuarina equesatiafolia using RAPD analysis Mentor: Prof. A.K. Gutta Center of Plant Molecular Biology, School of Biotechnoloy, MKU July 2001–June 2002 Effect of Prenatal sound over stimulation on synapses of chick brain stem auditory nuclei: A Transmission Electron Microscopic study Mentor: Prof Shashi Wabhwa Neurobiology Lab Department of Anatomy All India Institute of medical Sciences New Delhi, India October 2002 onwards Functional Expresion and Characterization of histamine - gated chloride channels Mentor: Prof. Dr Hans Hatt Ruhr Univertiy Bochum, International Graduate School of Neuroscience,Bochum, Germany Conferences\ Workshops attended 1. National Symposium on Relevance of Plant Biochemistry and Biotechnology –

Modern trends. Paper presentation.

The American College, Madurai, India. March 1 – 3, 2001

2. Summer cource in Bioinformatics Nijemegen, Netherland, August 18 – 29, 2003 3. SFB – symposium Neuro-vision, Ruhr University Bochum, Germany. March 21 – 23, 2004 4. GRK workshop Borkum `Disease of the Brain` Borkum, Germany October 2 – 5, 2004 5. American Society for Cell Biology Meeting, Poster presentation, Washington DC, USA December 4 – 8, 2004

4

6. Berlin Brain Day Berlin, Germany November 18 – 23, 2004 7. 6th NRW Neuroscience Conference

Gottingen, Germany Febuary 17-20, 2005

Extracurricular Activities 1. Hindi Academy – Poem Competition

Consulation Prize for the poem. 1992-93, New Delhi, India

2. Inter – College Health Awareness Competition

Cartoon Making Competition, Ist prize, New Delhi, India. October 2 – 6, 1996

3. Inter – College Health Awareness Competition Extrempore speech Competition, III rd prize, New Delhi, India. October 2 – 6, 1996

4. Perfect Health Mela Special certificate from Ministry of Health and Welfare, Govt. of India and Ministries of Transport, Environment and and Prohibition, Govt. of Delhi October 2 – 6 , 1996 New Delhi, India

5. Lions Club International VI Dr. N.S. Pradhan Memeorial Debate, 3rd Prize in Inter – University Hindi Debate, January 28th , 1998

6. Inter College Chess Competition

Certificate of Merit 3rd Place University of Delhi, 1998-99

5

Publication

1. A Novel Chloride Channel in Drosophila melanogaster Is Inhibited by Protons*

Katrin Schnizler, Beate Saeger, Carsten Pfeffer, Alexander Gerbaulet, Ulrich Ebbinghaus-Kintscher, Christoph Methfessel, Eva-Maria Franken, Klaus Raming,

Christian H. Wetzel, Arunesh Saras, Hermann Pusch, Hanns Hatt, and Günter Gisselmann.

J. Biol. Chem., Vol. 280, Issue 16, 16254-16262, April 22, 2005