Methoxy-6-Methylflavone and Kavain at Recombinant GABAA Receptors

Home , GABA analogue, GABA receptor agonist, Gaboxadol, Honokiol, Ionotropic GABA receptor, Kavain

Pharmacology profile of 2′-methoxy-6-methylflavone

and kavain at recombinant GABAA receptors

Han Chow Chua BPharm (Hons)

A thesis submitted in fulfilment of the requirements for the award of Doctor of Philosophy

Faculty of Pharmacy The University of Sydney 2016

Acknowledgements

Firstly, I would like to express my sincere gratitude to my supervisor Professor Mary

Collins for her continuous support, guidance, patience and motivation throughout my

PhD. Besides my supervisor, I would like to thank Professor Jane Hanrahan, Dr Nathan

Absalom and Dr Petra van Nieuwenhuijzen for their insightful comments and encouragement, which were precious for the writing of this thesis. My sincere appreciation also goes to Associate Professor Philip Ahring for providing me with valuable guidance in molecular biology and electrophysiology; Associate Professor

Thomas Balle for his immense knowledge in molecular modelling; Dr Raja Viswas for the synthesis of various chemicals used in this study.

I would also like to thank my colleagues and lab buddies (in no particular order): Vivian,

Irene, Radhika, Zirong, Taima, Leonny, Jia, Bryan, Ting, Steve, Terry, Tim, Izumi, Ida and Maja for their friendship, support, shared knowledge and experiences over the years, which helped me stay sane through those hair-pulling moments.

Last but not least, my heartfelt gratitude goes to my family, as none of this would have been possible without their support. They have been a constant source of love, concern, support and strength throughout all these years and I deeply appreciate their faith in me.

Abstract

GABAA receptors (GABAARs) are a class of physiologically- and therapeutically- important ligand-gated ion channels. These pentameric receptors occur in the brain with diverse subunit composition, which confers highly complex pharmacology to this receptor class. An impressive range of clinically-used therapeutics are known to bind to distinct sites found on GABAARs to modulate receptor function. Numerous experimental approaches have been used over the years to elucidate the binding sites of these drugs, but unequivocal identification is challenging due to subtype- and ligand-dependent pharmacology. This thesis focuses on two compounds with anxiolytic effects thought to be mediated via GABAARs. However, as the subunit requirement for their actions at

GABAARs is poorly understood, we sought to investigate this using electrophysiological and mutational approaches on various human recombinant GABAAR subtypes expressed in Xenopus oocytes.

2′MeO6MF

2′-Methoxy-6-methylflavone (2′MeO6MF) is an anxiolytic flavonoid which has been shown to display GABAAR β2/3-subunit selectivity, a pharmacological profile similar to that of the general anaesthetic etomidate. Electrophysiological studies suggest that the full agonist action of 2′MeO6MF at α2β3γ2L GABAARs may mediate the flavonoid’s in vivo effects. However, we found variations in the relative efficacy of 2′MeO6MF

(2′MeO6MF-elicited current responses normalised to the maximal GABA response) at

α2β3γ2L GABAARs due to mixed receptor populations. To identify which receptor(s) underlie the variations observed, we conducted a systematic investigation of 2′MeO6MF

activity at all receptor combinations that could theoretically form (α2, β3, γ2L, α2β3,

α2γ2L, β3γ2L and α2β3γ2L) in Xenopus oocytes using the two-electrode voltage clamp technique. We found that 2′MeO6MF activated non-α-containing β3γ2L receptors. In an attempt to establish the optimal conditions to express a uniform population of these receptors, we found that varying the relative amounts of β3:γ2L subunit mRNAs resulted in differences in the level of constitutive activity, the GABA concentration-response relationships, and the relative efficacy of 2′MeO6MF activation. Like 2′MeO6MF, general anaesthetics such as etomidate and propofol also showed distinct levels of relative efficacy across different injection ratios. Based on these results, we infer that β3γ2L receptors may form with different subunit stoichiometries, resulting in the complex pharmacology observed across different injection ratios. Moreover, the discovery that

GABA and etomidate have direct actions at the α-lacking β3γ2L receptors raises questions about the structural requirements for their respective binding sites at

GABAARs.

Kavain

Extracts from the pepper plant kava (Piper methysticum) are effective in alleviating anxiety in clinical trials. However, the molecular target(s) of the pharmacologically active constituents, kavalactones have not been established. GABAARs are assumed as the in vivo molecular target of kavalactones, but evidence to date is equivocal. In this study, we sought to determine the activity of kavain, the major anxiolytic kavalactone at human recombinant α1β2, β2γ2L, αxβ2γ2L (x = 1, 2, 3 and 5), α1βxγ2L (x = 1, 2 and 3) and

α4β2δ GABAARs expressed in Xenopus oocytes using the two-electrode voltage clamp technique. We found that kavain positively modulated all receptors regardless of the subunit composition, but the enhancement was greater at α4β2δ GABAARs. The

benzodiazepine site antagonist flumazenil failed to block the action of kavain at α1β2γ2L receptors, supporting previous findings that kavalactones do not modulate GABAARs via the benzodiazepine binding site. As part of our efforts to investigate the interaction of kavain with other ligands on a receptor level, we found that kavain modestly inhibited the activity of general anaesthetics. Mutational studies showed that while α1M236W,

β2M286W and β3N265M point mutations impaired general anaesthetics’ sensitivity, kavain’s modulation was only reduced by β3N265M. This finding suggests that kavain and general anaesthetics may share a common mechanism, but differ in their molecular determinants. As transgenic mice bearing the β3N265M point mutation are resistant to anaesthetic effects, we infer that the significant attenuation of kavain modulation with the introduction of this point mutation in vitro supports a direct kavain-GABAAR interaction, in contrast to the non-specific interaction previously suggested.

Table of Contents

Chapter 1 Introduction

1.1 Ion channels and neuronal signalling…………………………………....…..… 1 1.2 Fast synaptic transmission is mediated by neurotransmitter-gated ion channels………………………………………………………………………..… 2 1.3 GABAergic inhibition is essential for neuronal function …………………... 3 1.4 GABA synthesis, storage, release, reuptake and degradation…………...... 3 1.5 GABA and its receptors: GABAA, GABAB and GABAC……………………... 6 1.6 GABAARs: Physiological and clinical significance…………………………... 10 1.7 Architecture of GABAARs and other pentameric LGICs…………………… 11 1.8 GABAAR assembly: selective subunit oligomerisation………………………. 15 1.9 Multiple subtypes, locations and actions of GABAARs…...…………………. 17 1.10 Subunit stoichiometry and arrangement of GABAARs…………………….... 20 1.11 Pharmacology of GABAARs………………………………………...………… 23 1.11.1. GABA binding sites: extracellular β+α‒ interfaces……………………. 24 1.11.2. Benzodiazepine binding sites…………………………………………... 27 1.11.2.1 High-affinity site: extracellular α+γ‒ interface……….……... 27 1.11.2.2 Low-affinity benzodiazepine binding site(s)……...... … 29 1.11.3 Anaesthetic binding sites in the TMD…………………………………. 30 1.11.3.1 Transmembrane β+α‒ interfaces………………………..… 31 1.11.3.2 Transmembrane α+β‒ and γ+β‒ interfaces…………………. 31 1.11.3.3 Transmembrane β+β‒ interface……………………….…… 32 1.11.3.4 The promiscuity of anaesthetics binding……………...…..… 32 1.11.3.5 β2/3N265: critical sensitivity determinant, elusive role in binding……………………………………………………… 33 1.11.4 Neurosteroid binding sites in the TMD………………………….……... 34 1.11.5 δ-selective ligands…………………………….……………………….. 35 1.12 Natural products of plant origin and GABAARs……………….………… 36 1.12.1 Flavonoids…………………………………………..…………………. 36 1.12.2 Valerenic acid………………………………………………………..… 39 1.12.3 Honokiol and magnolol………...……………………………………… 40 1.13. Thesis rationale………………………………………………………………… 41 1.13.1 2′MeO6MF……………………………………………………………. 41 1.13.2 Kavalactones…………………………………………….…………..… 41 References…………………………………………………………………………….. 43

Chapter 2 Materials and Methods

2.1 Introduction………………………...………………………………………… 56 2.2. Safety, regulations and ethics……………………………………………….…. 57 2.3 Materials…………………………………………………………...…………… 58 2.3.1 Chemicals and reagents…………………………….………………..… 58 2.3.2 Bacterial growth media………………………………………………... 59 2.3.3 Buffers…………………………………………………………………. 59 2.3.4. Enzymes, competent cells and kits…………………………………….. 60 2.3.5 General apparatus and equipment……………………………………… 61 2.3.6 GABAAR subunits…………………………………………………...… 62 2.3.7 Oligonucleotide primers……………………………………………….. 62 2.4 Methods………………………………………………………………………… 63 2.4.1 Molecular biology……………………………………………………... 63 2.4.1.1 Transformation of plasmid DNA into E. coli…….…………. 63 2.4.1.2 Growth of E. coli cultures……………………………..…….. 63 2.4.1.3 Bacterial glycerol stocks……………………………………. 64 2.4.1.4 Purification, quality assessment and quantification of plasmid DNA………………………………………..……………….. 64 2.4.1.5 Verification of purified plasmid DNA identities……………. 66 2.4.1.6 Linearisation of plasmid DNA……………………………… 67 2.4.1.7 Synthesis, quality assessment and quantification of mRNA... 67 2.4.1.8 Site-directed mutagenesis…………………………………... 69 2.4.2. Xenopus oocytes: harvesting, defolliculation and microinjection……… 71 2.4.3 Two-electrode voltage-clamp technique……………………………….. 72 2.4.4 Data analysis…………………………………………………………. 74 References…………………………………………………………………………….. 75

Chapter 3

The direct actions of GABA, 2′-methoxy-6-methylflavone and general anaesthetics at β3γ2L GABAA receptors: Evidence for receptors with different subunit stoichiometries

3.1 Introduction…………………………………………………………………. 76 3.2 Results………………………………………………………………………..…. 79 3.2.1 Variations in the relative efficacy of 2′MeO6MF activation at α2β3γ2L GABAARs………………………………….………………………...... 79 3.2.2 2′MeO6MF does not activate binary α2β3 or homomeric α2, β3 and γ2L GABAARs……………………………………………………………... 81

3.2.3 2′MeO6MF activates β3γ2L GABAARs with variable efficacy……..… 83 3.2.4 β3γ2L GABAARs of different subunit stoichiometries vary in pharmacological properties……………………………………………. 87 3.2.5 Differential activation of β3γ2L receptors with different subunit stoichiometries by general anaesthetics……………………………….. 91 3.3 Discussion……………………………………………………………………..... 94 References…………………………………………………………………………….. 99

Chapter 4

The modulation of recombinant GABAARs by kavain, the major constituent of the anxiolytic kava extract is attenuated by the β3N265M point mutation

4.1 Introduction……………………………………………………………...….. 102 4.2 Materials and Methods…………………………………………………...… 105 4.2.1 Primer sequences used in site-directed mutagenesis………………….. 105 4.2.2 Electrophysiological studies………………………………………….. 106 4.3 Results……………………………………………………………………..… 107 4.3.1 Effect of GABAAR subunit composition on kavain activity………….. 107 4.3.2 Kavain’s differential effects at α1β2γ2L and α4β2δ GABAARs…...… 110 4.3.3 Effect of membrane potential on kavain modulation…………………. 112 4.3.4 Interaction of kavain with various allosteric binding sites……………. 113 4.3.4.1 Benzodiazepines…………………………………………... 113 4.3.4.2 Allopregnanolone…………………………………………. 114 4.3.4.3 DS2……………………………………………………...… 115 4.3.4.4 General Anaesthetics……………………………………… 117 4.3.5 Kavain activity at receptors containing α1M236W, β2M286W and β3N265M mutations…………………………………………………. 120 4.3.5.1 α1M236W…………………………………………………. 121 4.3.5.2 β2M286W…………………………………………………. 121 4.3.5.3 β3N265M…………………………………………………. 122 4.4 Discussion…………………………………………………………………… 126 References………………………………………………………………………...…. 131

Chapter 5

General discussion: Implications from the functional expression of βγ GABAARs

5.1 Flexibility in the subunit requirement of ligand binding sites………...….. 134 5.1.1 GABA binding sites at the β+γ‒ interfaces?...... 135 5.1.2 High-affinity benzodiazepine site at the β+γ‒ interface?...... 136 5.1.3 Other allosteric binding sites…………………………………………. 139 5.2 Possibility of mixed βγ and αβγ GABAARs…………...……………………. 140 5.3 Perspective……………………….………………………………………….. 143 References…………………………………………………………………………… 145

Chapter 6

GABAARs as the molecular target for kavalactones: Future directions

6.1 Effect of enantiomerically pure kavain…………………………………….. 147 6.2 Potential for synergistic modulation of kavalactones………….………..… 148 6.3 Effect of kavalactones in transgenic mice………………………………….. 148 6.4 Concluding remarks………………………………………………………… 149 References…………………………………………………………………………… 150

List of figures

Figure Title Page 1.1 GABA synthesis, storage, release, reuptake and degradation. 4 1.2 GABA structure and conformations. 7 1.3 GABAA, GABAB and GABACRs and their selective ligands. 8 1.4 Conserved overall architecture of pLGICs. 12 1.5 The crystal structure of a human β3 homomeric GABAAR (PDB: 4COF). 14 Model structures of the ternary αβγ and binary αβ GABAARs 1.6 22 demonstrating their respective subunit stoichiometry and arrangement. Schematic illustration of an αβγ GABAAR highlighting the orthosteric 1.7 24 and allosteric binding sites. 1.8 Homology model of the GABA binding site at α1β2γ2 receptors. 25 Chemical structures of various classes of ligands that bind to the 1.9 27 orthosteric and allosteric binding sites at GABAARs. 1.10 Natural products of plant origin that modulate GABAARs. 37 The quality assessment and concentration measurement of purified 2.1 66 plasmid DNA. The quality assessment and concentration measurement of synthesised 2.2 69 mRNA. 3.1 2′MeO6MF directly activates native γ2-containing GABAARs. 77 Characterisation of α2β3γ2L GABAARs expressed at a 3:1:3 injection 3.2 80 ratio. Subunit combinations with detectable function but were not activated by 3.3 82 2′MeO6MF. 3.4 2′MeO6MF activity at β3γ2L GABAARs. 84 GABA concentration-response curves of β3γ2L GABAARs expressed at 3.5 86 1:20 ratio. Characterisation of β3γ2L GABAARs expressed at 1:15, 1:50 and 1:100 3.6 88 ratios. 3.7 Etomidate activates both β3γ2L and β3 GABAARs 92 Propofol activates β3γ2L (1:15) and (1:100) receptors with different 3.8 94 relative efficacies. 4.1 Chemical structures of the six major kavalactones found in kava. 104 4.2 Kavain potentiates GABAARs with no apparent subtype selectivity. 108 GABA concentration-response curves for all receptors subtypes expressed 4.3 109 in this study Kavain produced greater enhancement of GABA-elicited currents at 4.4 111 α4β2δ than α1β2γ2L GABAARs. Membrane potential has negligible effect on the degree of potentiation of 4.5 112 kavain Flumazenil-insensitive kavain potentiation has less-than-additive effect on 4.6 114 diazepam action

Figure Title Page The lack of effect of kavain on allopregnanolone potentiation at α1β2γ2L 4.7 115 GABAARs.

4.8 The lack of effect of kavain on DS2 potentiation at α4β2δ GABAARs. 116 Kavain modestly reduced etomidate potentiation, but did not affect the 4.9 118 direct activation caused by etomidate at α1β2γ2L GABAARs. Kavain did not affect propofol potentiation, but modestly reduced 4.10 119 propofol activation at α1β2γ2L GABAARs. The putative β+α‒ anaesthetic binding sites in a model structure of the 4.11 120 α1β2γ2 GABAAR. Effect of α1M236W and β2M286W point mutations on etomidate and 4.12 123 propofol sensitivity. The pronounced effect of β3N265M point mutation on etomidate and 4.13 124 propofol sensitivity. Effect of α1M236W, β2M286W and β3N265M point mutations on kavain 4.14 125 activity. Aligned sequence of the α1 subunit that forms the complementary face of 5.1 the GABA binding site, and the corresponding region in the γ2 subunit 135 (which forms the complementary face of the benzodiazepine site). 5.2 Diazepam potentiation at β2γ2 GABAARs. 138 5.3 Comparable actions of various allosteric ligands at αβγ and βγ GABAARs. 139 GABA concentration-response curves of αβ, αβγ, and βγ GABAARs 5.4 expressed in Xenopus oocytes using different injection ratios from 142 pharmacological studies. 5.5 Zn2+ sensitivity at α1β2, β2γ2L and α1β2γ2L receptors. 143

List of Tables

Table Title Page

The working list of native GABAAR subtypes categorised based on the 1.1 18 strength of evidence proposed by Olsen and Sieghart (2008). 2.1 General chemicals and reagents. 58 2.2 Chemicals used in electrophysiological studies. 58 2.3 Media used for bacterial growth. 59 2.4 Buffers used for molecular biology purposes. 59 2.5 Buffers used in the handling of Xenopus oocytes. 60 2.6 Enzymes, competent cells and kits. 60 2.7 General apparatus and equipment. 61 2.8 A summary of all GABAAR subunits used in this study. 62 2.9 mRNA transcription setup. 68 2.10 Site-directed mutagenesis reaction setup. 70 2.11 Cycling parameters for the site-directed mutagenesis reactions. 71 Characteristics of receptors expressed in various combinations of α2, β3 3.1 81 and γ2L GABAAR subunits. GABA, 2′MeO6MF, etomidate and propofol concentration-response 3.2 curve parameters at β3γ2L (1:15), (1:50) and (1:100) GABAARs derived 90 from curve-fitting procedures. Primers used to introduce the α1M236W, β2M286W, and β3N265M point 4.1 105 mutations. 4.2 Subunit mRNA ratios used to express different GABAAR subtypes. 106 GABA concentration-response curve parameters at various receptor 4.3 110 subtypes derived from curve-fitting procedures.

Publication Chua HC, Absalom NL, Hanrahan JR, Viswas R, & Chebib M (2015). The direct actions of GABA, 2'-methoxy-6-methylflavone and general anaesthetics at β3γ2L GABAA receptors: Evidence for receptors with different subunit stoichiometries. PLoS ONE. DOI: 10.1371/journal.pone.0141359.

Conferences

2015 ‘Pharmacological characterisation of kavain at GABAARs’ (poster), Society for Neuroscience 45th annual meeting (#477.25/A101), Chicago, US.

‘Pharmacological characterisation of kavain at GABAARs’ (oral), Inter- University Neuroscience and Mental Health Conference 2015, Brain Science UNSW, Sydney, Australia.

‘Evidence of stoichiometric forms and non-canonical binding sites for β3γ2L GABAARs’ (oral), The University of Sydney Pharmaceutical Student Symposium 2015, American Association of Pharmaceutical Scientists (AAPS) The University of Sydney Student Chapter, Sydney, Australia.

‘2′MeO6MF activity reveals multiple possible stoichiometric forms of β3γ2L GABAARs’ (oral), Gage Ion Channels and Transporters conference, Canberra, Australia.

2014 ‘2′MeO6MF activity reveals multiple possible stoichiometric forms of β3γ2L GABAARs expressed in Xenopus oocytes’ (poster), Society for Neuroscience 44th annual meeting (#124.10/B25), Washington D.C., US.

‘2′MeO6MF activity reveals multiple possible stoichiometric forms of β3γ2L th GABAARs expressed in Xenopus oocytes’ (poster), 24 Neuropharmacology Conference, Washington D.C., US.

Chapter 1

Introduction

______

1.1. Ion channels and neuronal signalling

The movement of ions into and out of neurons is a fundamental event that underlies neuronal signalling. One of the major pathways for ion transport is through ion channels, membrane-spanning proteins which form hydrophilic pores that selectively allow distinct ions to diffuse rapidly down the electrochemical gradient. These channels are not continuously open, but are activated or gated by specific stimuli such as voltage changes

(voltage-gated ion channels), mechanical distortion of membrane (stretch-gated ion channels) and the binding of chemical agonists (ligand-gated ion channels; LGICs). Ion channels fluctuate between open and close states, with different transition states such as inactivation and desensitisation in between that help in the regulation of current flow across the membrane.

In an unstimulated neuron, there is a difference in the ion concentrations across the cell membrane such that the electrical potential in the cytoplasm is more negative than the extracellular environment (resting membrane potential is typically around -60 mV).

When an ion channel opens in response to a stimulus, the subsequent current flow alters the resting membrane potential, either by making it less negative (depolarisation) or more negative (hyperpolarisation). If a depolarising stimulus is sufficiently strong, different types of ion channels are sequentially activated, resulting in a self-propagating electrical

2 signal called an action potential which is used to communicate information from the peripheral nervous system (PNS) to the central nervous system (CNS), and vice versa.

1.2. Fast synaptic transmission is mediated by

neurotransmitter-gated ion channels

Majority of neurons communicate via the release of neurotransmitters which bind and activate specific neurotransmitter-gated ion channels found on the postsynaptic neuron, resulting in fast synaptic conductances (< 10 ms). Depending on the type of receptors and the ionic conditions encountered, neurotransmitters can produce excitatory or inhibitory responses. Generally, synaptic excitation is mediated by the transmitter glutamate, whereas inhibition relies on γ-aminobutyric acid (GABA) and glycine.

Glutamate activates cation-permeable NMDA- (N-methyl-D-aspartate), AMPA- (α- amino-3-hydroxyl-5-methyl-4-isoxazole-propionate) and kainate-gated ion channels, resulting in Na+, K+ or Ca2+ ion influx into the postsynaptic neuron. The membrane depolarises as a result, and the probability of action potential generation increases. In the brain, GABA-mediated fast synaptic inhibition is achieved via the activation of anion-

- permeable GABA type A receptors (GABAARs). The subsequent flow of Cl ions into the postsynaptic neuron leads to membrane hyperpolarisation and hinders action potential firing. Like GABAARs, glycine receptors are anion-permeable channels, and are found in great abundance in the spinal cord and brain stem.

1.3. GABAergic inhibition is essential for neuronal function

Information relay from one neuron to the next is dependent on the use of glutamate to propagate excitatory signals. GABA-mediated (GABAergic) inhibition can be perceived simply as the force that counters excitation, but in reality, it participates in the coordination of neuronal activity in a highly complex manner. Specifically, GABAergic inhibition controls the rate of neuronal firing in response to increasing excitatory input

(i.e., gain control) [1, 2], progressively recruits firing neurons to match the intensity of stimulus across a large dynamic range [3, 4], governs the stimulus selectivity of individual neurons (i.e., tuning) [5-8], generates and synchronises rhythmic changes in membrane potential (i.e., oscillations) which enable neurons to cooperate to convey signals to common downstream targets [9-11]. Given its fundamental regulatory roles, it is not surprising that GABAergic inhibition is implicated in various physiological processes such as neuronal development [12], learning [13-15], memory [16], circadian rhythm [17-

19], sleep [20, 21] and pain [22, 23], despite the fact that GABAergic neurons only make up approximately 30 % of all neurons [24].

1.4. GABA synthesis, storage, release, reuptake and

degradation

In the GABAergic neurons, GABA is produced from the removal of a carboxyl group from glutamate, a process catalysed by the rate-limiting enzyme glutamic acid decarboxylase (GAD), which exists in two major isoforms, GAD65 and GAD67 (Figure

1.1). GAD67 is constitutively active, and is responsible for the synthesis of a basal pool of GABA (> 90 % of GABA in the CNS), whereas GAD65 is activated when there is an increased demand during rapid inhibitory transmission. Disrupted GAD activity has been

4 shown to lead to detrimental consequences, with GAD67 knockout mice suffering from neonatal fatality [25, 26], and GAD65 knockout mice prone to epileptic seizures and anxiety despite appearing normal at birth [27, 28].

Figure 1.1. GABA synthesis, storage, release, reuptake and degradation. (1) In the presynaptic neuron, GABA is synthesised from glutamate by glutamic acid decarboxylase (GAD). (2) GABA is packaged into vesicles by the vesicular GABA transporters (VGAT). (3) GABA release into the synaptic cleft involves complex intracellular trafficking machineries which dock and fuse GABA-loaded vesicles with the presynaptic neuronal membrane. (4) GABA transporters (GATs) mediate the reuptake of GABA into the presynaptic neuron and surrounding glial cells to terminate GABAergic transmission. GABA recycled into the presynaptic neuron is mainly repackaged into synaptic vesicles for subsequent use. (5) In glial cells, GABA undergoes enzymatic degradation in the mitochondria.

Once synthesised, GABA is loaded into synaptic vesicles by the vesicular GABA transporters (VGATs), a process which relies on the electrochemical gradient of protons across the vesicular membrane created by the H+-ATPase. The transport of GABA via

VGATs into vesicles is a prerequisite for GABA release into the synapses. Consistent with this critical role, when the VGAT gene was knocked out in mice, it was found that the postsynaptic GABA-mediated inhibitory currents were drastically reduced, and

VGAT-deficient mice were found to die perinatally [29, 30].

GABA-loaded vesicles are accumulated in the presynaptic neuron terminals. The release of GABA into the synaptic cleft is stimulated by the arrival of an action potential at the presynaptic terminal, which leads to the opening of voltage-gated calcium channels, allowing the influx of Ca2+ ions. The elevated Ca2+ concentration then triggers a cascade of complex intracellular trafficking machineries which dock and fuse GABA-loaded vesicles with the presynaptic nerve terminal membrane [31, 32]. Following synaptic vesicle fusion, the contents of the vesicles are discharged into the synaptic cleft where

GABA rapidly diffuses across to bind to a variety of GABA-gated receptors found on the postsynaptic neuron. The process of docking and fusing vesicles with the presynaptic nerve terminal membrane, and the subsequent release of GABA is referred to as exocytosis, which happens in an ultrafast manner (< 1 ms). Multiple receptors expressed presynaptically such as metabotropic GABA type B (GABAB) [33] and µ-opioid [34] receptors, ionotropic kainate [35] and nicotinic acetylcholine receptors [36, 37] are known to modulate the probability of GABA release.

Termination of GABA-mediated inhibition is achieved primarily through the reuptake of

GABA via GABA transporters (GATs) into the presynaptic neuron and neighbouring glial cells. GATs work as symporters to co-transport GABA with Na+ and Cl- ions, in other words, both GABA and the ions are transported in the same direction. The removal of GABA from the synaptic cleft serves to (1) govern its concentration and time spent in the synaptic cleft, thus preventing the excessive activation of GABA receptors, and (2) to

6 prevent GABA spillover to other synapses nearby, allowing individual synapses to work independently, thus ensuring synaptic specificity/fidelity. Four pharmacologically- distinct isoforms of GATs have been identified thus far (GAT-1, GAT-2, GAT-3 and

BGT-1). These transporters are distributed differentially, for instance, GAT-1 and GAT-

3 are predominantly expressed in neurons and glial cells respectively [38].

Most GABA molecules taken up into the presynaptic neuron are repackaged into synaptic vesicles for subsequent use, but some undergo enzymatic degradation. Unlike neurons, glial cells are not involved in synaptic transmission, and enzymatic degradation is the only fate for transported GABA in this cell type. In the mitochondria, GABA transaminase (GABA-T) is responsible for the conversion of GABA to succinate semialdehyde (SSA), which is then oxidised to succinic acid by SSA dehydrogenase.

Succinic acid enters the Krebs cycle and gets converted into α-ketoglutarate, which is used in the synthesis of glutamate, the precursor of GABA.

1.5. GABA and its receptors: GABAA, GABAB and GABAC

GABA is a four-carbon, non-protein amino acid which exists predominantly in the zwitterionic form at physiological pH (Figure 1.2A). The backbone structure of GABA is made up of single carbon-carbon bonds with low energy barriers for rotations, allowing

GABA to adopt diverse conformations ranging from fully extended, partially folded to fully folded (Figure 1.2B). The inherent flexibility of the GABA molecule is believed to be important for the differential actions of GABA across the large number of GABA- recognising macromolecules such as enzymes, transporters as well as ionotropic and metabotropic receptors. Over the last four decades, the use of conformationally-restricted

GABA analogues has helped in the delineation of the structural requirements of the

GABA binding sites on different proteins.

Figure 1.2. GABA structure and conformations. (A) The chemical structure of GABA (unionised; top), and its physiological zwitterionic form (below). (B) The fully-extended (top), partially-folded (middle) and fully-folded (bottom) conformations of GABA.

When GABA is released into the synaptic cleft, it binds and activates three distinct types of receptors known as GABAA, GABAB and GABAC receptors (GABAA/B/CRs).

Structurally, GABAA and GABACRs are five-subunit protein complexes with a central channel pore selectively permeable to Cl- ions (Figure 1.3). However, the subunit composition of these receptors are very different. GABAARs are heteromeric (i.e., made up of more than one type of subunit), with 16 different subunit genes (α1–6, β1–3, γ1–3,

δ, ε, θ and π) known to date. In contrast, GABACRs are homomeric receptors made up of one out of the three ρ (ρ1–3) subunits. However, there is evidence to suggest that these receptors can be heteromeric consisting of ρ1 and ρ2 [39] or ρ1 in combination with

GABAA α1 and γ2 subunits [40].

Unlike the ionotropic GABAA/CRs, GABABRs belong to the family of G protein-coupled receptors (GPCRs). GABABRs function as a heterodimer with a GABA-binding B1

8 subunit and a surface-expression mandatory B2 subunit. Each subunit has an extracellular domain, a seven-helix transmembrane domain and an intracellular domain which contains a coiled-coil helix that takes part in the dimerisation of GABABRs (Figure 1.3). The intracellular loops of the B2 subunit are coupled to guanidine-binding proteins (G- proteins), and upon activation, trigger different effector mechanisms that result in the inhibition of presynaptic neurotransmitter release or membrane hyperpolarisation.

Figure 1.3. GABAA, GABAB and GABACRs and their selective ligands. (Top) The physiological actions of GABA are mediated by GABAA, GABAB and GABACRs which are structurally, pharmacologically, and functionally different. GABAA and GABACRs - are pentameric Cl ion channels, whereas GABABRs are dimeric metabotropic receptors coupled to G-proteins. Following GABA binding, GABAA and GABACRs are activated and the subsequent Cl- influx causes membrane hyperpolarisation and reduced neuronal excitability. The inhibitory actions of GABABRs are dependent on the G-proteins downstream effector mechanisms which lead to the inhibition of neurotransmitter release or the gating of ion channels that result in membrane hyperpolarisation. (Bottom) The chemical structures of selective ligands for different types of GABA receptors.

Pharmacologically, GABAARs are conventionally defined by their selective inhibition by the competitive antagonist bicuculline and modulation by benzodiazepines and barbiturates. GABABRs are activated by the GABA analogues (R)-baclofen and are competitively antagonised by (R)-phaclofen and (R)-saclofen. GABACRs are insensitive to bicuculline and (R)-baclofen, are selectively activated by the conformationally- restricted GABA analogue cis-aminocrotonic acid (CACA), and are inhibited by the competitive inhibitor (1,2,5,6-tetrahydropyridin-4-yl)methylphosphinic acid (TPMPA).

The unique pharmacological profile of GABACRs has prompted its separation from

GABAA and GABABRs. Notably, an International Union of Pharmacology (IUPHAR) committee has classified GABACRs as a ‘specialised set’ of GABAARs and recommended against the use of the term ‘GABAC’ [41]. However, for the purpose of this thesis, ‘GABAC’ is used to describe the specific category of bicuculline- and (R)- baclofen-insensitive, ρ-containing GABA receptors.

GABAARs are the most widespread GABA receptors in the CNS, facilitating fast inhibitory responses (< 10 ms) in the regulation of neuronal excitability. These receptors are highly concentrated on postsynaptic neuronal membrane, but they are also expressed extrasynaptically and presynaptically (Figure 1.1). The differential subcellular localisation of GABAARs involves specific scaffolding proteins such as gephyrin [42, 43] and collybistin [44] for synaptic and radixin [45] for extrasynaptic receptor anchoring.

Like GABAARs, GABACRs are also involved in fast inhibitory transmission, but these receptors are more restricted in their topographical distribution, being highly enriched in brain areas related to the visual pathways such as the retina, thalamus and superior colliculus. The subcellular localisation mechanisms of GABACRs also appears to be different from GABAAR, with microtubule-associated protein 1B (MAP-1B) shown to specifically localise GABACRs at inhibitory synapses [46]. In contrast to GABAA/CRs,

GABABRs are involved in slow, prolonged inhibitory transmission that happens over the timescale of sub-seconds to minutes. GABABRs are widely distributed in the CNS, and are found pre-, post- and extrasynaptically (Figure 1.1). Presynaptic GABABRs can act as feedback autoreceptors to inhibit the release of GABA, or as heteroreceptors to inhibit glutamate release in nearby excitatory synapses. These effects are achieved either through

Gαi/o inhibiting adenylyl cyclase which interferes with vesicle fusion and release, or Gβγ inhibiting voltage-gated calcium channels, preventing Ca2+-dependent neurotransmitter release [47]. Postsynaptic GABABRs can be found at synaptic and extrasynaptic sites.

These receptors contribute to membrane hyperpolarisation via the activation of inwardly rectifying K+ channels which allows K+ ions to flow out of the cell.

1.6. GABAARs: Physiological and clinical significance

As the main mediator of GABAergic inhibition in the CNS, GABAARs represent a class of physiologically- and clinically-important receptors. Several lines of evidence support this notion. The mutation or deletion of various genes encoding for GABAAR subunits in mice is highly disruptive to the normal phenotype, causing developmental defects, sensorimotor dysfunction, hypersensitive behaviour, anxiety, epilepsy and/or reduced lifespan [48-51]. In humans, aberrant GABAAR trafficking, expression and/or gating effects have been implicated in autism [52], schizophrenia [53] and a range of idiopathic epileptic syndromes [54]. Genetic association studies have also linked GABAAR subunit genes with alcohol dependence [55], eating disorder outcomes [56], autism [57, 58] and bipolar disorders [59, 60]. Further evidence comes from the successful clinical utilisation of GABAAR modulators in the treatment of CNS-related disorders such as insomnia, anxiety and epilepsy, as well as in the induction of anaesthesia in surgical patients. While clinically useful, the misuse of these drugs poses risks of dependence, addiction, abuse

11 and life-threatening conditions associated with overdose or withdrawal. Understanding the functions of GABAARs and the underlying mechanisms of the clinical and undesirable effects of GABAAR-targeting drugs to improve the selectivity and safety of these therapeutics are topics of intensive research. The properties of GABAARs are also the focus of this thesis.

1.7. Architecture of GABAARs and other pentameric LGICs

The neurotransmitter-gated ion channels can be divided into three main structural classes: the trimeric P2X receptor family, the tetrameric glutamate receptor family and the pentameric LGIC (pLGIC) superfamily. GABAARs are a member of the pLGIC superfamily which includes the nicotinic acetylcholine receptors (nAChRs), 5- hydroxytryptamine type 3 receptors (5-HT3Rs) and glycine receptors. The term ‘Cys-loop receptors’ has also been used to describe pLGICs due to a highly conserved 13-amino- acid loop found in the extracellular domain which is connected by a disulphide bridge between two cysteine (Cys) residues. This characteristic Cys-loop is only found in eukaryotic pLGICs but not in their prokaryotic counterparts.

Due to the inherent technical challenges associated with the overexpression, solubilisation, purification and crystallisation of mammalian pLGICs, the initial understanding of their three-dimensional structures was inferred from other homologues including the acetylcholine binding protein (AChBP) from the snail Lymnaea stagnalis

[61], the heteromeric nAChR from the Torpedo marmorata electric organ [62], the bacterial Erwinia chrysanthemi LGIC (ELIC) [63] and Gloeobacter violaceus LGIC

(GLIC) [64], and the Caenorhabditis elegans glutamate-gated chloride channel α

12 homopentamer (GluClα) [65]. More recently, X-ray crystal structures of a mouse 5-

HT3AR [66] and a human GABAAR [67] have been reported. Collectively, these studies reveal a similar overall architecture among these receptors. Each receptor subunit has an extracellular domain (ECD), a transmembrane domain (TMD) and an intracellular domain (ICD) which is absent in ELIC, GLIC and GluClα (Figure 1.4). The ECD is mostly made up of β sheets and contains agonist binding site, the TMD consists of the pore-forming α-helices and the structurally variable ICD is involved in receptor assembly, trafficking and clustering.

Figure 1.4. Conserved overall architecture of pLGICs. Published structures of the soluble AChBP from the snail Lymnaea stagnalis (PDB: 1I9B) which is homologous to the ECD of nAChRs, the heteromeric nAChR from the Torpedo marmorata electric organ (PDB: 2BG9), the bacterial homologues ELIC (PDB: 2VL0) and GLIC (PDB: 3EAM), mouse 5-HT3AR (PDB: 4PIR) and human GABAAR (PDB: 4COF) show three distinct domains: an extracellular domain (ECD; red), a transmembrane domain (TMD; blue) and an intracellular domain (ICD; green) which is absent in ELIC, GLIC and GluCl. The resolutions (in Å) are indicated above individual structures.

Besides providing structural details, these published receptor structures are also useful models to study various ligand binding sites, the different transition states underlying channel gating, and the mechanism of ion permeation through the channel pore [68]. For instance, molecular dynamics simulation using the closed-pore ELIC and the open-pore

GLIC and GluClα crystal structures has allowed for the modelling of structural events

13 which unfold at distinct receptor states [69, 70]. However, this information has limited application due to the low sequence identities of these structures.

For the interest of this thesis, the crystal structure of the β3 homomeric GABAAR obtained at 3 Å resolution will be described in more detail [67]. This receptor subtype is unlikely to be physiologically relevant, but its functional expression in heterologous systems is well known, and has been used as a model to study heteromeric GABAARs [71-77]. The receptor stands approximately 110 Å in height when viewed parallel to the membrane

(Figure 1.5A). The five subunits assemble in a doughnut-like shape with a diameter around 80 Å, when viewed from the extracellular space, down the channel pore (Figure

1.5B). The large ECD (~65 Å) of each subunit is made up of an N-terminal α-helix followed by a β-sandwich core with ten anti-parallel β-sheets (Figure 1.5C). The TMD consists of four membrane-spanning α-helices (M1–M4), with the M2 helices of all five subunits arranging themselves to form a tapered ion-conducting pore (Figure 1.5 B and

C). The outermost M4 helix harbours the C-terminus on the extracellular end. The ICD contains a small M1–M2 loop and a much larger M3–M4 loop (G333–N446; residue numbering follows sequence of P28472 in UniProt) which was replaced with a 7-amino- acid linker for the crystallisation of this structure (Figure 1.5C).

Figure 1.5. The crystal structure of a human β3 homomeric GABAAR (PDB: 4COF). (A) Cartoon representation of GABAAR viewed parallel to the membrane, coloured according to secondary structures (α-helices are in red except for the pore-forming helices (orange), β-sheets are in blue). (B) Left, GABAAR viewed from the extracellular space, with the five subunits labelled 1–5. Right, Transmembrane region of GABAAR, with the ECDs simplified as blue ovals for clarity. The arrangement of the four TMDs (M1–M4), the C-terminus (purple circles), and the M2–M3 loop (green) are illustrated. (C) Topology of a single subunit of GABAAR, rainbow coloured from the N-terminus (red) to the C- terminus (purple). The β-sheets of the ECD, the α-helices of the TMDs, the characteristic Cys-loop and other relevant loops are indicated. Note: the intracellular M3-M4 loop (G333–N446) was replaced with a 7-amino-acid linker for crystallisation. Figures were prepared using Maestro, v. 9.5.014, Schrӧdinger, LLC.

1.8. GABAAR assembly: selective subunit oligomerisation

Human genome sequencing has identified at least 16 GABAAR subunit genes (α1–6, β1–

3, γ1–3, δ, ε, θ and π), excluding the ρ subunits (ρ1–3). Given the heteromeric nature of

GABAARs in vivo, this long list of subunits, together with the splice variants of some of the subunits (e.g., alternate splicing of precursor GABAA γ2 mRNA results in two splice variants, a short (γ2S) and a long (γ2L) variant) allow for an enormous range of theoretically possible subunit combinations. However, experimental evidence suggests that only a few dozen combinations exist in vivo (see section 1.9), indicating that

GABAAR assembly is a selective, and not a random process. A hierarchical assembly mechanism has been proposed, in which certain subunits are preferred over others to form dimeric intermediates that ultimately assemble into pentameric complexes [78].

Different methods have been used to define the rules underlying receptor assembly. In concert, data obtained using functional, immunoimaging and sucrose gradient centrifugation techniques suggest that both α and β subunits are obligatory for the surface expression of fully functional pentameric receptors in heterologous cell systems [79-82].

The additional third γ subunit has been shown to enhance the efficiency of receptor assembly [81]. In contrast, the recombinant expression of individual α, β and γ subunits, and the α+γ and β+γ combinations mainly yielded di-, tri- and tetrameric oligomers which were retained in the endoplasmic reticulum [80, 83]. There are a few notable exceptions, however, with the homomeric β1 and β3 and the heteromeric β3γ2 receptors expressing readily in heterologous systems [72-74, 76, 84].

Amino acid residues important for assembly have been identified in several studies using the chimeric receptor and site-directed mutagenesis approaches (reviewed in [78]). These residues are found mainly in the ECD, and to a lesser extent in the intracellular M3-M4 loop. In accordance with these data, the β3 GABAAR crystal structure revealed extensive energetically favourable interactions such as hydrogen bonds, salt bridges and van der

Waals forces along the interfaces between subunit ECDs [67]. Disruption to these interactions could affect receptor assembly, and may be the reason for impaired GABAAR surface expression with epilepsy-associated mutations found in the N-terminal regions such as β3G32R and γ2R43Q [85-87].

While considerable insights have been provided by these studies, the molecular determinants could not be firmly established for several reasons. Firstly, these determinants differ depending on the partner subunits. For example, four residues in the

ECD of the β3 subunit (G171, K173, E179 and R180) have been identified to be critical for the assembly of β3 and β3γ2 receptors [73]. However, they are not compulsory for the assembly of αβ receptors. In another study that investigated the role of the N-terminal regions in the expression of α1β2γ2 GABAARs, subunit-specific contributions to receptor assembly were found [88]. When deleted, the N-terminus of the α1 subunit had the most prominent effect on the expression of α1β2γ2 receptors, whereas deletion in similar regions of the β2 and γ2 subunits had minimal effect on surface expression. Second, multiple residues may be involved in oligomerisation with the same neighbouring subunit

[89]. As such, when a putative binding residue is mutated and has no effect on receptor expression, it does not necessarily indicate no participation in intersubunit linking.

Conversely, an expression-impairing mutation does not validate its significance in subunit oligomerisation, as the residue may indirectly contribute to this process (e.g., by stabilising interacting regions). All in all, GABAAR assembly is a highly complex, multi-

17 step process which involves subunit-specific determinants that govern the subunit composition of GABAARs found natively.

1.9. Multiple subtypes, locations and actions of GABAARs

In recent years, it has become evident that the multiplicity in GABAAR subunit composition (or subtypes) is one of the main reasons for the heterogeneity observed in their cellular and subcellular distributions, biophysical characteristics, pharmacological properties, in addition to physiological functions [90-92]. Furthermore, the subunit composition of GABAARs is plastic. Alterations in brain GABAAR subtypes have been reported under various developmental and pathological conditions [93-95]. As such, answering the question “which GABAAR subtypes actually occur in vivo?” is essential to understanding the diverse roles played by GABAergic inhibition.

Currently, existing experimental techniques are unable to unequivocally identify

GABAAR subunit composition in neurons. To help determine the likelihood of a receptor subtype being expressed physiologically, the IUPHAR committee has introduced five classification criteria [41]. Briefly, (1) the expression and co-assembly of the subunit proteins of a receptor candidate must be demonstrated in heterologous systems, along with (2) the characterisation of its biophysical and pharmacological properties. In the brain, (3) the subunit mRNAs or proteins should co-localise on the cellular and subcellular levels, and (4) the co-precipitation of these subunits would provide stronger evidence. More importantly, (5) a receptor candidate has to show unique functional characteristics that match those of the recombinant receptor subtype, and disruption to

18 the expression of this receptor subtype in genetically engineered mice should lead to altered receptor responses and behavioural profile.

A working list of native GABAAR subtypes was proposed following stringent consideration of the criteria discussed (Table 1.1). These receptors are categorised as

“identified”, “existence with high probability” or “tentative”. Evidence from immunochemical, pharmacological and genetic studies collectively indicate that the majority of GABAARs are composed of α, β and γ2 subunits in the CNS. The α1β2γ2 subtype is the most prevalent isoform (approximately 50–60 % of all GABAARs), and are expressed in almost all brain regions. The γ2 subunit also co-assembles with other α and

β variants in the brain, but these receptors are found in considerably less abundance and are restricted in their regional (e.g., α5-containing receptors are expressed in high density in hippocampus) and cellular (e.g., the α2β3γ2 and α3β3γ2 subtypes are highly enriched in hippocampal pyramidal neurons and cholinergic neurons of the basal forebrain, respectively) distributions.

Table 1.1. The working list of native GABAAR subtypes categorised based on the strength of evidence proposed by Olsen and Sieghart (2008) [41].

Identified High probability Tentative α1β2γ2 α1β3γ2 αβγ1 Synaptic α2βγ2 α1βδ αβγ3 α3βγ2 α5β3γ2 αβθ α4βγ2 αβ1γ/αβ1δ αβε α5βγ2 αβ αβπ α6βγ2 α1α6βγ/α1α6βδ αxαyβγ2a Extrasynaptic α4β2/3δ α6β2/3δ a αxαy represents two different α subunit isoforms (α1–6).

In addition to the region- and cell-specific expression patterns, these receptors also show distinct subcellular localisation. The α1βγ2, α2βγ2 and α3βγ2 isoforms are typically found postsynaptically, whereas the α4βγ2, α5βγ2 and α6βγ2 isoforms are found extrasynaptically (Figure 1.1). In contrast, the δ subunit which preferentially assembles with the α6 and β2/3 subunits in the cerebellar granule cells [96], and with the α4 and

β2/3 subunits in the forebrain [97], are exclusively expressed at extrasynaptic sites. Due to their proximity to the site of vesicle release, synaptic GABAARs are exposed to saturating GABA concentrations (~1–3 mM) that diffuse away from the synaptic cleft rapidly (clearance time < 1 ms) [98, 99]. This transient activation of synaptic GABAARs is responsible for phasic inhibition, a form of fast, point-to-point synaptic communication used to control the flow of specific signals. Extrasynaptic GABAARs, on the other hand, are randomly activated by low levels of ambient GABA (low micromolar range) which has escaped from the synaptic cleft on the same or nearby neurons. These receptors are persistently activated, thus provide a basal inhibitory conductance, termed tonic inhibition, which is believed to be important in the regulation of the overall excitability in the brain [91]. Synaptic and extrasynaptic GABAARs are equipped with distinct properties that facilitate their unique roles in their local environments. For example, δ subunit-containing receptors display characteristics such as higher GABA affinity, minimal desensitisation and longer channel open time, which are ideal for continual activation with sustained exposure to low levels of GABA [100]. Synaptic αβγ isoforms have lower GABA sensitivity, and rapid activation and desensitisation kinetics, making them well suited to mediate responses of the phasic vesicular release of GABA.

Contrary to the belief that binary αβ combination has no physiological relevance, there is ample evidence suggesting that these receptors are highly probable to be found in vivo.

The defining characteristics of these receptors such as low conductance states and high

20 sensitivity to Zn2+ inhibition have been detected in single-channel recordings conducted in rat neurons [101-103]. Besides that, receptors consisting only of α and β subunits have also been immunoprecipitated from brain membranes derived from normal or mutant rodents [48, 104-106]. The exact α and β isoforms that make up these receptors remains unclear, but they are thought to constitute a small portion of GABAARs, likely to be expressed at extrasynaptic sites in the brain [103]. To add to the diversity, GABAARs comprised of different types of α subunits (αxαyβγ2) have also been detected in the CNS

[41], with high prevalence suggested in the spinal cord [107]. Other combinations that incorporate the minor subunits (γ1, γ3, θ, ε or π) are likely to occur in vivo, but more evidence is required to confirm their presence. It is expected that the list of native

GABAAR subtypes will expand as the identification techniques continue to improve over time.

1.10. Subunit stoichiometry and arrangement of GABAARs

Theoretically, a pentameric GABAAR composed of three types of subunits (e.g., αβγ) will have 6 configurations with different number of subunits (1α:1β:3γ, 2α:1β:2γ, 3α:1β:1γ,

1α:2β:2γ, 1α:3β:1γ and 2α:2β:1γ). A greater number of unique configurations arises when the spatial orientation of these subunits is taken into consideration. For instance, a receptor with a 2α:2β:1γ subunit stoichiometry can have up to six different arrangements of γ-α-α-β-β, γ-α-β-α-β, γ-α-β-β-α, γ-β-β-α-α, γ-β-α-β-α and γ-β-α-α-β in a counter- clockwise direction around the pore when viewed from the extracellular side. These configurations vary in the number and type of subunit interfaces, and as almost all known ligand binding sites are located interfacially, establishing the physiologically relevant configuration(s) is a prerequisite to understanding the mechanisms of receptor activation and modulation.

Numerous approaches including mutagenesis [108, 109], subunit counting in combination with immunoprecipitation [81] and fluorescence energy transfer microscopy

[110] have been used to determine the stoichiometry of GABAARs. There are limitations associated with each technique, but in combination, these studies support a 2α:2β:1γ stoichiometry for αβγ receptors (Figure 1.6). The subunit arrangement of this receptor subtype has also been elucidated with the help of different concatenated constructs, i.e.,

GABAAR subunit genes fused in tandem with a linker of optimised length joining the C- terminus of a preceding subunit to the N-terminus of a following subunit. These subunits are expressed as a single polypeptide with predetermined orientation, thus constraining the stoichiometry and arrangement of receptors expressed. The combination of various di- and trimeric concatemers suggests that only the γ-β-α-β-α arrangement (Figure 1.6; read counter-clockwise when viewed from the extracellular space) yields receptors with similar pharmacological and kinetic properties established in receptors assembled from free subunits [111-113]. However, the ambiguity remained whether these constructs were sufficient to force the receptors to assemble in the expected arrangement [114]. To address this issue, the pentameric γ-β-α-β-α concatemer was generated, and functional characterisation of this construct supports the findings from other studies [115]. It is worth noting that most of the studies mentioned have been conducted on the most prevalent subtype α1β2γ2, and it is presumed that all αβγ2 subtypes share the same subunit stoichiometry and arrangement.

Figure 1.6. Model structures of the ternary αβγ and binary αβ GABAARs demonstrating their respective subunit stoichiometry and arrangement. (A) The most prevalent native GABAAR subtype αβγ has a stoichiometry of 2α:2β:1γ, arranged γ-β-α-β-α counter-clockwise around the pore when viewed from the extracellular side. (B) The binary αβ receptors have been shown to assemble in two different stoichiometries of 2α:3β or 3α:2β, with a β-β-α-β-α or α-β-α-β-α arrangement respectively.

In the case of the binary αβ receptors, concatemers with stoichiometry of either 2α:3β or

3α:2β (Figure 1.6), but never both in the same study, form functional receptors with wild type-like properties [111, 113, 116]. It remains unclear whether one stoichiometry predominantly forms or both stoichiometries co-exist when these subunits are allowed to assemble freely. For δ-containing receptors, the accurate determination of the exact number and arrangement of the subunits is hindered by the promiscuous assembly properties of the δ subunit. Several studies suggest αβδ receptors share similar stoichiometry and arrangement as αβγ receptors, with the δ subunit replacing the γ subunit

(2α:2β:1δ; δ-β-α-β-α) [117, 118]. However, 1α:1β:3δ, 1α:2β:2δ, 1α:3β:1δ and 2α:1β:2δ stoichiometries have also been detected with the subunit-counting and concatemer strategies [119-121]. To make things more complicated, the δ subunit is flexible in its positioning in the pentameric complex, producing receptors with distinct pharmacological properties [119, 121]. Similar promiscuity has also been observed with the ε subunit, which co-assembles with α and β subunits to form the αβε subtype with potential stoichiometries of 2α:2β:1ε and 2α:1β:2ε or with α, β and γ subunits to form the

23 quaternary αβγε subtype with multiple arrangements [122, 123]. Less is known about the stoichiometry and arrangement of receptors containing the θ and π subunits.

1.11. Pharmacology of GABAARs

GABAARs have been described as the most complicated member of the superfamily of

LGICs for the plethora of receptor subtypes and the rich pharmacology. The orthosteric

GABA binding sites aside, it has been estimated that there are more than 10 potential ligand binding sites which are distributed at various locations throughout the receptor

(Figure 1.7). A range of ligands such as benzodiazepines, barbiturates, anaesthetics, neurosteroids and convulsants can bind to distinct allosteric (non-orthosteric) sites to exert their physiological or clinical effects by influencing GABAAR function, either by causing or stabilising conformational changes in the receptor. Some of these sites are found at specific receptor subtypes, which define the unique pharmacological signatures of these receptors. The properties of the GABA binding sites and several well-studied allosteric binding sites are discussed below.

Figure 1.7. Schematic illustration of an αβγ GABAAR highlighting the orthosteric and allosteric binding sites. For the purpose of discussion, the approximate binding site locations are indicated by the prototype ligands. The principal (+) and complementary (‒ ) faces of each subunit are also labelled.

1.11.1. GABA binding sites: extracellular β+α‒ interfaces

A common feature of Cys-loop receptors is that the neurotransmitter binding pocket is located at the extracellular interface between two adjacent subunits. Each subunit contributes three (in some cases four) non-contiguous regions that form the binding site.

By convention, the principal (+) face of the neurotransmitter binding site is made up of three loops (named loop A, B and C) and the complementary (‒) face has three β-strands and/or one loop (loop D, E, F and/or G; Figure 1.8A) [124]. For GABAARs, the β and α subunits bear the principal and complementary components of the GABA binding site respectively. In the most prevalent αβγ subtypes (2α:2β:1γ; γ-β-α-β-α), there are two

GABA binding sites per receptor at the β+α‒ interfaces (Figure 1.7). Channel opening occurs with GABA occupying just one site, but the probability increases greatly when both sites are engaged [125-127]. These identical sites contribute asymmetrically to

25 receptor activation, a phenomenon which has been tentatively correlated with the binding sites assuming different conformations depending on their flanking subunits (one site sits between γ and β, the other sits between α and γ; Figure 1.7) [127]. The existence of other

GABA binding interfaces consisting of the promiscuous δ subunit has also been proposed, but the molecular details of these sites are poorly understood [128].

Figure 1.8. Homology model of the GABA binding site at α1β2γ2 receptors. (A) The GABA binding site is located at the β+α‒ extracellular interface. The principal (loop A, B and C) and the complementary (loop D, E and F) components which form the binding pocket are highlighted in different colours. (B) GABA (green; ionised form) binding mode predicted by induced fit docking. Residues on both the β and α subunits which are implicated in GABA binding are indicated. Figures were taken from Bergmann et al. (2013) [129].

Mutational, radioligand binding and photoaffinity labelling studies have identified residues with aromatic, hydroxylated and charged side chains on both the α and β subunits that are critical determinants in GABA sensitivity [130, 131]. It is currently inconclusive how these residues are located spatially to interact with GABA, but homology modelling has shed some light on the GABA binding mode. In 2013, Bergmann et al. created a model of the α1β2γ2 GABAAR based on the C. elegans GluClα and ELIC crystal structures [129]. When docked into the orthosteric binding site, GABA forms (1) salt bridges with α1Arg66 and β2Glu155, (2) hydrogen bonds with α1Thr129 and β2Thr202,

26 and (3) cation-π interaction with β2Tyr205 (Figure 1.8B). These predictions are in good agreement with experimental evidence, but crystal structures of GABA-bound GABAARs will be necessary for validation.

Besides GABA, other structurally similar chemicals can access the orthosteric binding sites to elicit distinct functional responses. Some prototypical ligands include agonists muscimol and isoguvacine, partial agonists THIP/gaboxadol and P4S, and competitive antagonists bicuculline (Figure 1.3) and gabazine (Figure 1.9). The classification of ligands as agonists or partial agonists is not absolute, and is relative to the function of

GABA, which is dependent on receptor subtype. For instance, GABA is known to act as a partial agonist at αβδ receptors, exhibiting higher affinity and lower intrinsic efficacy, unlike its full agonist activity at αβγ receptors [132, 133]. Such functional difference has resulted in the agonist muscimol and partial agonist THIP (at αβγ receptors) to exhibit superagonism (higher maximal efficacy than GABA) at αβδ receptors [134, 135].

Figure 1.9. Chemical structures of various classes of ligands that bind to the orthosteric and allosteric binding sites at GABAARs.

1.11.2. Benzodiazepine binding sites

1.11.2.1. High-affinity site: extracellular α+γ‒ interface

Homologous to the β+α‒ GABA binding sites, a structurally-related pocket exists at the extracellular α+γ‒ interface (Figure 1.7). This allosteric site is commonly referred to as the benzodiazepine binding site, named after the prototype drugs. Benzodiazepines do not directly activate GABAARs, but they are able to modulate receptor function.

Benzodiazepine site agonists (e.g., diazepam; Figure 1.9) potentiate GABAARs to produce clinically useful effects such as sedation (for insomnia), anxiolysis (for anxiety), anticonvulsant (for epilepsy), myorelaxant (for muscle spasms) and amnesia (for invasive

28 medical procedures). Conversely, benzodiazepine site inverse agonists (e.g., DMCM) inhibit GABAARs to exert anxiogenic, convulsant and memory-enhancing effects.

Benzodiazepine site competitive antagonists (e.g., flumazenil) block the actions of other benzodiazepines, and can be used clinically as antidote for benzodiazepine toxicity.

Ligands that are structurally unrelated to the benzodiazepines can also access the high- affinity benzodiazepine site (e.g., zolpidem) to elicit similar responses.

Extensive effort has gone into the delineation of benzodiazepine pharmacology using the representative ligand diazepam. It is now clear that diazepam, at low concentrations, indiscriminately modulates α1βγ2, α2βγ2, α3βγ2 and α5βγ2 GABAARs and is insensitive at α4βγ2 and α6βγ2 GABAARs. A conserved histidine residue found in the α1, α2, α3 and

α5 subunits confers high-affinity diazepam sensitivity, whereas the homologous arginine in the α4 and α6 subunits renders these receptors diazepam insensitive. The histidine-to- arginine mutation has been shown to selectively abolish diazepam activity at

α1H101Rβ2γ2, α2H101Rβ2γ2, α3H126Rβ2γ2 and α5H105Rβ2γ2 recombinant receptors, and the reverse mutation produced diazepam-responsive α4R99Hβ2γ2 and

α6R100Hβ2γ2 receptors [136-138]. Genetic studies have exploited this functional switch to engineer point-mutated mice bearing diazepam-insensitive α1, α2, α3 and/or α5 subunits. Behavioural analysis of single point-mutated (only one type of diazepam- insensitive α subunit) and triple point-mutated (only one type of diazepam-sensitive α subunit) mice in response to diazepam has helped define the functional role(s) of individual α subunits [90, 107, 139]. Taken together, these studies strongly suggest that sedation is primarily mediated by α1βγ2, anxiolysis by α2βγ2, myorelaxant by both α2- and α3βγ2, and amnesia by α5βγ2 GABAARs. In light of these findings, as well as the fact that the α1-preferring zolpidem displays mainly sedative hypnotic effects, there is a

29 surge of interest in the development of subtype-selective benzodiazepine site agonists with improved therapeutic profiles [140]. In particular, α2-selective modulators with anxioselectivity devoid of sedating property are highly sought after [141].

It is also worth mentioning that while the α4- and α6-containing receptors lack high sensitivity to diazepam, it does not mean that the high-affinity benzodiazepine site is absent altogether. Indeed, members of the same ligand class such as bretazenil and Ro

15-4513 are able to bind to both diazepam-sensitive and insensitive GABAARs with high affinities [142]. Furthermore, the molecular determinants of these benzodiazepines are different from those of diazepam, as their functions are not diminished when the conserved histidine is mutated to arginine [138].

1.11.2.2. Low-affinity benzodiazepine binding site(s)

Many benzodiazepines also display additional low-affinity components at GABAARs.

The binding site(s) responsible for these actions are poorly characterised, but evidence thus far suggests that the receptor responses mediated by these site(s) do not require the

γ subunit, and may vary depending on the ligand, the concentration tested, and the type of subunits present [143]. The classical benzodiazepine diazepam, for instance, modulates

α1β2γ2 GABAARs via two distinct mechanisms manifested as a biphasic potentiation with nanomolar and micromolar potencies [144]. The nanomolar component is mediated by the high-affinity benzodiazepine site, and can be selectively blocked by the antagonist flumazenil. The micromolar component, however, is flumazenil insensitive, and has been shown to be affected by mutations in the TMD where anaesthetics usually bind (section

1.11.3). The physiological function of the low-affinity component is unclear, but has been

30 postulated to underlie diazepam’s anaesthetic property. Like diazepam, CGS 9895

(Figure 1.9) has high- and low-affinity components at α1β3γ2 GABAARs [145]. At nanomolar concentrations, CGS 9895 acts as a neutralising modulator to block diazepam action. At micromolar concentrations, CGS 9896 potentiates GABAARs, an effect selectively mediated by a cluster of residues found on the α and β subunits (α1V211,

α1S204, and β3Q64). Homology modelling has located this low-affinity benzodiazepine site on the extracellular α+β‒ interfaces, homologous to the classical benzodiazepine site

(Figure 1.7). The therapeutical potential for subtype-selective ligands targeting this site is an area of interest [146-148].

1.11.3. Anaesthetic binding sites in the TMD

The transmembrane region of GABAARs harbours many solvent-accessible pockets which are the site of action for a range of structurally diverse chemicals including volatile anaesthetics (e.g., enflurane, isoflurane), general anaesthetics (e.g., etomidate, propofol), anticonvulsants (e.g., barbiturates, loreclezole), and sedative hypnotics (e.g., methaqualone) (Figure 1.9). These chemicals exhibit multiphasic activity at GABAARs, enhancing GABA-elicited current at clinical concentrations, directly activate receptors at high micromolar concentrations, and in some cases, act as open-channel blockers at even higher concentrations [149-151]. These binding sites are traditionally described in the context of their prototypical ligands. However, as more experimental evidence has become available to indicate that these sites are not unique to the prototype drugs, it is more appropriate to address them according to their locations.

1.11.3.1. Transmembrane β+α‒ interfaces

Two key approaches have hitherto contributed to the elucidation of the first class of anaesthetic binding sites, commonly referred to as the etomidate binding sites. First, using photoreactive analogues of etomidate ([3H]azietomidate and [3H]TDBzl-etomidate) which covalently label binding residues upon UV irradiation, photoaffinity labelling studies have jointly identified the βM286 (located in the M3 domain) and αM236 (located in the M1 domain) residues [152, 153]. Etomidate is able to inhibit the photolabelling of these residues in a concentration-dependent manner, further supporting etomidate’s interaction with both residues. Second, conventional and cysteine-substitution mutagenesis studies have also demonstrated that these methionine residues are important determinants of etomidate binding and function [154-156]. With the help of homology models based on various crystal structures of related receptors, βM286 and αM236 are predicted to be located at the β+α‒ interfaces in the TMD, just below the orthosteric binding sites (Figure 1.7). Other residues in the vicinity implicated in etomidate binding include βF289 (M3), βV290 (M3), αL232 (M1), αT237 (M1) and αI239 (M1) [153, 156].

1.11.3.2. Transmembrane α+β‒ and γ+β‒ interfaces

Another class of anaesthetic binding sites has recently been identified in α1β3γ2

3 GABAARs using R-[ H]mTFD-MPAB, a photoreactive barbiturate analogue [157]. These binding pockets are located at the α+β‒ and γ+β‒ interfaces, in homologous positions to the etomidate binding sites (Figure 1.7). Binding residues labelled by R-[3H]mTFD-

MPAB include αA291 (M3; equivalent to βM286), αY294 (M3) and γS301 (M3; equivalent to βM286) found on the principal faces of the respective subunits, whereas the

βM227 (M1; equivalent to αL232) is found on the complementary face.

1.11.3.3. Transmembrane β+β‒ interface

Studies conducted in α1β3 GABAARs have also demonstrated that the photoreactive etomidate analogues and R-[3H]mTFD-MPAB which preferentially label β+ and β‒ residues respectively, also contact residues found on the opposite faces [153, 158].

[3H]Azietomidate and [3H]TDBzl-etomidate have been shown to photolabel βM227 on the complementary side, whereas R-[3H]mTFD-MPAB also photolabelled βM286 and

βF289 located on the principal face. These findings suggest that a homologous cavity at the β+β‒ interface, which is absent in the αβγ subtypes is also accessible to these anaesthetics.

1.11.3.4. The promiscuity of anaesthetics binding

While [3H]azietomidate and [3H]TDBzl-etomidate are highly selective for the β+α‒ interfaces, and R-[3H]mTFD-MPAB prefers the α+/γ+β‒ interfaces, the different classes of binding pockets are not exclusive to etomidate or barbiturates. Structurally dissimilar anaesthetics are able to compete with these photoreactive analogues to prevent residue photolabelling [157, 159]. Evidence from mutagenesis studies are in concordance with these findings. The substitution of methionine by tryptophan at residue 286 of the β subunit (βM286W) attenuates etomidate, propofol, volatile anaesthetics and methaqualone activity at GABAARs [160-162]. Moreover, the substituted cysteine accessibility method (SCAM) has also revealed that propofol protects the modification of

α1M236C and β2M286C mutants by sulfhydryl-reactive reagent, indicating that propofol also contacts these residues [163, 164].

Thus, it appears that a structurally diverse set of chemicals including, but most likely not limited to the ligands investigated to date are able to access different number and classes of interfacial anaesthetic binding sites to exert their clinical effects. This heterogeneity is best exemplified by the convulsant S-mTFD-MPAB which selectively binds to the γ+β‒ interface to inhibit GABAARs [165], and the general anaesthetic propofol that binds non- selectively to the β+α‒, α+β‒,γ+β‒ and β+β‒ interfaces to potentiate GABAARs [157,

158]. More complexity is likely as other binding sites in the α+γ‒ transmembrane region, intrasubunit helical bundles and intracellular loop have also been suggested [143].

However, due to limitations associated with available experimental techniques, and the lack of crystal structures of anaesthetic-bound GABAARs, their existence remains to be confirmed.

1.11.3.5 β2/3N265: critical sensitivity determinant, elusive role in binding

The modulatory and agonistic actions of etomidate are more efficacious at β2/3- than β1- containing GABAARs [149]. This functional difference is determined by a homologous residue at position 265 (M2 domain) of the β subunits (β1: serine; β2/3: asparagine).

Mutational studies have demonstrated that etomidate sensitivity at β1-containing

GABAARs is enhanced when the point mutation β1S265N is introduced, whereas the reverse mutation β2/3N265S negatively affects etomidate sensitivity [166]. This functional switch is shared with other β2/3-selective ligands such as loreclezole [167] and methaqualone [162]. When the βN265 residue is replaced with a methionine, the point mutation βN265M not only completely eliminates etomidate sensitivity, but also dramatically diminishes activity of propofol and enflurane [160, 168, 169]. Transgenic mice bearing the β2N265S or β3N265M point mutations are also resistant to distinct pharmacological actions of etomidate, propofol and pentobarbital [170-172].

Homology models built based on the recent β3 homomeric GABAAR as well as other related LGIC crystal structures have revealed that the N265 residue is located in the proximity of the β+α‒ etomidate/propofol binding sites [157, 158, 164]. However, none of the photoreactive anaesthetic analogues mentioned above has labelled this residue.

Furthermore, the chemical modification of βN265C mutant by sulfhydryl-reactive probes was not protected by etomidate and propofol in SCAM studies, which argues against the direct interaction of anaesthetics with the residue [163, 164]. The ability of a single point mutation to have such drastic effects on different anaesthetics has led to the speculation that a common anaesthetic transduction pathway is impaired. Nevertheless, until more conclusive evidence becomes available, whether β2/3N265 mediates the binding and/or gating of anaesthetics remains tentative.

1.11.4. Neurosteroid binding sites in the TMD

Endogenous steroids such as allopregnanolone (Figure 1.9) and their synthetic analogues have neuroactive effects such as anxiolysis, analgesia, sedation, anticonvulsant and anaesthesia mediated via GABAARs [173]. These neurosteroids potently enhance

GABAAR function at low nanomolar concentrations, and activate GABAARs at submicromolar-to-micromolar concentrations. The mechanism(s) of action of neurosteroids are different from those of benzodiazepines and anaesthetics as demonstrated in mutational [160], photoaffinity labelling [174], and transgenic mice behavioural studies [171, 175]. Through the construction of chimera receptors followed by site-directed mutagenesis, an electrophysiological study has shown that αQ241 (M1),

αN307 (M4) and αN410 (M4) residues are responsible for the potentiation by neurosteroids, whereas the agonistic action of neurosteroids is mediated by αT236 (M1) and βY284 (M3) residues [176]. Homology modelling located the modulatory site at the

M1-M4 interface within the α subunit, and the activation site occurs at the β+α‒ interfaces, between the M3 and M1 domains of the β and α subunits respectively (Figure

1.7). These residues are conserved in all α and β subunits, which may explain the lack of subtype selectivity in neurosteroid activity [177].

Naturally-occurring neurosteroids that block GABAARs (e.g., pregnenolone sulphate) also exist, but their physiological actions are not well defined. Evidence from binding and mutational studies suggest that inhibitory neurosteroids do not share the same sites with neurosteroids that augment GABAAR function. A residue found in the M2 domain of the

α1 subunit (V256) which is located further down the channel pore has been implicated, but this putative inhibitory neurosteroid binding cavity is controversial and awaits further clarification [178]. All in all, neurosteroids appear to interact in a complex manner with at least three distinct transmembrane sites to potentiate, activate and inhibit GABAARs.

1.11.5. δ-selective ligands

The δ subunit is exclusively found in extrasynaptic GABAARs with pronounced physiological significance and therapeutic relevance. However, the understanding of the properties of these receptors is hindered by their promiscuous assembly properties and also the scarcity of δ-selective pharmacological tools. The hypnotic drug THIP/gaboxadol directly activates αβδ with higher potency and efficacy (relative to GABA) than αβγ

GABAARs [132]. However, THIP does not discriminate between αβ and αβδ receptors expressed in vitro [134], which may reflect the difficulty in achieving subunit selectivity by targeting the structurally conserved orthosteric binding sites. Unlike THIP, 4-chloro-

N-(2-thiophen-2-ylimidazo[1,2-a]pyridin-3-yl)benzamide (DS2; Figure 1.9)

36 predominantly acts as an efficacious positive modulator at α4/6βδ, has limited efficacy at

αβγ and is inactive at αβ GABAARs [179]. Consistent with this δ-selective profile, the enhancement of tonic currents by DS2 in neurons is absent in δ-knockout mice [179, 180].

The location of the DS2 binding site(s) is not known, but the orthosteric and allosteric sites that etomidate, classical benzodiazepines, allopregnanolone and pentobarbital bind to have been excluded by binding and mutational studies [179]. Besides DS2, ketamine,

AA29504 and JM-11-43-A also have δ-preferring activity, but there is limited evidence to confirm their selectivity [181-183].

1.12. Natural products of plant origin and GABAARs

GABAARs are the molecular target for a remarkable range of natural products of plant origin. Prominent examples of these plant-derived ligands include the GABAAR-defining antagonist bicuculline (from Dicentra cucullaria), the potent psychoactive agonist muscimol (from the intoxicating mushroom amanita muscaria), and the pore-blocking convulsant picrotoxin (from the Menispermaceae family) [184-186]. Interestingly, trace amounts of diazepam have also been detected in wheat and potato [187]. Given the long history of herbal medicine use in alleviating symptoms associated with CNS-related disorders implicating GABAARs such as anxiety and insomnia, the pharmacological active constituents of these herbs are of great therapeutical interest. Several emerging families of natural products-inspired GABAAR ligands are discussed below.

1.12.1. Flavonoids

Flavonoids are found in great abundance in fruits, vegetables, herbs and beverages such as tea, coffee and red wine. There is tremendous structural diversity among flavonoids,

37 leading to division into eight classes, namely flavones, flavanones, flavonols, flavan-3- ols, flavandiols, dihydroflavonols, isoflavones and anthocyanins (Figure 1.10A). These polyphenolic compounds are associated with health-promoting properties such as anti- oxidant, anti-inflammatory, and anti-tumour [188]. In addition, lipophilic flavonoids are able to cross the blood-brain barrier to influence brain function [189, 190]. Among these neuroactive flavonoids, some modulate GABAARs with high affinity via the benzodiazepine site, and are anxioselective in mice [191, 192]. Subsequently, several synthetic efforts based on the flavonoid templates were undertaken, leading to the rapid expansion of a novel class of benzodiazepine site ligands [191, 193].

Figure 1.10. Natural products of plant origin that modulate GABAARs. (A) Major structural classes of flavonoids. (B) Synthetic flavones and flavan-3-ols with diverse pharmacology at GABAARs. (C) Valerenic acid isolated from Valeriana officinalis. (D) Honokiol and magnolol, bioactive constituents of Magnolia officinalis.

More recently, synthetic flavonoids with pharmacology clearly unrelated to the high- affinity benzodiazepine site have been reported [194]. The anxiolytic flavan-3-ol derivative (2S,3R)-trans-3-acetoxy-4′-methoxyflavan (Fa131; Figure 1.10B) has a triphasic profile at GABAARs in vitro [195]. Mutations that impair anaesthetic sensitivity such as β2N265S and α1M236W reduce the modulatory action of Fa131, suggesting that the flavonoid may bind to the transmembrane anaesthetic sites [196]. A structurally- related flavan-3-ol ester (2S,3S)-cis-3-acetoxy-4′-dimethoxyflavan (Fa173; Figure

1.10B) is a neutralising modulator at GABAARs. While it lacks intrinsic activity, Fa173 is able to selectively block the actions of Fa131, etomidate, loreclezole and the low- affinity diazepam component [196]. Fa173 is the first antagonist with such profile known to date, and is expected to serve as a pharmacological tool in the study of the transmembrane anaesthetic sites, akin to flumazenil for the high-affinity benzodiazepine site. Flavone analogues such as 2′-methoxy-6-methylflavone (2′MeO6MF) and 3- hydroxy-2′-methoxy-6-methylflavone (3-OH-2′MeO6MF; Figure 1.10B) also have anti- anxiety effects in mice mediated via GABAARs in a flumazenil-insensitive manner [197,

198]. These flavones exhibit complex pharmacology in vitro ranging from positive modulation, direct activation and no activity at distinct GABAAR subtypes. Besides that, a series of synthetic isoflavones have been demonstrated to produce flumazenil- insensitive efficacious enhancement at GABAARs in vitro [199].

Clearly, both natural and synthetic flavonoids are capable of influencing GABAAR function by directly binding to the high-affinity benzodiazepines site or the TMD where most anaesthetic sites reside. A flavonoid binding site on GABAARs has also been proposed, but its existence is yet to be systematically investigated. Nonetheless, the drug- like physiochemical properties and anxiolytic-like effects of flavonoids make them a class of attractive lead compounds for drug development. Moreover, the diverse pharmacology

39 of flavonoids represents an opportunity to unravel the complex mechanisms of activation and modulation at GABAARs.

1.12.2. Valerenic acid

Valerenic acid (Figure 1.10C), the major constituent of valerian root (Valeriana officinalis) is an allosteric modulator at GABAARs with anxiolytic and anticonvulsive effects. Functionally, valerenic acid positively modulates, activates and inhibits

GABAARs across its effective concentration range (1–300 µM) [200]. Furthermore, valerenic acid displays an etomidate-like selectivity profile, with more efficacious modulation recorded at β2/3- than β1-containing GABAARs. Consistent with this finding, mutations at β265 residue (β1S265N, β2N265S and β3N265M) were found to affect the activity of valerenic acid in vitro and in vivo [200, 201]. With the use of homology modelling, molecular docking, pharmacophore and site-directed mutagenesis, Luger et al. (2015) identified residues critical for the activity of valerenic acid on the β3 subunit, some of which correspond to the established etomidate/propofol binding residues, suggesting that valerenic acid also binds to the cavities in the transmembrane β+α‒ interfaces [202]. However, as other homologous interfacial sites were not investigated, the possibility of other binding pockets cannot be excluded. There is also interest in the clinical application of valerenic acid as an anxiolytic or anticonvulsant, with several lead optimisation attempts made to improve its pharmacokinetic and pharmacodynamic parameters [203, 204].

1.12.3. Honokiol and magnolol

The bark of Magnolia officinalis is used in traditional Chinese and Japanese medicine to treat insomnia and anxiety. Honokiol and magnolol (Figure 1.10D) are the pharmacologically active constituents with a variety of enzymes and receptors proposed to be the site of action [205]. GABAARs are thought to be one of the most plausible targets, on the basis of their remarkably similar properties to the benzodiazepines such as sedative, anxiolytic, muscle relaxant, anticonvulsant and anaesthetic. Animal behavioural studies appear to support the CNS actions of honokiol and magnolol being mediated via the high-affinity benzodiazepine site, as these effects are sensitive to flumazenil antagonism [206-209]. Electrophysiological studies show that both honokiol and magnolol are able to potentiate native GABAARs found in rat neurons [210, 211]. These compounds also positively modulates heterologously expressed αβ, αβγ and αβδ

GABAARs with no subtype selectivity, thus argues against the α+γ‒ benzodiazepine site as the site of action [211]. The reason for the discrepancy between in vitro and in vivo studies is not clear. Furthermore, mutations in the transmembrane M2, M3 and M3 domains which impair the action of several different classes of anaesthetic agents have insignificant effect on the action of honokiol and magnolol [211]. The binding site(s) for these natural products on GABAARs remain obscure at this stage, but has not hindered the search for structural analogues with improved potency and efficacy. Extensive structure-activity studies have led to a vast array of honokiol and magnolol derivatives, with some of them possessing subtype selectivity and extraordinary efficacy [212-214].

1.13. Thesis rationale

The focus of this thesis revolves around the action of 2′MeO6MF, a synthetic flavonoid, and kavain, a naturally occurring kavalactone at GABAARs.

1.13.1. 2′MeO6MF

The synthetic flavonoid 2′MeO6MF is an allosteric ligand with complex pharmacology at GABAARs [197]. The selective agonistic action of 2′MeO6MF at α2β2/3γ2L

GABAARs is thought to be the reason for its in vivo anxioselectivity. The mechanism(s) behind its unique pharmacological profile, however, is not known. Efforts to identify the specific binding region(s) of 2′MeO6MF have been futile as considerable variability of its action is observed within our laboratory and between different laboratories. We hypothesise that mixed receptor populations, a common problem in the heterologous expression of multimeric receptors, is the culprit. The first project aims to address the variability of 2′MeO6MF activity at α2β3γ2L GABAARs by systematically expressing all possible receptor combinations in Xenopus oocytes using α2, β3 and γ2L subunits, and test 2′MeO6MF function on individual receptor combination using electrophysiological methods. It is expected that data generated by this study will provide hints to explain the variability observed in 2′MeO6MF action.

1.13.2. Kavalactones

The intoxicating pepper plant kava has been used for millennia to treat insomnia and anxiety by the Pacific Islanders. Kava’s bioactive constituents, commonly known as kavalactones (Figure 4.1) are in the limelight due to their efficacy in reducing anxiety in several small-scale clinical trials [215, 216]. There is also an ongoing phase III clinical

42 trial attempting to establish the efficacy and safety of kavalactones in treating generalised anxiety disorder in a larger and longer study [217]. Despite a growing body of clinical evidence in support of kavalactones’ medicinal values, the basic pharmacology of kavalactones remains enigmatic. GABAARs have been proposed due to the frequent association of benzodiazepines with anti-anxiety effects. However, only a number of binding studies have investigated the interaction of kavalactones with GABAARs to date, and the evidence is inconsistent due to the use of kava extracts containing an assortment of chemicals. As such, the second project aims to characterise the function of individual kavalactones at various physiologically relevant human GABAAR subtypes expressed in

Xenopus oocytes. Besides that, it is also our aim to elucidate the mechanism of action of kavalactones with the use of pharmacological tools that target distinct allosteric binding sites at GABAARs. Through this study, we hope to obtain more conclusive proof which will establish the direct interaction of kavalactones with GABAARs.

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198. Karim N, Gavande N, Wellendorph P, 206. Maruyama Y, Kuribara H. Overview Johnston GAR, Hanrahan JR, Chebib of the pharmacological features of M. 3-Hydroxy-2′-methoxy-6- honokiol. CNS Drug Reviews. methylflavone: A potent anxiolytic 2000;6(1):35-44. with a unique selectivity profile at GABAA receptor subtypes. 207. Chen CR, Tan R, Qu WM, Wu Z, Biochemical Pharmacology. Wang Y, Urade Y, et al. Magnolol, a 2011;82(12):1971-83. major bioactive constituent of the bark of Magnolia officinalis, exerts 199. Gavande N, Karim N, Johnston GAR, antiepileptic effects via the Hanrahan JR, Chebib M. GABA/benzodiazepine receptor Identification of benzopyran-4-one complex in mice. British Journal of derivatives (isoflavones) as positive Pharmacology. 2011;164(5):1534-46.

208. Chen C-R, Zhou X-Z, Luo Y-J, Huang relationship. Journal of medicinal Z-L, Urade Y, Qu W-M. Magnolol, a chemistry. 2011;54(15):5349-61. major bioactive constituent of the bark of Magnolia officinalis, induces sleep 213. Fuchs A, Baur R, Schoeder C, Sigel E, via the benzodiazepine site of GABAA Müller CE. Structural analogues of the receptor in mice. Neuropharmacology. natural products magnolol and 2012;63(6):1191-9. honokiol as potent allosteric potentiators of GABAA receptors. 209. Qu W-M, Yue X-F, Sun Y, Fan K, Bioorganic & Medicinal Chemistry. Chen C-R, Hou Y-P, et al. Honokiol 2014;22(24):6908-17. promotes non-rapid eye movement sleep via the benzodiazepine site of 214. Bernaskova M, Schoeffmann A, the GABAA receptor in mice. British Schuehly W, Hufner A, Baburin I, Journal of Pharmacology. Hering S. Nitrogenated honokiol 2012;167(3):587-98. derivatives allosterically modulate GABAA receptors and act as strong 210. Squires RF, Ai J, Witt MR, Kahnberg partial agonists. Bioorganic & P, Saederup E, Sterner O, et al. Medicinal Chemistry. Honokiol and magnolol increase the 2015;23(20):6757-62. number of [3H]muscimol binding sites three-fold in rat forebrain membranes 215. Pittler MH, Ernst E. Kava extract in vitro using a filtration assay, by versus placebo for treating anxiety. allosterically increasing the affinities Cochrane Database of Systematic of low-affinity sites. Neurochem Res. Reviews. 2003;(1):Cd003383. 1999;24(12):1593-602. 216. Sarris J, LaPorte E, Schweitzer I. 211. Alexeev M, Grosenbaugh DK, Mott Kava: A comprehensive review of DD, Fisher JL. The natural products efficacy, safety, and magnolol and honokiol are positive psychopharmacology. Australian and allosteric modulators of both synaptic New Zealand Journal of Psychiatry. and extra-synaptic GABAA receptors. 2011;45(1):27-35. Neuropharmacology. 2012;62(8):2507-14. 217. Savage KM, Stough CK, Byrne GJ, Scholey A, Bousman C, Murphy J, et 212. Taferner B, Schuehly W, Huefner A, al. Kava for the treatment of Baburin I, Wiesner K, Ecker GF, et al. generalised anxiety disorder (K- Modulation of GABAA-receptors by GAD): study protocol for a honokiol and derivatives: Subtype randomised controlled trial. Trials. selectivity and structure–activity 2015;16:493.

Chapter 2

Materials and Methods ______

2.1. Introduction

The expression of recombinant ion channels in heterologous systems such as mammalian cell lines and Xenopus laevis frog oocytes is a critical approach in the study of ligand- gated ion channels. In conjunction with molecular biological and electrophysiological recording techniques, the structure-function relationship of a receptor of interest can be explored under controlled conditions. Defined mutations could also be introduced in these receptors to facilitate the elucidation of the binding site of a ligand of interest.

For the purpose of this thesis, Xenopus oocytes were used to study GABAARs for numerous reasons [1, 2]. First, their large sizes and robust nature allow mRNA microinjection and electrophysiological recording to be conducted relatively easily.

Second, by varying the ratio of the mRNA encoding for different subunits, the subunit composition of GABAARs expressed could be controlled (assuming that all subunits express equally well). Third, the lack of endogenous GABAARs in Xenopus oocytes allows the investigation of the function of a single, defined receptor subtype of interest without the contamination from native receptors. Lastly, as Xenopus oocytes translate exogenous mRNA very efficiently, the amount of mRNA injected could be adjusted to achieve desirable expression levels.

This chapter details the materials and methods used throughout this thesis. A comprehensive list of chemicals, reagents, media, buffers and equipment is provided. The general methodology involving molecular biological techniques, preparation of Xenopus oocytes, two-electrode voltage clamp electrophysiology recording and data analysis is also described in detail.

2.2. Safety, regulations and ethics

All experiments conducted followed the general guidelines stated in the Safety Handbook of Faculty of Pharmacy, University of Sydney. Quarantine Awareness training (endorsed by the Department of Agriculture, Australian Government) and Quarantine Approved

Premises accreditation (endorsed by the Australian Quarantine and Inspection Service,

Australian Government) were obtained prior to the commencement of any procedures which required handling of Xenopus laevis frogs as well as their oocytes. All the procedures involved in the use of Xenopus laevis frogs for the purpose of this thesis were approved by the Animal Ethics Committee of the University of Sydney (Reference number: 2013/5915).

2.3. Materials

2.3.1. Chemicals and reagents

Table 2.1. General chemicals and reagents.

Chemicals/reagents Source

Ambion® Nuclease-free water Thermo Fisher Scientific (Scoresby, VIC, Australia) Ampicillin sodium salt Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) Ethanol (for molecular biology) Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) Gentamicin solution (50 mg/mL) Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) Glycerol (for molecular biology) Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) Mineral oil (for molecular biology) Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) SBYR® Safe DNA Gel Stain Thermo Fisher Scientific (Scoresby, VIC, Australia) Sodium hydroxide (NaOH) pellets Asia Pacific Specialty Chemical Ltd Tetracycline HCl Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) UltraPure™ Agarose Thermo Fisher Scientific (Scoresby, VIC, Australia) 1 kb DNA ladder Genesearch Pty Ltd (Arundel, QLD, Australia)

Table 2.2. Chemicals used in electrophysiological studies.

Chemicals Source

Allopregnanolone Tocris Bioscience (Bristol, UK) Diazepam Apin Chemicals LTD (Abingdon, Oxon, UK) DMSO Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) DS2 Tocris Bioscience (Bristol, UK) Synthesised by Dr. Raja Viswas (Faculty of Pharmacy, University Etomidate HCl of Sydney, NSW, Australia) following protocols described in Janssen et al. [3] and Janssen et al. [4] Flumazenil Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) GABA Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) propofol Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) DL-kavain Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) Synthesised by Dr. Raja Viswas (Faculty of Pharmacy, University 2′MeO6MF of Sydney, NSW, Australia) as described in Karim et al. [5]

2.3.2. Bacterial growth media

The following media were prepared as described and stored at room temperature.

Table 2.3. Media used for bacterial growth.

LB Agar (Lennox L Agar) Terrific Broth 10 g SELECT Peptone 140 11.8 g SELECT Peptone 140 Ingredients 5 g SELECT Yeast Extract 23.6 g yeast extract (amount/L 5 g NaCl 9.4 g dipotassium hydrogen phosphate of media) 12 g SELECT Agar 2.2 g potassium dihydrogen phosphate 4 mL of glycerol (carbon source) 35 g powder/1 L deionised water; 47.6 g powder/1 L deionised water; Preparation autoclaved at 121 °C for 1 hour autoclaved at 121 °C for 1 hour Thermo Fisher Scientific Thermo Fisher Scientific Source (Scoresby, VIC, Australia) (Scoresby, VIC, Australia)

2.3.3. Buffers

The following buffers were used throughout this thesis. Buffers with proprietary ingredients were not included.

Table 2.4. Buffers used for molecular biology purposes.

Buffers Ingredients Source 50 mM potassium acetate, 20 mM tris-acetate, CutSmart® Buffer Genesearch Pty Ltd 10 mM magnesium acetate (1×) (Arundel, QLD, Australia) 100 µg/mL BSA (pH 7.9 at 25 °C). 95 % Formamide 18 mM EDTA Gel Loading Buffer Thermo Fisher Scientific 0.025 % SDS II (Scoresby, VIC, Australia) 0.025 % Xylene Cyanol 0.025 % Bromophenol Blue 15 mM ATP NTP/CAP (2×), 15 mM CTP mMessage Thermo Fisher Scientific 15 mM UTP mMachine® T7 (Scoresby, VIC, Australia) Transcription Kit 2 mM GTP 8 mM cap analog 40 mM Tris-acetate Sigma-Aldrich Pty Ltd TAE Buffer (10×) 5.7 % (v/v) glacial acetic acid (Castle Hill, NSW, 0.01 M EDTA (pH 8.0) Australia)

The recipes of buffers used in the defolliculation, storage and electrophysiological recording of Xenopus oocytes are summarised in Table 2.5. All buffer ingredients were purchased from Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia).

Table 2.5. Buffers used in the handling of Xenopus oocytes.

Buffers (1×) Ingredients ND96 OR2* ND96 (storage)* (electrophysiology) † NaCl 82.5 mM 96 mM 96 mM KCl 2 mM 2 mM 2 mM

MgCl2 1 mM 1 mM 1 mM

CaCl2 – 1.8 mM 1.8 mM HEPES – – 5 mM HEPES hemisodium 5 mM 5 mM – Theophylline – 0.5 mM – Na pyruvate – 2.5 mM –

* Filtered with the Stericup® Filter Units (Merck Millipore, Bayswater, VIC, Australia). † pH was adjusted to 7.4 with NaOH using a benchtop pH meter.

2.3.4. Enzymes, competent cells and kits

The following enzymes, competent cells and kits were used for the purpose of this thesis.

Table 2.6. Enzymes, competent cells and kits.

Source Boehringer Mannheim Australia Collagenase A Pty Ltd (North Ryde, NSW, Enzymes Australia) Restriction enzymes Genesearch Pty Ltd (Arundel, (HpaI, NheI-HF®, NotI-HF®, SmaI) QLD, Australia) Genesearch Pty Ltd (Arundel, NEB 10-beta Competent QLD, Australia) E. coli cell Thermo Fisher Scientific lines TOP10/P3 (Scoresby, VIC, Australia) mMessage mMachine® T7 Thermo Fisher Scientific Kits Transcription (Scoresby, VIC, Australia)

Qiagen (Chadstone, VIC, QIAprep® Spin Miniprep Australia) Qiagen (Chadstone, VIC, QIAquick® Nucleotide Removal Australia) Qiagen (Chadstone, VIC, QIAquick® PCR Purification Australia) QuikChange II Site-Directed Agilent Technologies Australia Mutagenesis (Mulgrave, VIC, Australia)

2.3.5. General apparatus and equipment

Table 2.7. General apparatus and equipment.

Equipment Source Autoclave Atherton (Thornbury, VIC, Australia) Block heater (Stuart) Bibby Scientific Ltd (Staffordshire, OSA, UK) Thermo Fisher Scientific (Scoresby, VIC, Centrifuge (Eppendorf, 5417R) Australia) Drummond Scientific glass capillaries Thermo Fisher Scientific (Scoresby, VIC, (OD: 1.4 mm, ID: 0.53 mm, l: 3.5′′)* Australia) GeneClamp 500B Amplifier ADInstruments, Sydney, NSW, Australia Harvard Apparatus glass capillaries SDR Scientific (Chatswood, NSW, Australia) (OD: 1.2 mm, ID: 0.94 mm, l: 100 mm)* Microprocessor-controlled vertical World Precision Instruments, Inc. (Sarasota, pipette puller (PUL-100) FL, USA) Microscopes (SZ-61) Olympus (North Ryde, NSW, Australia) Thermo Fisher Scientific (Scoresby, VIC, NanoDrop 1000 Spectrophotometer Australia) Nanoliter 2000 microinjection system Drummond Scientific (Broomali, PA, USA) PowerLab 2/25 ADInstruments, Sydney, NSW, Australia Bio-Rad Laboratories Pty Ltd (Gladesville, PowerPack™ Basic Power Supply NSW, Australia) Shaking incubators Labec (Marrickville, NSW, Australia) Single-stage glass microelectrode puller Narishige (Tokyo, Japan) (PP-830) Stericup® Filter Units Merck Millipore (Bayswater, VIC, Australia) Bio-Rad Laboratories Pty Ltd (Gladesville, Thermal Cycler (T100™) NSW, Australia) Warner OC-725C Oocyte Clamp Warner Instrument Corp, Hamden, CT, USA

* OD: outside diameter, ID: inside diameter, l = length

2.3.6. GABAAR subunits

A total of 10 different GABAAR subunits (α1–5, β1–3, γ2L and δ) were used in this thesis, and their origins, subcloned plasmid vectors, sources and the chapters they appeared in are summarised in Table 2.8.

Table 2.8. A summary of all GABAAR subunits used in this study. GABA R UniProt Restriction A Origin Plasmid vector Source Chapter subunit Entry enzyme α1 Human P14867 pCDM8 NotI-HF † 4 α2 Human Q8TBI4 pCMV6-XL5 SmaI ‡ 3 α4 Human P48169-1 pCDNA1/Amp HpaI § 4 α5 Human P31644-1 pCDNA3 NotI-HF § 4 β1 Human* P18505-1 pCDM8 HpaI § 4 β2 Human P47870-1 pCDM8 NotI † 4 β3 Human P28472-1 pGEMHE NheI-HF § 3, 4 γ2L Human P18507-2 pCDM8 NotI-HF † 3, 4 δ Human O14764-1 pCDNA1/Amp HpaI § 4

* The β1 subunit is of the human origin, but has a bovine signal sequence. † Gift from Dr. Paul Whiting (Merck Sharpe and Dohme, Harlow, UK). ‡ Purchased from OriGene Technologies, Rockville, MD, USA. § Gift from Professor Bjarke Ebert (H. Lundbeck A/S, Alby, Denmark).

2.3.7. Oligonucleotide primers

Oligonucleotide primers used in site-directed mutagenesis reactions were synthesised by

Sigma-Aldrich Pty Ltd (Castle Hill, NSW, Australia) and Thermo Fisher Scientific

(Scoresby, VIC, Australia). The sequences and other information about specific primers are provided in the relevant sections.

2.4. Methods

2.4.1. Molecular biology

2.4.1.1. Transformation of plasmid DNA into E. coli

Desired wild-type or mutant plasmid DNAs were transformed into appropriate competent

E. coli cell lines. TOP10/P3 cells were used for subunits subcloned into the pCDM8 vector (α1, β1, β2 and γ2L subunits), whereas the NEB 10-beta competent cells were used for other subunits (Table 2.8). A tube containing 50 µL of competent cells was thawed on ice before the addition of 1–5 µL containing 100–300 ng of plasmid DNA. After 30 minutes of incubation on ice, the cell mixture was heat-shocked at 42 ºC on a block heater for 45 seconds, and then placed on ice for another 10 minutes. Following that, 500 µL of

Terrific Broth was added to the cell mixture, and the tube was then incubated at 37 ºC in a shaking incubator (220 rpm) for 60 minutes. The cell mixture was mixed thoroughly after incubation before they were spread on LB Agar plates containing the appropriate antibiotics (TOP10/P3: 100 µg/mL ampicillin + 10 µg/mL tetracycline; NEB 10-beta:

100 µg/mL ampicillin only). The plates were air dried, inverted and incubated at 37 ºC overnight (12–18 hours, depending on growth rate).

2.4.1.2. Growth of E. coli cultures

For transformed NEB 10-beta cells, a single bacterial colony was picked from the LB

Agar plate using a pipette tip which was then dropped into a 50 mL Falcon tube containing

5 mL Terrific Broth supplemented with 100 µg/mL ampicillin. The inoculated media was then incubated at 37 ºC in a shaking incubator (220 rpm) overnight (≥ 16 hours, depending on growth rate). For transformed TOP10/P3 cells, due to the relatively low yields of plasmid DNA, inoculated Terrific Broth was incubated at 37 ºC with shaking at 220 rpm

64 for 1–2 hours to initiate growth before the addition of antibiotics (100 µg/mL ampicillin

+ 10 µg/mL tetracycline). Incubation continued at 37 ºC with shaking at 220 rpm overnight (≥ 16 hours).

2.4.1.3. Bacterial glycerol stocks

For long-term storage of plasmids, bacterial glycerol stocks were created and stored at -

80 ºC. Glycerol is a cryoprotectant that helps with cell viability by protecting bacteria from freezing damage. Bacterial glycerol stocks were created by mixing 500 µL of overnight cultures with 500 µL of 50 % glycerol in a 2 mL screw top tube, snap-frozen in a dry ice and ethanol bath, and then stored at -80 ºC.

2.4.1.4. Purification, quality assessment and quantification of plasmid DNA

Plasmid DNA was isolated and purified using the QIAprep Spin Miniprep Kit following the manufacturer’s instructions. Briefly, bacteria in overnight cultures were pelleted by centrifugation at 10, 000 rpm for 3 minutes using the Eppendorf Centrifuge at room temperature. Pelleted bacteria were resuspended in 250 µL of Buffer P1 (supplemented with RNase A to hydrolyse RNA, and thus prevent co-purification of RNA with plasmid

DNA) before being lysed by the addition of 250 µL of Buffer P2 (contains NaOH and

SDS). SDS helps cell lysis by solubilising phospholipid and proteins of cell membrane, whereas the strong base NaOH denatures chromosal and plasmid DNAs, as well as proteins released from the bacteria. 350 µL of Buffer N3 (contains guanidinium chloride and acetic acid; pH 4.3) was then added to neutralise the alkaline conditions of the lysate, allowing the macromolecules to renature. The high salt concentrations, however, causes the larger chromosal DNA, proteins, debris and SDS to aggregate and precipitate out of

65 the solution, while the small plasmid DNA renatures properly and stays soluble in the solution. The precipitant was pelleted by centrifugation at 14, 000 rpm for 10 minutes, and the clear supernatant containing the plasmid DNA was applied to the spin column where selective adsorption of the plasmids occur at the silica-gel membrane in high-salt conditions. 750 µL of Buffer PE (contains ethanol) was then added to the spin column to remove salts, and the column was spun twice to ensure no residual washing buffer remained. 30–50 µL of Buffer EB (10 mM Tris.Cl, pH 8.5) was used to elute the plasmid

DNA from the spin column.

The presence of purified plasmid DNAs was confirmed using agarose gel electrophoresis.

Agarose gels of 1 % (w/v) were made by dissolving 0.50 g UltraPure™ Agarose powder in 50 mL of 1× TAE buffer with 5 µL of SBYR® Safe DNA Gel Stain. DNA samples were mixed at a 1:1 ratio with Gel Loading Buffer II. A 1 kb DNA ladder was run alongside the DNA samples for each gel. Electrophoresis was performed at 100 V and

400 mA using the PowerPac™ Basic Power Supply for 30–45 minutes. DNA bands were visualised using the Gel Doc™ EZ System and the Image Lab™ software. Clear and defined bands on the gel indicate the presence of high quality plasmid DNA (Figure

2.1A). The quality and quantity of the plasmid DNA were further analysed using the

NanoDrop 1000 Spectrophotometer. The spectrophotometer measures the concentration of DNA (ng/µL) in a sample, besides providing information on sample purity in the form of ratio of sample absorbance at a wavelength of 260 nm and 280 nm (260/280), as well as 260/230 (Figure 2.1B). DNA samples with a ratio of ~1.8 for 260/280 and 1.8–2.2 for

260/230 are generally accepted as “pure”.

Figure 2.1. The quality assessment and concentration measurement of purified plasmid DNA. (A) An example of agarose gel electrophoresis of purified DNA from E. coli. Lane 1: 1 kb DNA ladder; 2: purified plasmid DNA of GABAAR β3 subunit; 3: linearised plasmid DNA of GABAAR β3 subunit. The two bands in lane 2 correspond to the two topologically-different forms of circular plasmid DNAs. The supercoiled form travels faster and is found further down the gel compared to the nicked, relaxed circle form. (B) The quantification of plasmid DNA samples using the NanoDrop 1000 Spectrophotometer. The sample measured contained plasmid DNA of GABAAR α2 subunit. The concentration of DNA measured was 154.8 ng/µL and the sample purity was confirmed by the 260/280 and 260/230 ratios (inset) within the recommended range (see section 2.4.1.5).

2.4.1.5. Verification of purified plasmid DNA identities

The identities of all wild-type and mutated plasmid DNAs were verified by sequencing the entire gene of interest using appropriate primers. Samples containing 600–1500 ng purified DNA and 0.8 pmol/µL primers were made up to 12 µL with nuclease-free water, and then sent to Sanger Sequencing, Australian Genome Research Facility Ltd

(Westmead, NSW, Australia) for sequencing. Raw traces (.ab1 files) were read using the

CodonCode Aligner software (version 4.0.4 demo, CodonCode Corporation) and translated to amino acid sequences using the online ExPASy translate tool. The amino acid sequences of DNA samples were then compared against the corresponding protein sequences found in the UniProt database using the online Clustal Omega multiple sequence alignment program.

2.4.1.6. Linearisation of plasmid DNA

Circular plasmid DNA was linearised using restriction enzymes to allow for the production of mRNA transcripts derived from the insert GABAAR subunit sequences only

(without the contamination from vector sequences). Briefly, 1–3 µg of plasmid DNA was treated with 1–2 U of restriction enzymes (Table 2.8) and 1× CutSmart® Buffer in a final reaction volume of 50 µL. All reactions were carried out on a Thermal Cycler at 37 ºC for 2 hours, except for when SmaI enzyme was used (25 ºC). Linearised DNA was purified using the QIAquick® Nucleotide Removal Kit. Briefly, 5 volumes of Buffer PNI were added to 1 volume of the reaction mixture, transferred to a spin column which was then centrifuged at 6000 rpm for 1 minute. The flow-through was discarded, and the spin column was washed with 750 µL of Buffer PE, followed by centrifugation at 6000 rpm for 1 minute. The flow-through was discarded, and the spin column was centrifuged at

13,000 rpm for another minute to remove any residual buffer. The linearised DNA was then eluted from the column to a clean 1.5 mL Eppendorf tube using 30–50 µL of Buffer

EB. To ensure complete cleavage of the circular DNA, samples were analysed using agarose gel electrophoresis. A clear, defined band indicates that all circular DNA has been cut (Figure 2.1A).

2.4.1.7. Synthesis, quality assessment and quantification of mRNA

The mMessage mMachine® T7 Transcription Kit was used for the synthesis of capped

RNA of a GABAAR subunit of interest. The term ‘mRNA’ was used instead of ‘capped

RNA’ in this thesis. The components and the required amounts for a transcription reaction is summarised in Table 2.9. A full reaction was used to make wild-type receptor subunit mRNA, whereas a half reaction was used for mutant mRNA.

Table 2.9. mRNA transcription setup.

Amount Component Full reaction Half reaction 2× NTP/CAP 10 µL 5 µL 10× Reaction Buffer 2 µL 1 µL Linearised DNA (50–200 ng/µL) 6 µL 3 µL T7 enzyme mix 2 µL 1 µL Total reaction volume 20 µL 10 µL

The transcription reaction was carried out on a Thermal Cycler at 37 ºC for at least 3 hours to ensure high yields. At the end of the reaction, 1 µL TURBO DNase was added to the mixture, followed by 15 minutes of incubation at 37 ºC to remove the template

DNA. The lithium chloride (LiCl) precipitation method was used to recover and purify mRNA. First, the reaction mixture was mixed thoroughly with 30 µL of LiCl Precipitation

Solution, and was stored at -20 ºC overnight. Then, the mRNA was pelleted by centrifugation (14, 000 rpm) at 4 ºC for 15 minutes. The supernatant containing unincorporated nucleotides and proteins was removed, and the mRNA pellet was washed with 500 µL of 70 % ethanol. The mixture was then re-centrifuged (4 ºC; 14, 000 rpm;

15 minutes), and the ethanol was removed. The ethanol wash step was repeated twice to maximise removal of unincorporated nucleotides. The mRNA pellet was resuspended in

10–20 µL of nuclease-free water when all residual ethanol had been removed.

The quality of mRNA was assessed using agarose gel electrophoresis as described in section 2.4.1.5, except the mRNA sample (+ Gel Loading Buffer II) was denatured at 94

ºC on a block heater before gel loading. Intact mRNA samples appeared as clear, distinct bands on the gel, whereas degraded samples produced a smear. The quantification of purified mRNA samples involved the use of NanoDrop 2000 Spectrophotometer (see section 2.4.1.5). The amount of mRNA in a sample is measured in ng/µL, and the purity

69 is indicated with 260/280 and 260/230 ratios. A 260/280 ratio of ~2.0 and a 260/230 ratio of 2.0–2.2 are generally accepted as pure.

Figure 2.2. The quality assessment and concentration measurement of synthesised mRNA. (A) An example of agarose gel electrophoresis of intact mRNA samples. Lane 1: 1 kb DNA ladder; 2 and 3: GABAAR β3 subunit mRNAs. (B) An example of agarose gel electrophoresis of degraded mRNA samples which appeared as smeared bands in lane 2 and 3. (C) The quantification of mRNAs using the NanoDrop 1000 Spectrophotometer. The sample measured contained mRNA of GABAAR α1 subunit. The concentration of mRNA measured was 1085.3 ng/µL and the sample purity was confirmed by the 260/280 and 260/230 ratios (inset) within the recommended range (see section 2.4.1.7).

2.4.1.8. Site-directed mutagenesis

Single amino acid substitution mutations were introduced to various GABAAR subunits of interest using the QuikChange II Site-Directed Mutagenesis Kit. To create a desired mutation, a forward primer containing the altered sequence positioned in the middle of approximately 30 wild-type nucleotide bases was designed. A reverse primer which has a reverse complement sequence to that of the forward primer was also designed to cover the opposite DNA strand. The forward and reverse primers sequences used to create specific mutations are provided in relevant chapters.

The mutagenesis reaction was carried out on a Thermal Cycler, and involves three major steps: denaturation at 95 ºC to separate plasmid DNAs into single strands; annealing at

55 ºC to allow for the primers to attach to the single DNA strands; and primer extension by DNA polymerase at 68 ºC. The reaction mixtures were assembled as described in

Table 2.10, and cycled using parameters detailed in Table 2.11. Following temperature cycling, 1 µL of Dpn I restriction enzyme (10 U/µL) was added to each reaction mixture, further incubated at 37 ºC for 1 hour to digest the parental (non-mutated) plasmid DNAs.

The amplified mutant plasmid DNA was purified using the QIAquick® PCR Purification

Kit. Briefly, 5 volumes of Buffer PB were added to 1 volume of the reaction mixture, transferred to a spin column which was then centrifuged at 13, 000 rpm for 1 minute. The flow-through was discarded, and the spin column was washed with 750 µL of Buffer PE, followed by centrifugation at 13, 000 rpm for 1 minute. The flow-through was discarded, and the spin column was centrifuged at 13,000 rpm for another minute to remove any residual buffer. The mutant DNA was then eluted from the column to a clean 1.5 mL

Eppendorf tube using 30 µL of Buffer EB. The purified mutant DNA was then transformed into E. coli (section 2.4.1.1), and steps 2.4.1.2–2.4.1.7 were repeated to obtain mutant subunit mRNA.

Table 2.10. Site-directed mutagenesis reaction setup.

Component Amount 10× Reaction Buffer 5 µL Plasmid DNA (5–50 ng) X µL Forward primer (125 ng) X µL Reverse primer (125 ng) X µL dNTP mix 1 µL Nuclease-free water to 50 µL PfuUltra HF DNA polymerase (2.5 U/µL) 1 µL

Table 2.11. Cycling parameters for the site-directed mutagenesis reactions.

Segment Cycles Temperature Time Step 1 1 95 ºC 30 seconds Denaturation 95 ºC 30 seconds Denaturation 2 16 55 ºC 1 minute Annealing 68 ºC 1 minute/kb of plasmid length Extension

2.4.2. Xenopus oocytes: harvesting, defolliculation and microinjection

Surgical harvesting

Before surgery, mature female Xenopus laevis frogs were anaesthetised with 0.17 % tricaine (buffered with 0.06 % sodium bicarbonate) for 12 minutes. The loss of righting reflex was confirmed before transferring on to ice where surgeries were performed. A small (1–2 cm) abdominal incision was made through both the skin and muscle layer with surgical knives. Ovary lobes were removed with a pair of forceps, and kept in OR2 buffer.

The skin and muscle layers were sutured separately, and frogs were allowed to recover for six months before they were reselected for surgeries. A total of five recoverable surgeries were allowed on each frog, before a terminal surgery was performed, in which a lethal dose of tricaine (0.5 %) was used.

Defolliculation

Removed ovary lobes were divided into small sections with the use of scissors, which were then treated with 2 mg/mL Collagenase A (dissolved in OR2 buffer) in a shaking incubator for an hour to defolliculate the oocytes. The concentration of Collagenase A was then diluted to 1 mg/mL, and the cells were checked at 15-minute intervals until defolliculation was complete. The oocytes were then rinsed in OR2 followed by ND96

72 storage solutions to remove Collagenase A and dead cells. Healthy-looking stage V–VI oocytes were isolated and kept in ND96 storage buffer at 18 °C until ready for injection.

Microinjection

To create injection micropipettes, Drummond Scientific borosilicate glass capillaries were pulled apart using a microprocessor-controlled vertical pipette puller with two pulling stages (Stage 1: Heat = 560, Pull = 0, Trip = 130, Delay = 40; Stage 2: Heat =

580, Pull = 0, Trip = 80, Delay = 40), and then blunted with a spatula to produce a tip with an approximate diameter of 10–20 µm. The micropipettes were filled with mineral oil using a syringe, ensuring no air bubbles were introduced in the process. Filled micropipettes were then placed into the injector of the Nanoliter 2000 microinjection system. Various combinations of GABAAR subunit mRNAs were mixed to different ratios and diluted accordingly to a desired concentration as stated in relevant sections.

The mRNA mixture was taken up into the micropipette, then injected into the cytoplasm of oocytes (50 nL/injection) using the Nanoliter 2000 microinjection system. Injected oocytes were stored in ND96 storage solution supplemented with 50 µg/mL gentamycin and 50 µg/mL tetracycline, and incubated with continuous shaking at 18 °C.

2.4.3. Two-electrode voltage-clamp technique

Two-electrode voltage clamp recordings were performed on oocytes 2–4 days post- injection at room temperature (20–25 °C) using a GeneClamp 500B amplifier or a Warner

OC-725C Oocyte Clamp. All experiments were conducted at a holding potential of –60 mV unless otherwise stated, and data were acquired with a PowerLab 2/25, and LabChart

(version 5.0.2) software. The recording microelectrodes (0.2–1.1 MΩ) were generated

73 from Harvard Apparatus glass capillaries using the single stage microelectrode puller

(heater level = 68) and were filled with 3 M KCl solution. The perfusion system was built using a series of polycarbonate, high-density polyethylene two-way stopcocks (Qosina,

Edgewood, NY, USA), connected through tubing to the recording chamber (100 µL), where the oocyte was continuously perfused with ND96 recording solution at an approximate rate of 5 mL/min.

GABA, etomidate HCl and ZnCl2 were dissolved in ND96 recording solution, whereas

2′MeO6MF, kavain, allopregnanolone, DS2, propofol, diazepam and flumazenil were dissolved in DMSO to make up stock solutions of desired concentrations. GABA stock solutions were prepared fresh on the day of experiments. When working with chemicals dissolved in DMSO, all perfusates were standardised to contain 0.8 % DMSO, which did not produce any alteration in the recordings. Perfusates containing these chemicals were made up fresh before each application. The concentration-response curves of agonists were obtained by exposing oocytes to increasing concentrations of agonists until the currents peaked. Maximal GABA current responses were used as controls to allow for comparisons between different oocytes. Depending on the concentration and the solubility of the chemicals applied on oocytes, the wash-out periods ranged from 3 to 15 minutes to allow time for receptors to recover from desensitisation. All experiments were performed using at least two different batches of oocytes.

2.4.4. Data analysis

Prism (version 5.04; GraphPad Software, La Jolla, CA, USA) was used for data analysis. Raw data from GABA concentration-response experiments were fitted to either a monophasic (1) or biphasic (2) Hill equation, and the fitted maximal values from the preferred model (Extra sum-of-squares F test) were used for normalisation of each dataset.

Imax I  ((logEC  [ A])n ) …………………………….……... (1) 110 50 H

I max  Frac I max (1 Frac) I  ((logEC [ A])n )  ((logEC [ A])n ) …….. (2) 110 50_1 H_1 110 50_ 2 H_2

Concentration-response curves of other agonists such as 2′MeO6MF and etomidate were constructed in a similar manner, except the data were normalised maximal GABA current responses as stated in the relevant chapters. In these equations, Imax represents the maximal agonist response, [A] represents the agonist concentration, EC50 represents the agonist concentration required to activate 50 % of the maximal response (there are two

EC50s for a biphasic curve), nH represents the Hill slope of the fitted curve (there are two

Hill slopes for a biphasic curve), and Frac represents the proportion of the maximal response mediated by the more potent component in a biphasic curve. In all cases, the bottom was constrained to 0, and when fitting data to the biphasic model, the Hill slopes were constrained to 1.

To illustrate the modulatory effect of a chemical of interest, the data were normalised and expressed as shown in equation 3, where I represents the current responses elicited by the

75 co-application of GABA and modulator, IGABA represents the amplitude of the control

GABA-elicited responses. Alternatively, the modulatory effect was expressed as the fold of potentiation, as shown in equation 4.

I  I GABA ………. (3) IGABA

I …………….. (4) IGABA

When comparing parameters across different groups (receptors subtypes/injection ratios), the means of the values obtained from individual dataset were analysed using either unpaired Student’s t test (comparing means for two groups) or one-way ANOVA with

Tukey’s post hoc test (comparing every mean with every other mean in three or more groups). The ANOVA Dunnett’s test was used when comparing every mean in three or more groups to a control mean. Statistical significance was attained at p < 0.05.

References

1. Tapper AR, George AL, Jr. Heterologous hypnotic agent in rats. Arzneimittel- expression of ion channels. Methods in Forschung. 1971;21(8):1234-43. Molecular Biology. 2003;217:285-94. 4. Janssen CGM, Thijssen JBA, Verluyten 2. Goldin AL. Expression of ion channels in WLM, Heykants JJP. Synthesis of (R)-(+)- Xenopus oocytes. Expression and 3H-etomidate. Journal of Labelled Analysis of Recombinant Ion Channels: Compounds and Radiopharmaceuticals. Wiley-VCH Verlag GmbH & Co. KGaA; 1987;24(8):909-18. 2006. p. 1-25. 5. Karim N, Curmi J, Gavande N, Johnston 3. Janssen PA, Niemegeers CJ, Schellekens GA, Hanrahan JR, Tierney ML, et al. 2'- KH, Lenaerts FM. Etomidate, R-(+)-ethyl- Methoxy-6-methylflavone: a novel 1-(α-methyl-benzyl) imidazole-5- anxiolytic and sedative with subtype carboxylate (R 16659), a potent, short- selective activating and modulating actions acting and relatively atoxic intravenous at GABAA receptors. British Journal of Pharmacology. 2012;165(4):880-96.

Chapter 3

The direct actions of GABA, 2′-methoxy-6- methylflavone and general anaesthetics at β3γ2L GABAA receptors: Evidence for receptors with different subunit stoichiometries ______

3.1. Introduction

2′MeO6MF (Figure 1.10B) is a synthetic flavonoid with anxiolytic effect in mice behavioural studies [1]. In two unconditioned models of anxiety (elevated plus maze and light/dark box tests), 2′MeO6MF (1–10 mg/kg, i.p.) increased exploratory behaviour of mice (indicative of anxiolysis) to similar extent as the standard anxiolytic compound diazepam (1–2 mg/kg, i.p.). However, unlike diazepam which showed sedative and myorelaxant effects within the same dose range, 2′MeO6MF induced sedation only at higher doses (30–100 mg/kg) and had no detectable muscle-relaxing action. The anxiolytic effect of 2′MeO6MF was found to be attenuated in the presence of the non- competitive GABAAR antagonist pentylenetetrazole (PTZ), but was unaffected by the benzodiazepine site-specific antagonist flumazenil. These data suggest that GABAARs may be a possible molecular target for the flavonoid, but the mechanism(s) involved are distinct from the benzodiazepines.

Single-channel recording experiments investigating 2′MeO6MF activity at native receptors in dissociated rat hippocampal neurons confirmed the interaction of 2′MeO6MF with GABAARs. In the absence of any ligand, the flavonoid directly elicited current with

- properties consistent with GABAAR-mediated Cl current. Furthermore, the co- application of a competitor peptide mimicking the intracellular MA helix of the GABAAR

γ2 subunit (residues 381–401) reduced the average current amplitude and open channel probability following 2′MeO6MF application (Figure 3.1). The γ2(381-401) MA peptide has been shown to selectively reduce diazepam potentiation, possibly by competing with the γ2 MA helix to interact with essential proteins involved in ion conductance [2]. It is inferred from these data that γ2-containing GABAARs are at least in part responsible for the 2′MeO6MF-induced conductance observed in neurons.

Figure 3.1. 2′MeO6MF directly activates native γ2-containing GABAARs. The left panel traces show the 2 s single-channel activity in excised inside-out patches from cultured rat hippocampal neurons, and the right panel shows the current amplitude probability histogram obtained from 30 s of continuous activity. (A) Single-channel currents elicited by 100 µM 2′MeO6MF had an average maximum amplitude of 2.8 pA. (B) The γ2 MA peptide reduced the current amplitude and open channel probability. Figure taken from Karim et al. [1].

To determine the subunit requirements for 2′MeO6MF activation, Karim et al. (2012) examined the flavonoid activity at numerous recombinant human GABAAR subtypes consisting of (α1/2/3/5) + (β1/2/3) +/- (γ2L) subunits [1]. 2′MeO6MF exhibited a complex pharmacological profile ranging from positive modulation at all α1-containing and

α2β1γ2L receptors, activation at α2β2/3γ2L receptors and no activity at α3- and α5- containing receptors. However, the activity of 2′MeO6MF at βγ receptors was not investigated. Flumazenil failed to antagonise the direct activation of 2′MeO6MF at

GABAARs, in agreement with mice behavioural studies. The distinct activities of

2′MeO6MF at α2β1γ2L and α2β2/3γ2L receptors resembles that of the general anaesthetic etomidate, it was hypothesised that the flavonoid may work via a similar mechanism. Consistent with this hypothesis, the homologous residues found on position

265 of the β subunit, which are critical determinants of etomidate sensitivity at GABAARs

(the effects of these mutations on etomidate sensitivity are discussed in section 1.11.3), were shown to affect 2′MeO6MF activity. Interchanging the asparagine residue of β2 subunit and the homologous serine residue of β1 subunit resulted in positive modulation of α2β2N265Sγ2L receptors and direct activation of α2β1S265Nγ2L receptors.

The non-selective benzodiazepines such as diazepam, while clinically useful in treating anxiety, produce undesirable sedative and amnestic effects, and are also associated with abuse potential and severe withdrawal symptoms. In contrast, 2′MeO6MF displays clear segregation of anxiolytic and sedation doses in mouse behavioural studies and unique pharmacological profile at GABAARs which is not related to the benzodiazepines. These properties of 2′MeO6MF are deemed more favourable than those of diazepam, and its mechanism(s) of action are a topic of interest in our laboratory. However, in our attempt to replicate the activity of 2′MeO6MF at α2β3γ2L receptors, we found considerable

79 variations in the efficacy of 2′MeO6MF activation which prompted us to investigate the underlying cause(s).

3.2. Results

3.2.1. Variations in the relative efficacy of 2′MeO6MF activation at

α2β3γ2L GABAARs

2′MeO6MF has been previously described as a full agonist at α2β3γ2L GABAARs expressed at a 1:1:10 injection ratio in Xenopus oocytes [1]. We expressed α2β3γ2L receptors at a 3:1:3 injection ratio and recorded pharmacological responses to GABA,

Zn2+, diazepam (1 µM) and flumazenil (10 µM) that were consistent with those previously reported (Figure 3.2 and Table 3.1) [3-5]. In contrast to what was reported by Karim et al. (2012) [1], we found that 300 µM 2′MeO6MF only activated α2β3γ2L receptors 8.2

± 3.6 % of the 3 mM GABA response (n = 5; Figure 3.2E and F). The EC50 of 2′MeO6MF was estimated to be approximately 74 µM (95 % CI: 6.4–850 µM). We hypothesised that the variations in the relative efficacy of 2′MeO6MF measured at α2β3γ2L receptors expressed at different ratios may be due to the presence of mixed receptor populations.

Figure 3.2. Characterisation of α2β3γ2L GABAARs expressed at a 3:1:3 injection ratio. (A) Left panel, Representative traces of α2β3γ2L (3:1:3) GABAARs responses to 3 2+ mM GABA and 100 µM Zn alone. Right panel, Mean holding current of oocytes expressing α2β3γ2L (3:1:3) GABAARs (-93 ± 19 nA; n = 8). (B) Left panel, Continuous traces demonstrating two consecutive applications of control (100 µM GABA) followed 2+ by the co-application of 10 µM Zn with control. Right panel, Modulation of 100 µM GABA responses by 10 µM Zn2+ (n = 9). (C) Left panel, Continuous traces demonstrating two consecutive applications of control (1 µM GABA) followed by the co-application of 1 µM diazepam with control; 1 µM diazepam and 10 µM flumazenil with control; and control. Right panel, Potentiation by 1 µM diazepam of 1 µM GABA responses (n = 6). Representative traces demonstrating (D) GABA current responses from 1 µM to 10 mM and (E) 2′MeO6MF’s direct activation from 1 to 300 µM (red) in comparison to 3 mM GABA response. Concentration-response curves of GABA (black; n = 6) and 2′MeO6MF (red; n = 5) are shown in (F). Data are presented as mean ± SEM. Bars indicate durations of drug application. The holding current values are represented by the dotted lines.

Table 3.1. Characteristics of receptors expressed in various combinations of α2, β3 and γ2L GABAAR subunits.

Subunit Rati mRNA amount Expression Holding n 2′MeO6MF activity n combination o (ng/oocyte) level (nA)a current (nA) α2 ― 28–35 No current -20 ± 7.5 8 No current 5 β3 ― 5 290 ± 67 -1000 ± 420 6 Inverse agonism 5 γ2L ― 20–35 No current -370 ± 41 7 No current 5 α2β3 20:1 5 1100 ± 220 -30 ± 7.0 7 Insignificant activation 5 α2γ2L 1:1 5 No current -42 ± 16 4 No current 4 1:1 5 110 ± 34 -470 ± 88 4 Inverse agonism 4 1:5 5 270 ± 80 -220 ± 60 5 Inverse agonism/activation 5 1:10 5 310 ± 61 -150 ± 13 8 Inverse agonism/activation 8

β3γ2L 1:15 5 490 ± 47 -260 ± 40 30 Activation 30 1:20 5 860 ± 87 -180 ± 31 49 Activation 49 1:50 5 1000 ± 160 -28 ± 6.0 15 Activation 15 1:10 5 1100 ± 100 -15 ± 5.0 21 Activation 21 0 3:1: α2β3γ2L 5 4300 ± 940 -93 ± 19 8 Activation 5 3 a Expression level is expressed as the current responses ± SEM (nA) elicited by 3 mM GABA, except for homomeric and α2γ2L receptors (60 mM GABA).

3.2.2. 2′MeO6MF does not activate binary α2β3 or homomeric α2, β3

and γ2L GABAARs

The presence of αβ receptors when expressing αβγ receptors has been shown to result in submaximal measurement of diazepam modulation [6, 7] as these receptors lack the α+γ‒ interface where diazepam binds. To determine if this was the cause for differences in

2′MeO6MF relative efficacy at α2β3γ2L receptors, we expressed α2:β3 subunit mRNAs at a 20:1 ratio to prevent β3 homomeric receptors from forming, and found that

2′MeO6MF (100 µM) had negligible activity at these receptors, eliciting currents only

1.7 ± 0.35 % of the 3 mM GABA responses (n = 5; Figure 3.3A). Therefore, α2β3 receptors are unlikely to contribute to the much higher 2′MeO6MF relative efficacy previously reported at α2β3γ2L GABAARs.

Figure 3.3. Subunit combinations with detectable function but were not activated by 2′MeO6MF. (A) Representative traces illustrating the robust response elicited by 3 mM GABA, and the lack of activity of 100 µM Zn2+ and 100 µM 2′MeO6MF at α2β3 (20:1) GABAARs (n = 6). (B) The injection of γ2L mRNA at high amounts (28–35 ng/oocyte) resulted in constitutively active channels which were inhibited by 100 µM Zn2+, but were not sensitive to 60 mM GABA, 100 µM etomidate and 100 µM 2′MeO6MF (n = 5). (C) The injection of β3 mRNA (5 ng/oocyte) resulted in the formation of functional receptors. Representative traces of β3 homomeric receptors responses to 1–60 mM GABA (top;), 3–300 µM of 2′MeO6MF and 100 µM Zn2+ (bottom). (D) 2′MeO6MF concentration- response curve for β3 homomeric receptors (n = 5). The efficacy of 2′MeO6MF as an inverse agonist is expressed as a fraction of the inhibited spontaneous current (I) normalised against the holding current (Iholding). Data are presented as mean ± SEM. Bars indicate durations of drug application. The holding current values are represented by the dotted lines.

To determine if homomeric GABAARs are activated by 2′MeO6MF, we injected α2, β3 and γ2L mRNAs alone and examined the effect of 2′MeO6MF at these receptors. We did not detect function with 60 mM GABA and 100 µM 2′MeO6MF in Xenopus oocytes injected with α2 mRNA (n = 5; Table 3.1). γ2L homomeric receptors showed Zn2+- sensitive constitutive activity, but were not responsive to 60 mM GABA and 100 µM

2′MeO6MF (n = 5; Figure 3.3B and Table 3.1). Consistent with previous findings [8, 9], the injection of β3 subunit mRNA resulted in the expression of constitutively active receptors, which are activated by high concentrations of GABA (Figure 3.3C and Table

3.1). In contrast to the direct activation observed at α2β3γ2L GABAARs, 2′MeO6MF inhibited the constitutive activity of β3 homomeric receptors in a concentration- dependent manner, with a maximal inhibition of 54 ± 3.0 % (normalised against oocyte holding current) measured at 300 µM 2′MeO6MF (n = 5; Figure 3.3C and D). The lack of activation by 2′MeO6MF at homomeric receptors demonstrates that these receptors are unlikely to be the cause for the variations observed.

3.2.3. 2′MeO6MF activates β3γ2L GABAARs with variable efficacy

We then determined whether binary α2γ2L or β3γ2L receptors are activated by

2′MeO6MF. Binary α2γ2L receptors showed no function to 60 mM GABA, 100 µM Zn2+ and 100 µM 2′MeO6MF (n = 4; Table 3.1). In contrast, injection of β3 and γ2L mRNAs at a 1:20 ratio to prevent the expression of β3 homomers yielded receptors that responded to GABA. 2′MeO6MF efficaciously activated these receptors, with 100 µM eliciting currents approximately 34 ± 4.0 % of the 3 mM GABA responses (n = 49; Figure 3.4).

Notably, there was a high variability in the relative efficacy of 2′MeO6MF, ranging from as low as 1.0 to 120 % of the 3 mM GABA responses. While the standard error of mean efficacy was only 4.0 %, the standard deviation (SD) remained high (31 %). This

84 variability was observed both between different batches of oocytes and within the same batch. The substantial variability in the relative efficacy of 2′MeO6MF suggests that the

1:20 ratio was not optimal for the uniform expression of β3γ2L receptors.

Figure 3.4. 2′MeO6MF activity at β3γ2L GABAARs. (A) 2′MeO6MF exhibits a complex spectrum of activity across β3γ2L GABAARs expressed at various injection ratios. Representative traces of 3 mM GABA (black) and 100 µM 2′MeO6MF (red) are shown for each ratio. Bars indicate durations of drug application. The holding current values are represented by the dotted lines. (1:1; n = 7), 2′MeO6MF inhibited the constitutive activity of receptors expressed. (1:5; n = 5) and (1:10; n = 10), 2′MeO6MF exhibited mixed agonist and inverse agonist activity. (1:15; n = 30), 2′MeO6MF directly activated receptors expressed efficaciously. (1:20; n = 49), 2′MeO6MF activated β3γ2L (1:20) receptors with variable efficacy. Sample traces of cell 1 and 2 were taken from a simultaneous experiment conducted on two different oocytes injected at the same time. (1:50; n = 15) and (1:100; n = 21), 2′MeO6MF showed activation with low efficacy at these receptors. (B) Mean efficacy of 100 µM 2′MeO6MF direct activation at β3γ2L (1:15), (1:20), (1:50) and (1:100) GABAARs. Data are normalised to the 3 mM GABA response. The mean efficacy of 100 µM 2′MeO6MF at various ratios was compared using Tukey’s test, and the significance levels are indicated with n.s. (not significant) and *** (p ≤ 0.001).

To establish the optimal ratio(s) for the uniform expression of β3γ2L GABAARs, we varied the β3:γ2L mRNAs ratio (1:1, 1:5, 1:10, 1:15, 1:50 and 1:100) while keeping the total amount of mRNA constant at 5 ng in each oocyte. We then measured the relative efficacy of 100 µM 2′MeO6MF and compared it across these receptors. At 1:1 ratio,

β3γ2L receptors were constitutively active and 2′MeO6MF acted as an inverse agonist

(Figure 3.4A). This is likely due to the presence of a substantial amount of β3 homomer subpopulation. At 1:5 and 1:10 ratios, where the relative amounts of γ2L mRNA were greater than β3 mRNA, 2′MeO6MF displayed a mixed inverse agonist and agonist effect

(Figure 3.4A), indicating that the pharmacological responses of 2′MeO6MF observed at these ratios were still confounded by β3 homomers.

We increased the injection ratio to 1:15, and consistently observed activation with 100

µM 2′MeO6MF (Figure 3.4A). The 2′MeO6MF-elicited currents were 87 ± 4.5 % of the

3 mM GABA current responses (n = 30), which was significantly higher than that at 1:20 ratio (p ≤ 0.001; Tukey’s test; Figure 3.4B). While the low-efficacy component of

2′MeO6MF was not observed at 1:15 ratio, there was still considerable variability in the relative efficacy measured (SD = 25 %). Increasing the injection ratios further to 1:50 and

1:100 abolished the high-efficacy component of 2′MeO6MF. 100 µM 2′MeO6MF showed efficacy around 6.7 ± 1.2 % (n = 15) at 1:50 ratio and 5.1 ± 0.72 % (n = 21) at

1:100 ratio, which is significantly lower than at the 1:15 ratio (p ≤ 0.001; Tukey’s test;

Figure 3.4B). The relative efficacy of 2′MeO6MF measured at α2β3γ2L receptors was similar to that of β3γ2L receptors expressed at 1:50 and 1:100 ratios (p > 0.5; Dunnett’s test), but was significantly lower than 1:15 (p ≤ 0.001) and 1:20 (p ≤ 0.5) ratios.

The extensive variability in 2′MeO6MF relative efficacy measured at 1:20 ratio was likely to be the result of mixed stoichiometries being expressed. This is further supported by variability observed in the GABA concentration-response relationships at β3γ2L (1:20) receptors, in which out of 13 datasets, 6 were better fitted to a biphasic Hill equation, whereas a simpler monophasic model was preferred to describe the rest of the datasets

(Figure 3.5). Under our conditions, the 1:15, 1:50 and 1:100 ratios were most optimal to express the different stoichiometries of β3γ2L receptors, allowing us to measure

2′MeO6MF responses more reliably. As such, these ratios were chosen for detailed pharmacological characterisation.

Figure 3.5. GABA concentration-response curves of β3γ2L GABAARs expressed at 1:20 ratio. The experiment was conducted on 13 Xenopus oocytes expressing β3γ2L (1:20) receptors, and 6 of the concentration-response curves were found to be better fitted to a biphasic Hill equation (left panel), whereas the other 7 datasets prefer a simpler monophasic model (middle panel). The unique code for each experiment (ddmmyy-cell number) was indicated. The averaged GABA concentration-response curve was significantly better fitted to a biphasic model (p < 0.05; extra sum-of-squares F test; right panel). EC50_1 = 275 µM; EC50_2 = 17.1 mM; Data are presented as mean ± SEM.

3.2.4. β3γ2L GABAARs of different subunit stoichiometries vary in

pharmacological properties

To obtain an understanding of the properties of different β3γ2L receptor stoichiometries, we assessed constitutive activity, 2′MeO6MF and GABA potencies of the receptors expressed at 1:15, 1:50 and 1:100 injection ratios. Oocytes expressing β3γ2L (1:15) receptors exhibited significantly larger, more negative holding currents (-260 ± 40 nA) in comparison to 1:50 (-28 ± 6.0 nA) and 1:100 (-15 ± 5.0 nA) ratios (p ≤ 0.001; Tukey’s

2+ test; Figure 3.6A). The application of 100 µM Zn generated a reduction in inward current at β3γ2L (1:15) receptors (Figure 3.6B). In contrast, 100 µM Zn2+ did not have any effect at β3γ2L (1:50) and (1:100) receptors.

Figure 3.6. Characterisation of β3γ2L GABAARs expressed at 1:15, 1:50 and 1:100 ratios. The level of constitutive activity is indicated by (A) holding current of injected 2+ oocytes and (B) the inhibition of baseline current by 100 µM Zn . (A) β3γ2L GABAARs expressed at 1:15 ratio showed significantly larger holding current (-260 ± 40 nA; n = 30) than at 1:50 (-28 ± 6.0 nA; n = 15) and 1:100 (-15 ± 5.0 nA; n = 21) ratios (p ≤ 0.001; Tukey’s test). (B) Representative traces demonstrating current responses of 3 mM GABA, 100 µM 2′MeO6MF and 100 µM Zn2+. At 1:15 ratio, receptors were sensitive to

89 the inhibition of 100 µM Zn2+ (reduction in inward current; n = 10). Zn2+ did not have any effects at 1:50 (n = 7) and 1:100 (n = 8) ratios. (C) Representative traces demonstrating 2′MeO6MF’s direct activation from 1 to 300 µM in comparison to 3 mM GABA response. (D) 2′MeO6MF concentration-response curves of β3γ2L (1:15; n = 8), (1:50; n = 7) and (1:100; n = 6) GABAARs. Data are normalised to the 3 mM GABA response. (E) Representative traces of GABA current responses from 1 µM to 30 mM at β3γ2L (1:15), (1:50) and (1:100) GABAARs. (F) GABA concentration-response curves of β3γ2L (1:15; n = 6), (1:50; n = 8) and (1:100; n = 9) GABAARs. Data are presented as mean ± SEM. Bars indicate durations of drug application. The holding current values are represented by the dotted lines.

Despite the difference in maximal efficacy (Figure 3.6C), 2′MeO6MF showed similar potency at all ratios (EC50 = 45 µM (1:15), 62 µM (1:50), 63 µM (1:100); p > 0.05 for all logEC50 comparisons; Tukey’s test; Figure 3.6D and Table 2). The potency of

2′MeO6MF measured at α2β3γ2L receptors is also not significantly different from β3γ2L receptors at all three ratios (p > 0.05 for all comparisons; Dunnett’s test). We then measured the GABA potency, and found that β3γ2L (1:15) receptors displayed a monophasic GABA concentration-response curve with low potency and a shallow Hill slope (EC50 = 1.4 mM; nH = 0.54; n = 6; Figure 3.6E, F and Table 3.2). In contrast, the data of 1:50 and 1:100 ratios were significantly better fitted to a biphasic Hill equation than a monophasic model (p < 0.0001 for both ratios; extra sum-of-squares F test; Figure

3.6E and F). At 1:50 ratio, GABA has a high-sensitivity component with an EC50 of 63

µM and a low-sensitivity component with an EC50 of 5.5 mM. The mean EC50 values for

β3γ2L (1:100) receptors were very similar to those of β3γ2L (1:50) receptors (EC50_1: 64

µM; EC50_2: 4.0 mM). β3γ2L GABAARs expressed at 1:50 and 1:100 ratios exhibit nearly identical properties (no statistical significant differences in all aspects; p > 0.05), but are very different in comparison to the receptors expressed at 1:15 ratio.

Table 3.2. GABA, 2′MeO6MF, etomidate and propofol concentration-response curve parameters at β3γ2L (1:15), (1:50) and (1:100) GABAARs derived from curve- fitting procedures.

β3γ2L (ratio)a

1:15 1:50 1:100 c Imax ± SEM 780 ± 220 nA 1000 ± 250 nA 1000 ± 97 nA

EC50_1 (95 % CI) 1.4 (0.87–2.2) mM 63 (47–84) µM 64 (49–83) µM

EC50_2 (95 % CI) – 5.5 (1.3–23) mM 4.0 (1.4–11) mM b GABA nH_1 ± SEM 0.54 ± 0.040 1.0 1.0

nH_2 – 1.0 1.0 Frac (95 % CI) – 0.74 (0.67–0.80) 0.71 (0.65–0.77) n 6 8 9 d Emax ± SEM 1.2 ± 0.10 0.086 ± 0.018 0.10 ± 0.020

EC50 (95 % CI) 45 (31–64) µM 62 (22–172) µM 63 (27–143) µM 2′MeO6MF nH ± SEM 1.8 ± 0.42 1.4 ± 0.88 1.6 ± 0.76 n 8 7 6 e Emax ± SEM 20 ± 1.8 5.5 ± 0.80 3.2 ± 0.40

Etomidate EC50 (95 % CI) 8.7 (6.7–11) µM 55 (27–110) µM 43 (25–76) µM

(-60 mV) nH ± SEM 2.0 ± 0.32 1.3 ± 0.37 1.5 ± 0.46 n 5 6 7 f Emax ± SEM 9.9 ± 0.84 2.8 ± 0.50

Etomidate EC50 (95 % CI) 19 (11–31) µM 60 (19–190) µM N.D.g (-30 mV) nH ± SEM 1.3 ± 0.33 1.2 ± 0.50 n 4 5 h Emax ± SEM 5.7 ± 0.55 0.64 ± 0.23

EC50 (95 % CI) 85 (33–220) µM 74 (33–170) µM Propofol N.D.g nH ± SEM 1.4 ± 0.50 2.0 ± 1.2 n 4 4 a The total amount of mRNA injected per oocyte was adjusted to be approximately 5 ng for each ratio. b Data of 1:15 ratio were better fitted to a monophasic model. Data of 1:50 and 1:100 ratios were significantly better fitter to a biphasic model (p < 0.0001 for both ratios; extra sum-of-squares F test). Concentration-response curves are shown in Figure 3.6F. c Current amplitude elicited by 30 mM GABA. d 300 µM 2′MeO6MF-elicited current normalised against 3 mM GABA response. e 30 µM (for 1:15) and 300 µM (for 1:50 and 1:100) etomidate-elicited current normalised against 3 mM GABA response. f 300 µM etomidate-elicited current normalised against 3 mM GABA response. g N.D. = not determined. h 300 µM propofol-elicited current normalised against 3 mM GABA response.

3.2.5. Differential activation of β3γ2L receptors with different subunit

stoichiometries by general anaesthetics

2′MeO6MF activity has previously been shown to be affected by a well-established mutation that perturbs etomidate activity [1], suggesting that both ligands could share a common activation mechanism. Like 2′MeO6MF, etomidate did not activate homomeric

2L (Figure 3.3B) or 22L receptors (n = 4; data not shown). However, etomidate activated β3 homomeric receptors efficaciously (Emax = 15 ± 0.79, normalised against 60 mM GABA), with an EC50 of 7.7 µM (95 % CI: 5.6–11 µM; n = 5; Figure 3.7E and F), in contrast to 2′MeO6MF inhibition of the receptors’ constitutive activity (Figure 3.3C).

We then evaluated etomidate at β3γ2L (1:15), (1:50) and (1:100) receptors. Etomidate directly activated β3γ2L receptors expressed at all three ratios, albeit with different relative efficacies (Figure 3.7A). At 10 µM, etomidate directly activated β3γ2L (1:15) approximately 12 ± 1.1 fold higher relative to 3 mM GABA (n = 7), significantly greater than at 1:50 (0.46 ± 0.10; n = 6) and 1:100 (0.27 ± 0.05; n = 7) ratios (p ≤ 0.001; Tukey’s test; Figure 3.7B). At β3γ2L (1:50) and (1:100) receptors, consistent with the almost identical GABA and 2′MeO6MF profiles, etomidate activated β3γ2L receptors at both ratios with no significant differences in potencies and maximal efficacies (p > 0.05; unpaired t test; Figure 3.7C and Table 3.2). At 1:15 ratio, we could not construct a complete concentration-response curve as the current elicited by etomidate at concentrations above 10 µM frequently exceeded the measuring range of the amplifier (>

10 µA).

Figure 3.7. Etomidate activates both β3γ2L and β3 GABAARs. (A) Representative traces of etomidate (10 µM) direct activation in comparison to 3 mM GABA at β3γ2L (1:15), (1:50) and (1:100) GABAARs. (B) Mean efficacy of 10 µM etomidate activation at β3γ2L (1:15; n = 7), (1:50; n = 6) and (1:100; n = 7) receptors. Data are normalised to the 3 mM GABA response. The mean efficacy of 10 µM etomidate at various ratios was compared using Tukey’s test, and the significance levels are indicated with n.s. (not significant) and *** (p ≤ 0.001). (C) Concentration-response curves of etomidate

93 activation at β3γ2L receptors expressed at 1:15 (n = 5), 1:50 (n = 6) and 1:100 (n = 7) ratios. Recording was conducted at -60 mV. Data are normalised to the 3 mM GABA response. (D) Concentration-response curves of etomidate activation at β3γ2L receptors expressed at 1:15 (n = 4) and 1:100 (n = 5) ratios. Recording was conducted at -30 mV. Data are normalised to the 3 mM GABA response. (E) Representative traces of β3 homomeric receptors responses to 0.1–300 µM etomidate in comparison to 60 mM GABA response. (F) Etomidate concentration-response curve for β3 homomeric receptors (n = 5). Data are normalised against 60 mM GABA responses. Data are presented as mean ± SEM. Bars indicate durations of drug application.

To investigate etomidate activity across the whole concentration range at β3γ2L (1:15) receptors, we conducted electrophysiological recording at -30 mV. The same experiment was repeated at β3γ2L (1:100) receptors as a control. As expected from the voltage- dependent profile of etomidate [10], we observed reduced etomidate activation at -30 mV.

However, etomidate still exhibited significantly larger relative efficacy at 1:15 (9.9 ±

0.84) than 1:100 (2.8 ± 0.50) ratio (p ≤ 0.01; unpaired t test; Figure 3.7D; Table 3.2).

Etomidate also showed a modest, but significantly higher potency at 1:15 (EC50 = 19 µM) than 1:100 (EC50 = 60 µM) ratio (p ≤ 0.01 for logEC50 comparison; unpaired t test; Table

3.2).

We also investigated the action of propofol, a non-selective general anaesthetic which has been shown to bind with similar affinity to at least 4 binding sites at GABAARs found on

β+α‒, α+β‒, β+β‒ and γ+β‒ subunit interfaces [11, 12]. We found that propofol directly activated β3γ2L receptors, in agreement with previous findings reported on β2γ2S receptors [13]. Like etomidate, propofol directly activated β3γ2L (1:15) and (1:100) receptors differentially (Figure 3.8A). At 300 µM, propofol elicited current responses approximately 6-fold larger than maximal GABA currents at β3γ2L (1:15) receptors, significantly larger than at β3γ2L (1:100) receptors (Emax = 0.64 ± 0.23; p ≤ 0.001; unpaired t test; Figure 3.8B; Table 3.2). However, the potencies of propofol measured at

1:15 (EC50 = 85 µM) and 1:100 (EC50 = 74 µM) ratios were not different (p > 0.05; unpaired t test; Table 3.2).

Figure 3.8. Propofol activates β3γ2L (1:15) and (1:100) receptors with different relative efficacies. (A) Representative traces of propofol (1–300 µM) direct activation in comparison to 3 mM GABA at β3γ2L (1:15) and (1:100) GABAARs. (B) Concentration- response curves of propofol activation at β3γ2L receptors expressed at 1:15 (n = 4) and 1:100 (n = 4) ratios. Data are normalised to the 3 mM GABA response.

3.3. Discussion

2′MeO6MF has been previously demonstrated to directly activate recombinant

α2β2/3γ2L GABAARs expressed in Xenopus oocytes [1], but the functional significance of each subunit remains unclear. In this study, we performed a systematic investigation of 2′MeO6MF activity at all receptor combinations expressed in Xenopus oocytes using

α2, β3 and γ2L subunits and found that 2′MeO6MF exhibits distinct pharmacology at β3,

β3γ2L and α2β3γ2L GABAARs. It acts as an inverse agonist at β3 homomers by inhibiting the constitutive activity of the receptors, and activates β3γ2L and α2β3γ2L GABAARs with different relative efficacies. The lack of activity of 2′MeO6MF at α2β3 (20:1) binary receptors, despite the presence of the β3 subunit, favours an interfacial 2′MeO6MF

95 binding site(s) containing the β3 subunit (β3-β3, β3-γ2L or γ2L-β3) over an intra-β3- subunit binding site.

The robust formation of the non-α-containing β3γ2L GABAARs raises a critical question of where GABA binds to activate this receptor as it lacks the β+α‒ interfaces. Despite the absence of any known GABA binding sites in β3γ2L receptors, GABA elicited robust responses within a similar concentration range that activates αβ and αβγ receptors. Thus, we infer that GABA binding site(s) which are previously unknown may exist. A GABA binding site at the β-β interface is possible, but the poor GABA activity at β3 homomeric receptors even at saturating concentrations suggests that other subunit interfaces, possibly between the β and γ subunits are more likely to contribute to the GABA sensitivity of

β3γ2L receptors.

There is a wealth of evidence supporting two β+α‒ interfacial binding sites for etomidate in the transmembrane domain [14-16]. Our data show that etomidate activates β3 homomeric and β3γ2L binary receptors efficaciously, but is insensitive at γ2L homomeric receptors. These findings indicate that the β, but not the α subunit is essential for etomidate sensitivity. This is supported by findings from a photoaffinity labelling study which suggested that etomidate can also bind to a homologous β+β‒ interfacial site with less selectivity [12]. Besides that, mutations found on the β subunit such as N265M [17] and M286W [18] have been shown to completely eliminate etomidate sensitivity, while mutations found on the α subunit tend to have less profound effects [19]. Taken together, we infer from these findings that the β subunit, which bears the principal (+) components of the etomidate binding sites, dominates ligand binding interactions, whereas the identity of the complementary (‒) subunit (α, β or γ) is well tolerated.

We also demonstrate that β3γ2L receptors expressed at different injection ratios exhibit distinct levels of constitutive activity, relative efficacies of 2′MeO6MF, etomidate and propofol activation as well as different GABA potencies. These findings strongly suggest that β3γ2L receptors with different subunit stoichiometries are being formed. The influence of subunit cDNA or mRNA ratios on receptor stoichiometry expressed in heterologous systems is well established in the case of α4β2 nicotinic acetylcholine receptors, in which α4-biased and β2-biased ratios are routinely used to selectively express the pharmacologically dissimilar 3(α4):2(β2) and 2(α4):3(β2) stoichiometries respectively [20, 21]. It is not clear from our experiments which stoichiometries of β3γ2L receptors exist. However, as the binary αβ GABAARs have been shown to assemble into two functional stoichiometries of 2α:3β and 3α:2β [22-24], by analogy, we propose that

β3γ2L GABAARs also exist in two different stoichiometries of 3β:2γ and 2β:3γ. It is likely that when the γ2L subunit mRNA was injected in greater abundance relative to the β3 subunit (e.g., 1:50 and 1:100 ratios), the 2β:3γ stoichiometry is predominantly expressed.

In contrast, when the injection ratio was reduced (e.g., 1:15), the 3β:2γ stoichiometry may be favourably expressed. The definitive mechanisms underlying the distinct pharmacological profiles of β3γ2L stoichiometric forms are beyond the scope of this investigation, but differences in the number of functional binding interfaces and intrinsic activation properties may have a role. Further experiments are required to understand how these factors result in the complex pharmacology observed for β3γ2L receptors.

We and others have shown that 2′MeO6MF, etomidate, pentobarbital and dopamine appear to be more efficacious agonists than GABA at β3γ2 receptors [25, 26]. In contrast, these ligands directly activate αβγ receptors with lower efficacies relative to the maximal

GABA responses [18, 26, 27]. Similar observations have been reported at αβδ receptors,

97 where GABA acts as a partial agonist with low intrinsic efficacy, and modulators such as neurosteroids [28], etomidate [29], propofol [30] and pentobarbital [27] are able to enhance maximal GABA currents significantly larger than they do at αβγ receptors, where

GABA acts as a full agonist. It is possible that GABA also acts as a weak partial agonist at βγ receptors. However, this inference is not conclusive as the intrinsic efficacies of

GABA and other ligands at βγ receptors are not well established and were not systematically explored in our study. Further studies are required to address these issues, and to clarify whether the apparent larger relative efficacies of various ligands at βγ receptors are due to subtype-dependent functional differences or the low GABA efficacy at these channels.

The expression of βγ receptors has been reported previously [13, 25, 26, 31-34]. In these studies, the overall pharmacological profiles of βγ receptors are very similar to that of the

αβγ receptors. This is concerning as pharmacological tools such as diazepam, which is routinely used as an indicator of the purity of the αβγ receptors heterologously expressed would not be able to distinguish βγ from αβγ receptors due to its comparable actions at both receptor subtypes [31, 33]. The GABAAR competitive antagonist bicuculline has recently been suggested to have β3γ2-selective pharmacology as it exhibits inhibitory action at α1β3γ2 but activates β3γ2 receptors [26]. However, as bicuculline has been shown to also activate β3 homomers [8], it is not known if the effect observed at β3γ2 receptors was mediated by a background population of β3 homomers. The great diversity in GABAAR subtypes dictates the need for subtype-specific ligands to understand the function of these receptors. As such, the search for βγ-selective ligands is pivotal to accurately determine the pharmacology of these receptors in vitro.

There is emerging evidence to suggest that native GABAAR subtypes are far more diverse than earlier studies have indicated, as exemplified by the in vivo expression of αβ receptors which were previously thought to only form in heterologous systems [35-39].

In single channel recording experiments conducted in dissociated rat hippocampal neurons, it has been shown that the activity of 2′MeO6MF, like diazepam, is significantly attenuated by the γ2(381-403) MA peptide which disrupts protein-receptor interactions involving only the γ2 subunit [1, 2]. This finding suggests that γ2-containing GABAARs are the molecular target of 2′MeO6MF, and the subunit composition of these receptors is most likely to make up of α, β and γ subunits since they are the most common native isoforms. It is currently not known if βγ receptors exist in neurons and thus contribute partially to the flavonoid activation observed. Again, βγ-selective ligands will be necessary to explore this possibility.

Overall, our findings have important implications in the study of GABAARs. We have shown that β3γ2L receptors assemble with different subunit stoichiometries in Xenopus oocytes, which can be differentiated by GABA, 2’MeO6MF, etomidate and propofol.

These receptors lack the α subunit which contributes the complementary (‒) components of the GABA and etomidate binding sites, but are still activated by GABA and etomidate, suggesting that the structural requirements for these sites need to be redefined. While the in vivo existence of βγ receptors remains elusive, it can serve as a tool to study the contribution of different subunits or subunit interfaces in the pharmacology of a chemical of interest.

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23. Baumann SW, Baur R, Sigel E. Subunit 33. Whittemore ER, Yang W, Drewe JA, arrangement of γ-aminobutyric acid type A Woodward RM. Pharmacology of the receptors. Journal of Biological Chemistry. human gamma-aminobutyric acidA 2001;276(39):36275-80. receptor α4 subunit expressed in Xenopus laevis oocytes. Molecular Pharmacology. 24. Boileau AJ, Pearce RA, Czajkowski C. 1996;50(5):1364-75. Tandem subunits effectively constrain GABAA receptor stoichiometry and 34. Miko A, Werby E, Sun H, Healey J, Zhang recapitulate receptor kinetics but are L. A TM2 residue in the β1 subunit insensitive to GABAA receptor-associated determines spontaneous opening of protein. The Journal of Neuroscience. homomeric and heteromeric γ- 2005;25(49):11219-30. aminobutyric acid-gated ion channels. Journal of Biological Chemistry. 25. Taylor PM, Thomas P, Gorrie GH, 2004;279(22):22833-40. Connolly CN, Smart TG, Moss SJ. Identification of amino acid residues 35. Günther U, Benson J, Benke D, Fritschy within GABAA receptor β subunits that JM, Reyes G, Knoflach F, et al. mediate both homomeric and heteromeric Benzodiazepine-insensitive mice generated receptor expression. The Journal of by targeted disruption of the γ2 subunit Neuroscience. 1999;19(15):6360-71. gene of γ-aminobutyric acid type A

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Chapter 4

The modulation of recombinant GABAARs by kavain, the major constituent of the anxiolytic kava extract is attenuated by the β3N265M point mutation ______

4.1. Introduction

The use of intoxicating substances to enhance mood, alter consciousness and to achieve spiritual enlightenment is known in virtually every culture. Alcohol is the most commonly used intoxicant globally, with the exception in a few regions [1]. For millennia, Pacific

Islanders have been using the root of a native pepper plant called kava (Piper methysticum) to prepare a non-alcoholic psychoactive beverage, which is also called kava. Kava drinking is an integral component of the Pacific Islander culture, with kava playing roles as a sacred drug in religious rituals, a social lubricant at formal gatherings, and a medicine to induce relaxation and sleep [2]. The contemporary use of kava extends beyond ritualised circumstances [3, 4]. In some western societies, kava is used as a prescription-free alternative to the benzodiazepines to relieve stress-induced anxiety and insomnia [5]. The recreational use of kava as a substitute for alcohol is also gaining popularity owing to kava’s calming effect which contrasts the aggressive tendencies associated with alcohol [6].

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There is a long-standing interest in the use of kava in the treatment of anxiety. In clinical trials, kava extracts are superior to placebo in reducing anxiety, and are generally well tolerated with negligible to mild side effects [7-9]. Despite reports of alleged kava- induced hepatotoxicity which led to the withdrawal and restriction of kava in several countries [10, 11], systematic reviews and meta-analyses conducted over the last 15 years found a clear positive benefit-to-risk ratio for kava [12, 13]. In view of the lack of direct causal evidence for liver injury, the ban of kava in Germany was recently overturned, leading to a resurgence of attention on kava [14]. Currently, there is an ongoing phase III clinical trial aimed to establish the efficacy and safety of kava in patients diagnosed with generalised anxiety disorder [15].

A group of structurally-related, lipophilic compounds known as kavalactones (or kavapyrones) is responsible for the clinical effects of kava [16]. Kavain, along with dihydrokavain, methysticin, dihydromethysticin, yangonin, and desmethoxyangonin are the most abundant kavalactones (Figure 4.1) [17]. Numerous proteins including

+ 2+ GABAARs, voltage-gated Na and Ca channels, opioid µ and δ receptors, dopamine type-2 (D2) receptor, histamine type-1 and 2 (H1/2) receptors, cannabinoid type-1 (CB1) receptor, and monoamine oxidase type B (MAO-B) have been suggested to be the molecular targets for kavalactones [16, 18-20]. However, due to the paucity in robust evidence, a consensus on the pharmacology of kavalactones could not be reached.

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Figure 4.1. Chemical structures of the six major kavalactones found in kava.

The prevailing view that GABAARs are a strong candidate target for kavalactones is on the basis of their benzodiazepine-like pharmacological actions. To date, evidence that indicates kavalactone-GABAAR interaction mainly comes from radioligand binding studies. In rodent brain membranes, kava extracts with enriched kavalactone content were able to enhance the binding of radiolabelled orthosteric ligands to native GABAARs [19,

21]. The interaction of pure kavalactones with GABAARs has also been investigated, but mixed results were obtained. Davies et al. (1992) demonstrated that kavalactones only enhanced [3H]GABA binding marginally even at high concentrations (up to 1 mM) [22].

The removal of lipid membrane with detergent resulted in the complete loss of modulation, a finding that prompted the authors to speculate that kavalactones, due to their lipophilicity, alter the physicochemical properties of lipid membrane to cause conformational changes in GABAARs [22]. However, another study found that pure kavalactones showed low-micromolar affinity and structure-dependent activity at

GABAARs, implying that kavalactones do directly bind to GABAARs [23]. Notably, none of these studies detected significant affinity of kavalactones for the benzodiazepine binding site, contrary to popular belief.

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There is limited information on kavalactone mechanism of action provided by these studies due to two factors. First, the contribution of non-kavalactone compounds in enhancing ligand binding cannot be ruled out with the use of kava extracts in some studies. Second, kavalactones were tested on a heterogeneous population of GABAARs isolated from rodent brain, thus the identity of specific GABAAR subtypes modulated by kavalactones is not known. To address these limitations, in this study, we set out to perform functional characterisation of a kavalactone at 9 different GABAAR subtypes expressed in Xenopus oocytes. Kavain is selected as the representative kavalactone due to its abundance in kava extracts commonly consumed [24], as well as its established anxiolytic and sedative properties in animal and human [25-27].

4.2. Materials and Methods

4.2.1. Primer sequences used in site-directed mutagenesis

The protocol of site-directed mutagenesis is detailed in section 2.4.1.8. Primers used for the generation of desired mutations in different GABAAR subunits are shown in Table

4.1, with the nucleotides coding for the mutations underlined:

Table 4.1. Primers used to introduce the α1M236W, β2M286W, and β3N265M point mutations. Mutations Primers Forward: CCTGCCGTGCATATGGACAGTTATTCTC α1M236W Reverse: GAGAATAACTGTCCATATGCACGGCAGG Forward: AGGCCATTGACATGTACCTGTGGGGGTGCTTTGTCTTCGT β2M286W Reverse: ACGAAGACAAAGCACCCCCACAGGTACATGTCAATGGCCT Forward: ACAATGACAACCATCATGACCCACCTTCGGGAG β3N265M Reverse: CTCCCGAAGGTGGGTCATGATGGTTGTCATTGT

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4.2.2. Electrophysiological studies

The subunit mRNA ratios, and the total amount of mRNA injected into individual oocyte used for the expression of various GABAAR subtypes in this study are listed in Table 4.2.

The screening of kavain at α2/3β2γ2L GABAARs was performed by Kirsten Høstgaard-

Jensen from the Department of Drug Design and Pharmacology, University of

Copenhagen, Denmark.

Table 4.2. Subunit mRNA ratios used to express different GABAAR subtypes.

Subunit combination Ratio mRNA amount (ng/oocyte) α1β2 1:1 10 β2γ2L 1:3 10 α1β2γ2L 1:1:3 2.5 α5β2γ2L 1:1:3 1.5 α1β1γ2L 5:1:5a 5 α1β3γ2L 10:1:10a 1.5 α4β2δb 5:1:5 6 α1M236Wβ2γ2L 1:1:3 2.5 α1β2M286Wγ2L 1:1:3 2.5 α1β3N265Mγ2L 10:1:10a 1.5 a The α1 and γ2L subunit mRNAs are in excess to reduce the expression of homomeric β1/3 receptors. b The mRNA mixture was a generous gift from Leonny Hartidi who characterised the pharmacological profiles of α4β2δ receptors expressed at different injection ratios. The 5:1:5 ratio was found to give rise to a relatively homogeneous receptor population in Xenopus oocytes with consistent pharmacology [28].

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4.3. Results

4.3.1. Effect of GABAAR subunit composition on kavain activity

We tested kavain (10–300 µM) at α1β2γ2L GABAARs, the most abundant native

GABAAR isoform, in the presence of GABA EC3 (10 µM), and found that kavain enhances GABA-elicited responses in a concentration-dependent manner (Figure 4.2A).

The modulatory effect of kavain is moderate, with only 170 ± 23 % of enhancement measured at 300 µM (n = 6; Figure 4.2C). We also investigated the ability of kavain to activate the receptors, and found negligible agonist activity, with 300 µM kavain eliciting currents < 1 % of the 10 mM GABA responses (n = 5; Figure 4.2B). To understand the subunit requirement for kavain modulation, we tested kavain (10–300 µM) at different

GABAAR subtypes with a low concentration of GABA (EC3-7 for all receptors except

α4β2δ, where the EC30 was used due to the low level of expression). The receptor subtypes investigated were α1β2, β2γ2L, αxβ2γ2L (x = 1, 2, 3 or 5), α1βxγ2L (x = 1, 2 or 3) and α4β2δ (GABA concentration-response relationships summarised in Figure 4.3 and Table 4.3). Kavain potentiated current responses elicited by low concentrations of

GABA at all receptors to a similar degree, exhibiting no apparent subtype selectivity (no significant difference was detected for all pairwise comparisons using Tukey’s test; p >

0.5; Figure 4.2C).

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Figure 4.2. Kavain potentiates GABAARs with no apparent subtype selectivity. (A) Representative traces demonstrating kavain (10–300 µM) enhancing current elicited by 10 µM GABA (EC3) at α1β2γ2L GABAARs in a concentration-dependent manner. (B) Representative traces of current responses elicited by a maximal concentration of GABA (10 mM), in comparison to 300 µM kavain alone. (C) Top panel, Potentiation of GABA EC3-7 by 300 µM kavain at α1β2, β2γ2L, αxβ2γ2L (x = 1, 2, 3 and 5) and α1βxγ2L (x = 1, 2 and 3) GABAARs. At α4β2δ GABAARs, the GABA control (1 µM) corresponds to an EC30. Data are presented as mean ± SEM. Numbers in bars indicate the number of experiments. No significant difference was found for kavain potentiation at these receptor subtypes (Tukey’s test; p > 0.05 for all pairwise comparisons). Bottom panel, Superimposed current traces of GABA alone (black) and GABA in combination with 300 µM kavain (red) at the corresponding receptor subtypes. The black bars above the traces indicate duration of drug application.

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Figure 4.3. GABA concentration-response curves for all receptor subtypes expressed in this study. Data are presented as mean ± SEM. The curve parameters are listed in Table 4.3.

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Table 4.3. GABA concentration-response curve parameters at various receptor subtypes derived from curve-fitting procedures.

Receptor subtypes GABA EC50 (95% CI) nH ± SEM n α1β2 7.8 (6.9–8.8) µM 1.2 ± 0.072 5 β2γ2L 1.3 (0.9–1.8) mM 0.63 ± 0.034 4 α1β2γ2L 136 (116–158) µM 0.99 ± 0.062 10 α1β2γ2L (+ 300 µM kavain) 99 (87–113) µMa 0.75 ± 0.026 6 α2β2γ2L 76 (72–80) µM 1.4 ± 0.041 4 α3β2γ2L 150 (136–162) µM 1.2 ± 0.054 4 α5β2γ2L 67 (56–80) µM 1.0 ± 0.078 5 α1β1γ2L 89 (79–100) µM 1.1 ± 0.059 6 α1β3γ2L 93 (83–104) µM 1.2 ± 0.070 5 α4β2δ 2.6 (1.7–4.1) µM 0.70 ± 0.095 4 α4β2δ (+ 300 µM kavain) 1.7 (0.93–3.1) µMa 0.59 ± 0.11 4 α1M286Wβ2γ2L 59 (48–73) µM**b 0.80 ± 0.053 7 α1β2M286Wγ2L 25 (23–27) µM****b 1.1 ± 0.036 5 α1β3N265Mγ2L 72 (65–80) µMc 1.4 ± 0.083 5 a The GABA potency was not significantly altered in the presence of 300 µM kavain (p > 0.05 for logEC50 comparisons; paired t test). b The GABA potency was significantly enhanced compared to wild-type α1β2γ2L GABAARs (** p < 0.01; **** p < 0.0001 for logEC50 comparisons; unpaired t test). c The GABA potency was not significantly different from the wild-type α1β3γ2L GABAARs (p > 0.05 for logEC50 comparisons; unpaired t test).

4.3.2. Kavain’s differential effects at α1β2γ2L and α4β2δ GABAARs

To understand the nature of kavain modulation in response to different GABA concentrations, we constructed GABA concentration-response curves with and without

300 µM kavain at α1β2γ2L and α4β2δ receptors. These receptors represent synaptic and extrasynaptic GABAARs respectively, which have distinct subcellular localisations

(section 1.9) and GABA sensitivities (α1β2γ2L: EC50 = 136 µM; α4β2δ: EC50 = 2.6 µM;

Figure 4.3; Table 4.3). At α1β2γ2L receptors, kavain showed modest enhancement at

GABA concentrations below EC45, but did not affect the EC50 and maximal efficacy of

GABA (p > 0.05 for both parameters; paired t test; Figure 4.4A and D; Table 4.3). In

111 contrast, kavain increased the maximal GABA efficacy by two fold (p < 0.0001; paired t test) at α4β2δ receptors without affecting the GABA potency (p > 0.05 for logEC50 comparison; paired t test; Figure 4.4B and D; Table 4.3).

Figure 4.4. Kavain produced greater enhancement of GABA-elicited currents at α4β2δ than α1β2γ2L GABAARs. (A) GABA concentration-response curves in the absence (black; n = 10) and presence (red; n = 6) of 300 µM kavain at α1β2γ2L GABAARs. The curve parameters are summarised in Table 4.3. (B) GABA concentration-response curves in the absence (black; n = 4) and presence (red; n = 4) of 300 µM kavain at α4β2δ GABAARs. The curve parameters are summarised in Table 4.3. (C) Superimposed current responses of maximal GABA in the absence (black) and presence (red) of 300 µM kavain at α1β2γ2L and α4β2δ GABAARs. (D) The effect of kavain on the maximal GABA current responses at α1β2γ2L (n = 6) and α4β2δ (n = 7) GABAARs was compared using the paired t test, and the significance levels are indicated with n.s. (not significant) and **** (p < 0.0001). Data are presented as mean ± SEM.

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4.3.3. Effect of membrane potential on kavain modulation

The activity of some GABAAR modulators is known to be voltage dependent. For instance, anaesthetic agents such as etomidate, propofol and isoflurane exhibit greater potentiation at negative membrane potentials, and the effect decreases as the membrane potentials become more positive [29]. To determine if kavain also shows voltage dependence, the enhancement of 10 µM GABA (EC3) responses mediated by 300 µM kavain across a range of membrane potentials (-80 to -30 mV, applied as 10 mV steps) was measured at α1β2γ2L GABAARs. We found that changes in membrane potential had no significant effect on kavain modulation (p > 0.05 for all pairwise comparisons;

Tukey’s test; n = 6–9 for each data point; Figure 4.5).

Figure 4.5. Membrane potential has negligible effect on the degree of potentiation of kavain. (A) Representative traces demonstrating the current responses to 10 µM GABA (EC3) alone and the co-application of 10 µM GABA and 300 µM kavain when the membrane potential at which the oocyte was voltage-clamped was changed from -80 to - 30 mV in 10 mV steps at α1β2γ2L GABAARs. (B) Potentiation of GABA EC3 by 300 µM kavain at α1β2γ2L GABAARs plotted against the holding potential. No significant difference was found for kavain potentiation at different membrane potential (Tukey’s test; p > 0.05 for all pairwise comparisons). Data are presented as mean ± SEM. n = 6–9 for each data point.

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4.3.4. Interaction of kavain with various allosteric binding sites

The psychoactive properties of kavalactones have sparked concern of potential adverse pharmacodynamic interactions with the co-administration of kava with other CNS depressants, but there is a lack of experimental evidence to support these postulations

[30]. Thus, we sought to determine whether kavain interacts with other GABAAR allosteric modulators in an additive, synergistic or antagonistic manner.

4.3.4.1. Benzodiazepines

We investigated the effect of combining benzodiazepine site ligands with kavain at

α1β2γ2L GABAARs (n = 5; Figure 4.6). In concordance with the literature, diazepam (1

µM) enhanced current responses elicited by 10 µM GABA in a flumazenil-sensitive manner, and flumazenil alone did not alter GABA responses, consistent with its neutralising modulator profile [31]. Unlike diazepam, the GABA-enhancing action of 300

µM kavain was unaffected in the presence of flumazenil. We also found that the co- application of kavain and diazepam resulted in significantly greater enhancement of

GABA current (350 ± 10 %) compared to their actions alone (GABA + diazepam: 260 ±

4.2 %, p < 0.001; GABA + kavain: 260 ± 11 %, p < 0.0001; paired t test; Figure 4.6B).

However, this degree of potentiation was less than additive (theoretical additive value =

420 %). The same experiment was repeated with 100 nM diazepam, and a less-than- additive interaction was detected again with the kava and diazepam combination (n = 3; data not shown).

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Figure 4.6. Flumazenil-insensitive kavain potentiation has less-than-additive effect on diazepam action. Top, Representative traces demonstrating responses to control (10 µM GABA); control and 1 µM diazepam; control, 1 µM diazepam and 10 µM flumazenil; control and 10 µM flumazenil; control and 300 µM kavain; control, 300 µM kavain and 10 µM flumazenil; and control, 300 µM kavain and 1 µM diazepam. Bottom, The modulatory effect of diazepam, flumazenil and kavain at α1β2γ2L GABAARs (n = 5). Kavain potentiation was unchanged in the presence of flumazenil (G + K vs. G + K + F; p > 0.05; paired t test). The combination of kavain and diazepam (G + K + D) resulted in greater potentiation than diazepam (G + D; p < 0.001; paired t test) and kavain (G + K; p < 0.0001; paired t test) alone, but the effect was less than the theoretical additive modulatory effect (dotted line). Data are normalised to the current responses elicited by 10 µM GABA, and are presented as mean ± SEM.

4.3.4.2. Allopregnanolone

Next, we investigated the interaction of kavain with the neurosteroid allopregnanolone at

α1β2γ2L GABAARs (n = 5; Figure 4.7). At 1 µM, allopregnanolone enhanced GABA

EC3 responses efficaciously to 730 ± 62 %. Co-application of allopregnanolone with 300

µM kavain did not result in a significant increase in enhancement (840 ± 60 %; p > 0.05; paired t test; Figure 4.7B). As we failed to detect agonist activity with allopregnanolone

115 even at 30 µM, in agreement with Hammer et al. [32], the presence of the neurosteroid activation site previously reported [33] could not be ascertained under our experimental conditions. Thus, the interaction of kavain with the neurosteroid activation site was not explored.

Figure 4.7. The lack of effect of kavain on allopregnanolone potentiation at α1β2γ2L GABAARs. Top, Representative traces demonstrating the current responses elicited by 10 µM GABA (control); control with 300 µM kavain; control with 1 µM allopregnanolone; and control, 300 µM kavain and 1 µM allopregnanolone. Bottom, The modulatory effect of kavain and allopregnanolone (n = 5). The degree of potentiation elicited by allopregnanolone alone is not significantly different from that of the co- application of allopregnanolone and kavain (G + A vs. G + K + A; p > 0.05; paired t test; n.s. = not significant). Data are normalised to the 10 µM GABA responses, and are presented as mean ± SEM.

4.3.4.3. DS2

We also investigated the modulatory effect of kavain and DS2, either alone or in combination on the maximal GABA responses (1 mM) at α4β2δ GABAARs (n = 7;

Figure 4.8). 300 µM kavain and 1 µM DS2 enhanced the GABA responses by 200 ± 11

% and 380 ± 32 % respectively (Figure 4.8B). Co-application of 300 µM kavain and 1

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µM DS2 resulted in 400 ± 29 % potentiation, which was not significantly different from the degree of potentiation elicited from 1 µM DS2 alone (p > 0.05; paired t test; Figure

4.8B).

Figure 4.8. The lack of effect of kavain on DS2 potentiation at α4β2δ GABAARs. Top, Representative traces demonstrating the current responses elicited by 1 mM GABA (control); control with 300 µM kavain; control with 1 µM DS2; and control, 300 µM kavain and 1 µM DS2. Bottom, The modulatory effect of kavain and DS2 alone and in combination (n = 7). The degree of potentiation elicited by DS2 alone is not significantly different from that of the co-application of kavain and DS2 (G + DS2 vs. G + K + DS2; p > 0.05; paired t test; n.s. = not significant). Data are normalised to the 1 mM GABA responses, and are presented as mean ± SEM.

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4.3.4.4. General anaesthetics

The effect of combining kavain and general anaesthetics (etomidate and propofol) on receptor modulation was also investigated at α1β2γ2L GABAARs. Both a low and a high concentration of the general anaesthetics were used (3 and 30 µM for etomidate; 10 and

100 µM for propofol) to produce the modulatory and agonist effects respectively.

Etomidate

At 3 µM, etomidate enhanced the GABA EC3 responses efficaciously (53 ± 2.6 % of maximal GABA responses; n =7; Figure 4.9). The addition of 300 µM kavain resulted in a modest, but significant reduction in potentiation (45 ± 3.1 % of maximal GABA responses; n = 7; p < 0.01; paired t test). At 30 µM, etomidate directly elicited current responses approximately 24 ± 4.2 % of the maximal GABA amplitude (n = 6). The agonist effect was not significantly altered in the presence of 300 µM kavain (27 ± 2.8 %; n = 6; p > 0.05; paired t test).

Propofol

At 10 µM, propofol enhanced the GABA EC3 responses efficaciously (45 ± 3.5 % of the maximal GABA responses; n =5; Figure 4.10). The addition of 300 µM kavain resulted in a slight, but insignificant enhancement in propofol potentiation (50 ± 4.8 %; p > 0.05; paired t test). Following exposure to 100 µM propofol, GABA-elicited current responses were highly variable, resulting in inconsistent measurement. Thus, the agonist effect of propofol was investigated in the absence of GABA. We found that when combined, the currents elicited by 300 µM kavain and 100 µM propofol were smaller than the

118 application of propofol alone (91 ± 1.5 % of the maximal propofol responses; n = 5; p <

0.01; paired t test; Figure 4.10).

Figure 4.9. Kavain modestly reduced etomidate potentiation, but did not affect the direct activation caused by etomidate at α1β2γ2L GABAARs. Top, Representative traces of current responses elicited by 10 mM GABA (control); 10 µM GABA; 10 µM GABA and 300 µM kavain; 10 µM GABA and 3 µM etomidate; 10 µM GABA, 300 µM kavain and 3 µM etomidate; 30 µM etomidate; 30 µM etomidate and 300 µM kavain. Bottom, Kavain caused a subtle but significant reduction in etomidate potentiation (G2 + E1 vs. G2 + K + E1; n = 7; p < 0.01; paired t test), but had no effect on etomidate activation (E2 vs. E2 + K; n = 6; p > 0.05; paired t test). Data are normalised to the 10 mM GABA responses, and are presented as mean ± SEM.

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Figure 4.10. Kavain did not affect propofol potentiation, but modestly reduced propofol activation at α1β2γ2L GABAARs. Top, Representative traces of current responses to 10 mM GABA (control); 10 µM GABA; 10 µM GABA and 300 µM kavain; 10 µM GABA and 10 µM propofol; 10 µM GABA, 300 µM kavain and 10 µM propofol. Middle, Continuous traces demonstrating two consecutive applications of control (100 µM propofol) followed by the co-application of 300 µM kavain with control; and control. Bottom, Receptor modulation produced by propofol alone (G2 + P) was not significantly different from the combination of kavain and propofol (G2 + K + P; n = 5; p > 0.05; paired t test). The agonist effect of propofol (P2) was significantly reduced in the presence of kavain (P2 + K; n = 5; p < 0.01; paired t test). Data are presented as mean ± SEM.

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4.3.5. Kavain activity at receptors containing α1M236W, β2M286W and β3N265M mutations

One of the possible reasons for the inhibitory effect of kavain on the activity of anaesthetics is that kavain competes with these ligands for the same sites on GABAARs.

However, given the lack of potency and efficacy of kavain, whether the nature of this inhibition is competitive could not be established. We hypothesised that if kavain binds to the same sites as etomidate and propofol, mutations in the β+α‒ transmembrane anaesthetic sites (α1M236W, β2M286W and β3N265M; Figure 4.11) which perturb etomidate and propofol actions would also affect kavain activity.

Figure 4.11. The putative β+α‒ anaesthetic binding sites in a model structure of the α1β2γ2 GABAAR. The subunits are colour coded: α1, orange; β2, turquoise; γ2: green. (A) Side view of the receptor structure, with a close-up view of the three residues which are implicated in anaesthetic binding (inset): α1M236 (dark green), β2/3N265 (red) and β2/3M286 (dark blue). (B) Transmembrane region of the model structure with the extracellular domains removed for clarity, illustrating the positions of α1M236, β2/3N265 and β2/3M286 residues. The arrangement of the TMDs (M1–M4) of an α1 and a β2 subunit is illustrated.

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4.3.5.1. α1M236W

The co-expression of α1M236W with β2 and γ2L subunits resulted in receptors with higher GABA potency (EC50 = 59 µM; n = 7) than wild-type α1β2γ2L GABAARs (EC50

= 136 µM; p < 0.01 for logEC50 comparisons; unpaired t test; Figure 4.3; Table 4.3). The point mutation significantly attenuated the modulatory effects of 3 µM etomidate (p <

0.0001; n = 5; unpaired t test; Figure 4.12) and 10 µM propofol (p < 0.01; n = 5; unpaired t test; Figure 4.12) on GABA EC3 (1 µM) responses. In contrast, the agonist effects of

30 µM etomidate and 100 µM propofol appeared to be greater, but were not significantly different from wild-type receptors (p > 0.05; n = 5 for both etomidate and propofol; unpaired t test; Figure 4.12). Overall, these functional characteristics are consistent with previous studies [34].

Next, we investigated the effect of kavain alone and in combination with GABA EC3 at

α1M236Wβ2γ2L GABAARs. At 300 µM, kavain elicited current responses approximately 63 ± 5.2 % of the GABA EC3 responses (n = 6), which was significantly greater than that observed at wild-type receptors (15 ± 3.6 %; n = 4; p < 0.001; unpaired t test; Figure 4.14). The degree of potentiation produced by the co-application of GABA

EC3 and 300 µM kavain, however, was not significantly different between α1β2γ2L (250

± 7.8 % of GABA EC3 responses; n = 4) and α1M236Wβ2γ2L (220 ± 11 %; n = 6)

GABAARs (p > 0.05; unpaired t test).

4.3.5.2. β2M286W

Receptors expressed from α1, β2M286W and γ2L subunits displayed enhanced GABA sensitivity (EC50 = 25 µM; n = 5; p < 0.0001 for logEC50 comparisons vs. α1β2γ2L; unpaired t test; Figure 4.3; Table 4.3). In agreement with previous studies, etomidate

122 sensitivity was abolished with the introduction of this point mutation, which is evident from the absence of modulatory and agonist effects (Figure 4.12) [34]. Receptor modulation produced by 10 µM propofol in the presence of GABA EC3 (1 µM) was also significantly diminished (p < 0.001; unpaired t test; Figure 4.12). However, the direct activation caused by 100 µM propofol was unaffected (p > 0.05; unpaired t test; Figure

4.12). Similar propofol profile at α1β2M286Wγ2S receptors has been reported [35]. Like

α1M236Wβ2γ2L, α1β2M286Wγ2L receptors showed increased efficacy in response to

300 µM kavain alone (96 ± 5.8 % of the GABA EC3 responses; p < 0.0001; unpaired t test; Figure 4.14). The modulatory effect of kavain on GABA EC3-elicited current responses, however, was not significantly different from α1β2γ2L GABAARs (270 ± 8.7

% of GABA EC3; p > 0.05; unpaired t test; Figure 4.14).

4.3.5.3. β3N265M

The asparagine-to-methionine mutation at position 265 of the β2/3 subunits is known to profoundly weaken anaesthetic sensitivity with minimal effect on GABA activity (section

1.11.3.5) [36, 37]. We found that GABA potencies at both α1β3γ2L and α1β3N265Mγ2L

GABAARs were not significantly different (p > 0.05 for logEC50 comparisons; unpaired t test; Figure 4.3; Table 4.3). Etomidate did not modulate (at 3 µM) nor activate (at 30

µM) the point-mutated receptors (Figure 4.13). Pronounced attenuation of propofol activity was also observed (Figure 4.13). In contrast to the α1M236W and β2M286W mutations, 300 µM kavain had negligible action by itself at α1β3N265Mγ2L receptors

(3.3 ± 1.0 % of GABA EC3 responses). However, kavain modulation of GABA EC3 responses was significantly reduced from 260 ± 18 % at α1β3γ2L to 110 ± 14 % at

α1β3N265Mγ2L receptors (p < 0.001; unpaired t test; Figure 4.14).

123

Figure 4.12. Effect of α1M236W and β2M286W point mutations on etomidate and propofol sensitivity. (A) Representative traces demonstrating the modulatory effect of 3 µM etomidate and 10 µM propofol on GABA EC3 (left) and the agonist effect of 30 µM etomidate and 100 µM propofol relative to 10 mM GABA (right) at α1β2γ2L, α1M236Wβ2γ2L, and α1β2M286Wγ2L GABAARs. (B) Etomidate and propofol modulation, but not activation were significantly reduced at α1M236Wβ2γ2L GABAARs. Both modulatory and agonist actions of etomidate were significantly diminished at α1β2M286Wγ2L GABAARs. Propofol modulation was also significantly weakened, while its agonist effect was unaffected. Data are normalised to current responses elicited by 10 mM GABA, and are presented as mean ± SEM. ** p < 0.01; *** p < 0.001; **** p < 0.0001; unpaired t test (mutants vs. wild-type). Numbers above bars indicate number of experiments. GABA + ETO: GABA EC3 + 3 µM etomidate; ETO: 30 µM etomidate; GABA + PRO: GABA EC3 + 10 µM propofol; PRO: 100 µM propofol.

124

Figure 4.13. The pronounced effect of β3N265M point mutation on etomidate and propofol sensitivity. (A) Representative traces demonstrating the modulatory effect of 3 µM etomidate and 10 µM propofol on GABA EC3 (left) and the agonist effect of 30 µM etomidate and 100 µM propofol relative to 10 mM GABA (right) at α1β3γ2L and α1β3N265Mγ2L GABAARs. (B) The modulatory and agonist effects of etomidate and propofol were markedly diminished at α1β3N265Mγ2L GABAARs. Data are normalised to current responses elicited by 10 mM GABA, and are presented as mean ± SEM. *** p < 0.001; **** p < 0.0001; unpaired t test (mutant vs. wild-type). Numbers above bars indicate number of experiments. GABA + ETO: GABA EC3 + 3 µM etomidate; ETO: 30 µM etomidate; GABA + PRO: GABA EC3 + 10 µM propofol; PRO: 100 µM propofol.

125

Figure 4.14. Effect of α1M236W, β2M286W and β3N265M point mutations on kavain activity. (A) Representative traces demonstrating current responses elicited by GABA EC3 (black bar), 300 µM kavain (red bar), and the combination of GABA EC3 and 300 µM kavain at α1β2γ2L, α1M236Wβ2γ2L, α1β2M286Wγ2L, α1β3γ2L, and α1β3N265Mγ2L GABAARs. (B) Top, Kavain displayed increased agonist efficacy at α1M236Wβ2γ2L and α1β2M286Wγ2L GABAARs, but the combination of GABA EC3 and kavain produced similar extent of potentiation compared to α1β2γ2L GABAARs. Bottom, Kavain modulatory action was significantly reduced at α1β3N265Mγ2L GABAARs. Data are normalised to GABA EC3 responses, and are presented as mean ± SEM. Numbers above bars indicate the number of experiments. Dotted lines indicate the current responses elicited by GABA EC3 alone. *** p < 0.001; **** p < 0.0001; unpaired t test (mutant vs. wild-type). Note: kavain-elicited current amplitudes were not deducted from the measurement of kavain-induced GABA potentiation.

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4.4. Discussion

In this study, we functionally characterised the effects of kavain at 9 different human

GABAAR subtypes expressed in Xenopus oocytes, and found no subtype dependence in kavain potentiation (Figure 4.2C). The concentration-response relationships of kavain at all receptor subtypes investigated were steep, with no plateau obtained even at its solubility limit. Thus, we could not accurately determine the potency of kavain. The inference on the lack of subtype selectivity was based on the modulation efficacy of currents elicited by a low concentration of GABA by a maximal concentration of kavain

(300 µM). Our results demonstrated that kavain potentiation did not require the α1 or γ2L subunit, as both β2γ2L and α1β2 GABAARs were modulated to similar extent as α1β2γ2L

GABAARs. Furthermore, varying the α (α1, 2, 3 or 5) and β (β1, 2 or 3) subunit isoforms in αβγ2L receptors did not result in significant differences in kavain potentiation.

However, kavain displayed functional differences at α1β2γ2L and α4β2δ GABAARs

(Figure 4.4). Kavain modulated α1β2γ2L GABAARs at GABA concentrations below the

EC45, but failed to potentiate peak GABA currents at higher concentrations. In contrast, the GABA concentration-response curve of α4β2δ GABAARs was significantly shifted upwards in the presence of kavain. The preference of kavain for α4β2δ GABAARs can be explained by the partial-agonist, weak-intrinsic-efficacy profile of GABA at this receptor subtype (section 1.11.1) that renders it more sensitive to kavain modulation. Similar differential enhancement effects that arise from the distinct GABA efficacies at synaptic

αβγ and extrasynaptic αβδ receptors have been described for other allosteric ligands such as etomidate [38], propofol [39], pentobarbital [40], and neurosteroids [41]. While our findings do not conclusively prove that GABAARs are the in vivo substrate for kavain, it is tempting to speculate that kavain may have a larger impact on extrasynaptic than

127 synaptic GABAARs physiologically. This conjecture is substantiated by previous studies that found more prominent kavain or kavalactones actions in brain regions such as the hippocampus [21, 42], in which tonic GABAergic conductances mediated via δ- containing GABAARs have been detected [43]. Future electrophysiological studies comparing kavain activity in neurons from wild-type and α4- or δ-knockout mice will be necessary to confirm this speculation.

The low affinity of kavain detected in our study is in agreement with most studies conducted in the past using kava extracts or pure kavalactones [21, 22, 44]. The high- micromolar concentrations required to observe the pharmacological actions of kavalactones in vitro raise the critical question of whether the concentrations used in these studies are indeed physiologically relevant. Studies conducted in mice showed that 100 mg/kg of kavain alone caused a rapid surge in brain concentration (up to 100 µM) within minutes after i.p. injection [22, 45]. A higher brain concentration of kavain was found

(120 µM) in the presence of other kavalactones (44 mg kavain in 120 mg/kg kavalactone mixture; i.p.), clearly demonstrating pharmacokinetic synergism [22, 45]. Davies et al.

(1992) predicted higher brain concentrations (300 µM) with the administration of larger doses (300 mg/kg) of kavain [22]. Thus, kavain concentrations used in our study appear to correlate well with the concentrations needed to induce anxiolysis and hypnosis in mice

[27, 46]. Unfortunately, the relevance of this concentration range cannot be established for humans as no data is currently available [47].

The possibility of kavalactones interacting non-specifically with lipid membrane of neurons to exert their psychoactive effects has been raised [21, 22]. This hypothesis is likely to stem from the historical Meyer-Overton theory which postulates that for a lipid-

128 soluble anaesthetic agent, once a threshold number of dissolved molecules in lipid bilayer is reached, the physical distortion of neuronal membrane triggers changes in brain functions [48]. However, this theory has been contested and fallen out of favour, as converging lines of evidence obtained using electrophysiological, mutational, and mouse genetic approaches prove that anaesthetics do bind to distinct sites found on LGICs to directly exert their clinical effects [49]. Our mutational studies revealed a near-complete loss of kavain potentiation at α1β3N265Mγ2L GABAARs (Figure 4.14). The extensively studied β3N265M point mutation is known to selectively abolish the in vitro and in vivo sensitivity of anaesthetics such as etomidate, propofol, pentobarbital and volatile agents

(section 1.11.3.5). As such, this finding argues strongly against the lipid hypothesis, and supports a direct kavain-GABAAR interaction.

We also investigated the impact of mutating two homologous methionine residues

(α1M236 and β2M286) that line the transmembrane β+α‒ anaesthetic binding sites

(section 1.11.3.1). Substituting the methionine residues with tryptophan decreases etomidate and propofol sensitivity (Figure 4.12), an effect which has been correlated with increased side-chain volume of tryptophan sterically hindering anaesthetic binding [34].

Unlike β3N265M, the α1M236W and β2M286W point mutations did not negatively affect the activity of kavain (Figure 4.14), suggesting that either the binding determinants of kavain are different from etomidate and propofol, or kavain does not bind to the transmembrane β+α‒ interfaces. The enhanced agonist efficacy of kavain observed can be explained by the higher open probability of these spontaneously-gated mutant receptors [34].

129

We tentatively infer, based on the preliminary mutational evidence, that kavain binds to the transmembrane anaesthetic sites. Two observations support this notion: (1) The

β3N265M point mutation impairs the sensitivity of anaesthetics agents that bind to homologous transmembrane cavities, but not benzodiazepines and neurosteroids which target binding sites of dissimilar locations (section 1.11.3.5). (2) Kavain displays a remarkably similar pharmacological profile to the volatile anaesthetics which exhibit low affinity (EC50 > 200 µM) and steep dose-response relationships, besides lacking subtype selectivity at GABAARs [50, 51]. Future investigation into the volatile anaesthetic determinants αS270 and αL277 [52]; other residues in the α+β‒, γ+β‒ and β+β‒ interfaces identified from photolabelling studies using anaesthetic analogues (section 1.11.3); and residues in other regions is warranted to provide a more comprehensive picture of kavain binding site(s). Given the absence of subtype specificity in kavain modulation at

GABAARs, the existence of multiple binding sites for kavain is highly probable.

As part of our efforts in characterising the interaction of kavain and other GABAAR modulators on a receptor level, we found that kavain action was not blocked by flumazenil

(Figure 4.6). This finding corroborates previous studies that failed to detect any significant interaction of kavalactones with the high-affinity benzodiazepine site [19, 21-

23, 46]. In addition, kavain and diazepam modulated GABAARs in a less-than-additive manner, contrary to the speculated synergistic (supra-additive) interaction [30]. The co- application of kavain and other GABAAR positive modulators such as allopregnanolone,

DS2, etomidate and propofol also produced similar less-than-additive potentiation.

Typically, two positive modulators that bind to distinct, independent binding sites give rise to additive potentiation when combined, as exemplified by sevoflurane and propofol

[53], and diazepam and valerenic acid [54]. While the possibility of a competitive nature of these interactions cannot be denied, it seems unlikely that kavain binds to all the

130 implicated binding sites which are geographically distinct. It is more plausible that the putative kavain binding site(s) are allosterically coupled to these sites in an inhibitory fashion, resulting in less-than-additive joint enhancement. Alternatively, kavain may affect a common transduction pathway that reduces the enhancement efficacy by these modulators at GABAARs. It should be mentioned that most of these experiments were conducted with single ligand concentrations (with the exception of diazepam), and accurate determination of the nature of these interactions demands isobologram analysis.

Despite the long history of kava consumption and the wealth of clinical evidence in favour of the efficacy of kavalactones in treating anxiety, there is a severe gap in our understanding of the molecular target(s) and the mechanism of action of these psychoactive compounds. The present study provides evidence that supports a direct interaction of kavain, the major kavalactone with GABAARs in vitro. The enhancement of GABAAR function by kavain occurs in a subtype-unselective and flumazenil- insensitive manner, and the underlying mechanism may be similar to anaesthetic agents.

Future studies comparing the behavioural responses of kavalactones in wild-type and mutant mice bearing the β3N265M point mutation will provide stronger evidence that

GABAARs mediate the actions of kavalactones in vivo. Once this is established, the identification of specific GABAAR-enhancing kavalactones will give impetus in the development and refinement of these kava-derived chemicals as novel anxiolytics.

131

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Chapter 5

General discussion: Implications from the functional expression of βγ GABAARs

______

Our work with the semi-synthetic flavonoid 2′MeO6MF revealed that the combination of the β3 and γ2L subunits formed fully functional receptors in Xenopus oocytes (Chapter

3). Furthermore, depending on the injection ratios used, β3γ2L receptors showed unique pharmacological profile in response to GABA, 2′MeO6MF, etomidate and propofol, indicating the existence of multiple stoichiometric forms. We believe that the functional expression of βγ receptors has important implications, both in the study of GABAARs, as well as in drug discovery. We discuss these implications in more detail below.

5.1. Flexibility in the subunit requirement of ligand binding sites

One of the most striking aspects of βγ receptors is that it defies current understanding of

GABAAR pharmacology. It lacks the α subunit which has been established to constitute the complementary faces of the GABA, etomidate and neurosteroid binding sites, as well as the principal face of the high-affinity benzodiazepine site (Chapter 1). Nevertheless, we and others have shown that βγ receptors are responsive to these ligands [1, 2]. While it is possible that hitherto unidentified binding sites are responsible for these responses, a more logical explanation would be alternative subunit interfaces harbouring homologous orthosteric and allosteric binding sites in these non-α-containing receptors. In other

135 words, the structural requirement for these binding sites may not be as strict as they are generally perceived.

5.1.1. GABA binding sites at the β+γ‒ interfaces?

One of the possible explanations for the GABA sensitivity of βγ receptors is that the γ subunit can substitute for the α subunit to accommodate for GABA binding at the β+γ‒ and/or γ+β‒ interfaces. However, as the γ+β‒ interface is present in the αβγ receptors, and is not implicated in GABA binding [3], the β+γ‒ interface is a more plausible candidate. The conventional β+α‒ interface has an “aromatic box” formed by βY97,

βY157, βF200, βY205, and αF64 residues which is essential in agonist recognition

(Figure 1.8B) [4-6]. The integrity of this aromatic box is likely to be maintained at the

β+γ‒ interfaces, as the homologous residue in the γ subunit that corresponds to αF64 is also a phenylalanine (γF77; Figure 5.1).

Figure 5.1. Aligned sequence of the α1 subunit that forms the complementary face of the GABA binding site, and the corresponding region in the γ2 subunit (which forms the complementary face of the benzodiazepine site). Residues important for binding or function of orthosteric ligands are highlighted in red, whereas the residues implicated in benzodiazepine sensitivity are highlighted in turquoise. Residue α1F64 (indicated by *) is an important determinant of GABA binding and gating. Homology modelling predicts α1R66 (indicated by ‡) to form salt bridges, and α1T129 (indicated by §) to form hydrogen bonds with GABA. The sequence numbering is based on the alignment presented in Bergmann et al. (2013) [4]. The secondary structures indicated below the sequence are based on the X-ray crystal structure of the human β3 homomeric GABAAR [7].

136

Furthermore, other α subunit residues implicated in GABA binding/function such as

α1L127, α1T129 (predicted to form a hydrogen bond with GABA) and α1R131 are conserved at the corresponding positions in the γ2 subunit, except for α1R66. As α1R66 was predicted to form salt bridges with GABA by Bergmann et al. (2013) [4], we postulate that this interaction is not possible in βγ receptors as the bidentate arginine is replaced with a non-polar alanine residue in the γ2 subunit (γ2A79), which may underlie the > 100-fold lower GABA potency at β2γ2L compared to α1β2 receptors (Figure 4.3,

Table 4.3). Hence, it is of interest to investigate whether the γ2A79R mutation will enhance GABA sensitivity of βγ receptors to resemble that of αβ or αβγ receptors. Besides that, mutational studies targeting residues found on the β+γ‒ interfaces which correspond to those found in the β+α‒ GABA binding sites will help establish the existence of GABA binding sites at βγ receptors.

5.1.2. High-affinity benzodiazepine site at the β+γ‒ interface?

The possibility for the β subunit to form the high-affinity benzodiazepine site with the γ subunit in the absence of the α subunit was suggested following the discovery that diazepam modulated α1β2γ2 and β2γ2 receptors to similar extent in several studies

(Figure 5.2A and B) [1, 2, 8]. In concordance with these findings, we also detected diazepam potentiation at β2γ2L receptors (Figure 5.2C). As part of the effort to identify the binding determinants of diazepam, Amin et al. (1997) showed that mutating two tyrosine residues in the principal face of the α1 subunit (α1Y159 and α1Y209) to serine significantly diminished diazepam sensitivity at α1β2γ2 receptors [2]. In contrast, when the corresponding conserved tyrosine residues in the γ2 subunit were mutated to serine

(γ2Y172S and γ2Y220S), modest reduction in diazepam modulation was detected at

α1β2γ2Y172S and α1β2γ2Y220S receptors. These mutations had no effects on diazepam

137 action at β2γ2Y172S and β2γ2Y220S receptors [2]. Based on these findings, we infer that the principal face of the γ2 subunit (i.e., γ+β‒ or γ+γ‒ interfaces) is unlikely to mediate diazepam modulation at the non-α-containing β2γ2 receptors.

While it is possible for the β+γ‒ interface to hold the benzodiazepine site, ascertaining this binding interface is expected to be challenging because residues found in the principal face of the β subunit (e.g., β2Y157 and β2Y205) are also involved in GABA binding

(Figure 5.2D). As such, mutations in this region will inevitably attenuate GABA sensitivity, which will complicate the interpretation of changes to diazepam’s GABA- modulatory action. Additionally, as we have proposed that the β+γ‒ interfaces to be the site of GABA action for βγ receptors, it is mechanistically unclear as to how diazepam can modulate these receptors via the same sites in the presence of GABA. Having said that, it is not impossible for benzodiazepines to bind to the GABA binding sites, as such possibility has recently been highlighted for flurazepam and zopiclone [9].

The α1H101 residue plays an integral role in diazepam sensitivity, and mutations of this residue greatly reduce diazepam affinity at α1βγ2 receptors (section 1.11.2.1). Thus, it is questionable that the non-conservative substitution of histidine with leucine at residue 99 in the β2 subunit (β2L99; Figure 5.2D) will confer high-affinity diazepam binding through the β+γ‒ interfaces. Despite the absence of this critical binding residue, diazepam potentiates β2γ2 receptors at nanomolar-to-low-micromolar concentrations. While a detailed mutational study is warranted to elucidate the mechanism underlying diazepam sensitivity at these α-lacking receptors, a possible explanation is that diazepam’s contact points at βγ receptors are different from those determined at αβγ receptors. This hypothesis is supported by the considerable variability reported previously for the binding

138 determinants of (1) different benzodiazepine site ligands in the α1+γ2‒ interface [10], and

(2) the same ligand in different αx+γ2‒ (x = 1, 2, 3, 5 or 6) interfaces [11].

Figure 5.2. Diazepam potentiation at β2γ2 GABAARs. Comparable diazepam actions at α1β2γ2 and β2γ2 GABAARs reported by (A) Whittemore et al. (1996) and (B) Amin et al. (1997). (B) The γ2Y172S and γ2Y220S point mutations had not effect on diazepam activity at β2γ2 receptors. The corresponding mutations in the α1 subunit significantly reduced diazepam potentiation at α1β2γ2 receptors. (C) Continuous traces demonstrating the application of control (10 µM GABA); control and 1 µM diazepam; followed by two consecutive applications of control at β2γ2L (1:20) receptors (n = 4). (D) Aligned sequence of the α1 subunit that forms the principal face of the benzodiazepine binding site, and the corresponding region in the β2 subunit (which line the principal face of the orthosteric site). Residues important for binding or function of benzodiazepines are highlighted in turquoise, whereas the residues implicated in GABA actions are highlighted in red. Residue α1H101 (indicated by *) is an important in vitro and in vivo determinant for diazepam (section 1.11.2.1). Homology modelling predicts α1Y159 (indicated by ‡), α1T206 (indicated by §) and α1Y209 (indicated by ¶) to form polar contacts with diazepam. The sequence numbering is based on the alignment presented in Bergmann et al. (2013) [4]. The secondary structures indicated below the sequence are based on the X-ray crystal structure of the human β3 homomeric GABAAR [7].

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5.1.3. Other allosteric binding sites

Besides GABA and diazepam, we and others have shown that many allosteric ligands such as etomidate, propofol, loreclezole, pentobarbital, 3α-hydroxy-5α-pregnan-20-one

(3α,5α-P) and mefenamic acid also have comparable actions at αβγ and βγ receptors

(Figure 5.3) [1, 12]. These functional data reflect that homologous ligand binding sites may exist in βγ receptors, but may not have been detected in αβ or αβγ receptors previously. In keeping with this postulation, evidence from photoaffinity labelling studies demonstrated that homologous transmembrane anaesthetic binding sites do exist at distinct subunit interfaces (β+α‒, α+β‒, γ+β‒, or β+β‒) depending on the GABAAR subtypes investigated (section 1.11.3).

Figure 5.3. Comparable actions of various allosteric ligands at αβγ and βγ GABAARs. (A) Modulatory effects of pentobarbital (PB), 3α-hydroxy-5α-pregnan-20- one (3α,5α-P), mefenamic acid (MA), and loreclezole (LZ) reported by Whittemore et al. (1996) [1]. (B) Direct activation of α1β2γ2S and β2γ2S receptors by propofol and its derivative 4-iodo-2,6-diisopropylphenol (4-I-Pro) reported by Sanna et al. (1999) [12].

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5.2. Possibility of mixed βγ and αβγ GABAARs

A common problem in the heterologous expression of recombinant GABAARs is the presence of mixed αβ and αβγ receptor populations which compromises the accuracy in measuring a receptor property of interest. To circumvent this problem, an injection ratio which is heavily biased toward the γ2 subunit is usually used when expressing αβγ receptors in Xenopus oocytes. Boileau et al. (2002) found that an injection ratio of 1:1:10 for α1:β2:γ2 subunit mRNAs was necessary to produce “a purer population of α1β2γ2 receptors” characterised by a lower GABA potency (~4-fold) than α1β2 (1:1) receptors

(Figure 5.4A), and consistent measurement of diazepam potentiation [13]. However, studies conducted by Baburin et al. (2008) showed that while increasing the relative amount of the γ subunit resulted in greater degree of diazepam potentiation, a great portion of αβ receptors was still present even at a 1:1:20 ratio, contributing to the discrepancy in GABA potency measured at α1β2γ2 receptors formed from free subunits and concatenated constructs (Figure 5.4B) [14].

In contrast to these studies, under our experimental conditions, we found that an injection ratio of 1:1:2 (α2:β2:γ2L) reduced the GABA potency approximately 7 fold compared to

α2β2 (1:1) receptors (Figure 5.4C). Increasing the ratio to 1:1:10 resulted in ~55-fold decrease in GABA potency, and the concentration-response curve displayed a low Hill slope (nH = 0.73). Notably, the GABA concentration-response curve of α2β2γ2L (1:1:10) receptors overlaps with that of β2γ2L (1:10) receptors, raising a cautionary note on the possibility of mixed βγ and αβγ populations. This hypothesis, however, will be difficult to test as both αβγ and βγ receptors are potentiated by diazepam (Figure 5.2) and are insensitive to Zn2+ inhibition (Figure 5.5), thus precluding these compounds as diagnostic tools. Nevertheless, based on these studies, it is clear that a one-ratio-fits-all solution does

141 not exist, and the ratio chosen needs to be tailored according to experimental parameters such as the quality of mRNA, the plasmid vector, the origin of subunit cDNAs and the subunit splice variants, as well as pharmacological evidence.

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Figure 5.4. GABA concentration-response curves of αβ, αβγ, and βγ GABAARs expressed in Xenopus oocytes using different injection ratios from pharmacological studies. The curve parameters are summarised in tables on the right panel. Figures were taken from (A) Boileau et al. (2002) [13], (B) Baburin et al. (2008) [14], and (C) the present study.

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Figure 5.5. Zn2+ sensitivity at α1β2, β2γ2L and α1β2γ2L receptors. Current traces demonstrating the co-application of 10 µM Zn2+ with a GABA concentration close to 2+ EC50. Zn inhibits the GABA-elicited current at α1β2 receptors (n = 8), but has no effects at β2γ2L (n = 5) and α1β2γ2L (n = 9) receptors.

5.3. Perspective

To our knowledge, there are at least 15 studies to date (excluding the present study) that have demonstrated the in vitro functional expression of βγ receptors [1, 2, 8, 12, 15-25].

Despite this level of evidence, βγ receptors are still not routinely used as a tool receptor in pharmacological studies, perhaps due to (1) the reluctance to accept the evidence for its formation as it contradicts the general consensus that only the αβ and αβγ combinations can access cell surface efficiently to form functional receptors; (2) the questionable physiological existence of βγ receptors; and/or (3) the “erratic” nature of βγ receptors reported by some studies [23]. Majority of studies have in the past inferred the subunit requirement of a chemical of interest’s binding site(s) based on its activity at recombinant

αβ and αβγ receptors, and mutational studies designed based on these data had led to the identification of binding/function determinants.

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In our view, this approach is incomplete and potentially misleading. The heteromeric nature of GABAARs, coupled with the interfacial nature of most ligand binding sites necessitate pharmacological analysis of different receptor subtypes to fully understand the subunit requirement for a ligand of interest. For these reasons, we believe that the inclusion of βγ receptors in pharmacological studies will provide a more comprehensive picture of a ligand binding site, besides preventing the overestimation of a ligand’s subtype selectivity. It is worth mentioning at this stage that the recent work of Maldifassi et al. (2016) provided evidence that etomidate, propofol and pentobarbital interact with the γ+β‒ interface in α1β2γ2 receptors [26]. This finding does not come as a surprise as these ligands have all been shown to be functional at βγ receptors.

The fact that we observed pharmacological differences with β3γ2L GABAARs expressed using different injection ratios (Chapter 3) adds another layer of complexity to this discussion. However, as the experimental approaches used in our study do not provide direct evidence that distinct subunit stoichiometries contribute to the remarkable variability in the pharmacology of β3γ2L GABAARs, the use of concatenated constructs in the future will be necessary to confirm our findings. Nevertheless, the functional data obtained using 2′MeO6MF (Figure 3.5) clearly show that non-optimal injection ratios not only cause receptors of different subunit composition to be expressed, but mixed stoichiometric forms of the same receptor subtype could also affect the accuracy of pharmacological studies.

In conclusion, there is now strong evidence indicating more complex GABAAR pharmacology than previously anticipated. Hence, it is of utmost importance that future

145 drug discovery and structure-function analysis efforts using recombinant receptors to be designed and interpreted with this level of complexity in mind.

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human gamma-aminobutyric acidA multiple drug-binding sites. In: Uwe R, receptor α4 subunit expressed in Xenopus editor. Advances in Pharmacology. laevis oocytes. Molecular Pharmacology. Volume 72: Academic Press; 2015. p. 53- 1996;50(5):1364-75. 96.

2. Amin J, Brooks-Kayal A, Weiss DS. Two 10. Morlock EV, Czajkowski C. Different

tyrosine residues on the α subunit are residues in the GABAA receptor crucial for benzodiazepine binding and benzodiazepine binding pocket mediate allosteric modulation of γ-aminobutyric benzodiazepine efficacy and binding.

acidA receptors. Molecular Pharmacology. Molecular Pharmacology. 2011;80(1):14- 1997;51(5):833-41. 22.

3. Amin J, Weiss DS. GABAA receptor needs 11. Luscher BP, Baur R, Goeldner M, Sigel E.

two homologous domains of the β-subunit Influence of GABAA receptor α subunit for activation by GABA but not by isoforms on the benzodiazepine binding pentobarbital. Nature. site. PloS one. 2012;7(7):e42101. 1993;366(6455):565-9. 12. Sanna E, Motzo C, Usala M, Serra M, 4. Bergmann R, Kongsbak K, Sorensen PL, Dazzi L, Maciocco E, et al. Sander T, Balle T. A unified model of the Characterization of the

GABAA receptor comprising agonist and electrophysiological and pharmacological benzodiazepine binding sites. PloS one. effects of 4-iodo-2,6-diisopropylphenol, a 2013;8(1):e52323. propofol analogue devoid of sedative- anaesthetic properties. British Journal of 5. Laha KT, Tran PN. Multiple tyrosine Pharmacology. 1999;126(6):1444-54. residues at the GABA binding pocket influence surface expression and mediate 13. Boileau AJ, Baur R, Sharkey LM, Sigel E,

kinetics of the GABAA receptor. Journal of Czajkowski C. The relative amount of Neurochemistry. 2013;124(2):200-9. cRNA coding for γ2 subunits affects

stimulation by benzodiazepines in GABAA 6. Lynagh T, Pless SA. Principles of agonist receptors expressed in Xenopus oocytes. recognition in Cys-loop receptors. Neuropharmacology. 2002;43(4):695-700. Frontiers in Physiology. 2014;5. 14. Baburin I, Khom S, Timin E, Hohaus A, 7. Miller PS, Aricescu AR. Crystal structure Sieghart W, Hering S. Estimating the of a human GABA receptor. Nature. A efficiency of benzodiazepines on GABAA 2014;512(7514):270-5. receptors comprising γ1 or γ2 subunits. British Journal of Pharmacology. 8. Boileau AJ, Kucken AM, Evers AR, 2008;155(3):424-33. Czajkowski C. Molecular dissection of benzodiazepine binding and allosteric 15. Sigel E, Baur R, Trube G, Möhler H, coupling using chimeric γ-aminobutyric Malherbe P. The effect of subunit acid receptor subunits. Molecular A composition of rat brain GABAA receptors Pharmacology. 1998;53(2):295-303.

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subunits. European Journal of K, Taylor PM, Moss SJ, et al. GABAA Neuroscience. 1992;4(1):1-9. receptor composition is determined by distinct assembly signals within α and β 17. Im HK, Im WB, Hamilton BJ, Carter DB, subunits. Journal of Biological Chemistry. Vonvoigtlander PF. Potentiation of γ- 2003;278(7):4747-55. aminobutyric acid-induced chloride currents by various benzodiazepine site 23. Hamon A, Morel A, Hue B, Verleye M, agonists with the α1γ2, β2γ2 and α1β2γ2 Gillardin J-M. The modulatory effects of

subtypes of cloned γ-aminobutyric acid the anxiolytic etifoxine on GABAA type A receptors. Molecular receptors are mediated by the β subunit. Pharmacology. 1993;44(4):866-70. Neuropharmacology. 2003;45(3):293-303.

18. Sanna E, Mascia MP, Klein RL, Whiting 24. Miko A, Werby E, Sun H, Healey J, Zhang PJ, Biggio G, Harris RA. Actions of the L. A TM2 residue in the β1 subunit general anesthetic propofol on determines spontaneous opening of

recombinant human GABAA receptors: homomeric and heteromeric γ- influence of receptor subunits. Journal of aminobutyric acid-gated ion channels. Pharmacology and Experimental Journal of Biological Chemistry. Therapeutics. 1995;274(1):353-60. 2004;279(22):22833-40.

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Chapter 6

GABAARs as the molecular target for kavalactones: Future directions

______

It has almost been 30 years since GABAARs were first proposed to mediate the in vivo effects of kavalactones. However, a lack in robust experimental evidence has led to ambiguity in the explication of a GABAergic mechanism for these natural products. In a pharmacological study conducted using kavain as a representative kavalactone on a range of human recombinant GABAAR subtypes expressed in Xenopus oocytes, we demonstrated that kavain directly potentiated GABAARs with no subtype selectivity

(Chapter 4). We also presented evidence that kavain potentiation did not occur via the high-affinity benzodiazepine binding site, and the underlying mechanism of kavain may be common to the GABAAR-targeting anaesthetics. While these findings are encouraging, many aspects of kavalactone actions at GABAARs remain to be explored.

6.1. Effect of enantiomerically pure kavain

With the exception of the achiral yangonin and desmethoxyangonin, other major kavalactones such as kavain, dihydrokavain, methysticin and dihydromethysticin have a stereocenter, and only the (+)-enantiomer forms occur naturally (Figure 4.1). Despite the homochirality, majority of the studies investigating the in vitro and in vivo effects of individual kavalactones have used the racemic mixture, as it is the only commercially available form. For the same reason, (±)-kavain (or DL-kavain) was used in our study.

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However, given the precedents for chiral ligands to exhibit opposite modulatory effects at GABAARs [1], the possibility of (-)-kavain confounding the determination of (+)- kavain pharmacology should not be excluded. The pharmacological characterisation of

(+)-kavain obtained through chemical synthesis [2] or chromatographic isolation [3] in the future will provide further insight into the structure-activity relationship of kavain activity at GABAARs.

6.2. Potential for synergistic modulation of kavalactones

A synergistic pharmacodynamic interaction of kavalactones has been suspected as the pharmacological activity of kava extracts is usually greater than individual kavalactones

[4]. We have preliminary data indicating comparable GABA enhancement effects by yangonin, methysticin and dihydromethysticin to kavain at α1β2γ2L GABAARs (data not shown). Hence, future studies investigating the effects of these kavalactones in different combinations (pairs, triplets, or quadruplets) will help clarify the nature of their interactions, as well as identifying the most efficacious kavalactone combination(s) at

GABAARs.

6.3. Effect of kavalactones in transgenic mice

The use of knock-in mice bearing the β3N265M point mutation has aided the establishment of GABAARs as one of the major in vivo targets of general anaesthetics

(section 1.11.3) [5]. Our mutational studies clearly demonstrated that the β3N265M mutation strongly impaired kavain potentiation in vitro. We expect that the analysis of kavalactones in the transgenic mouse line will allow us to correlate any attenuated behavioural responses with GABAARs containing the mutant β3 subunit. While

149 speculative, we do not think that the transgenic mice will be completely resistant to kavalactones-induced responses as other targets (different GABAAR subtypes or other receptors/enzymes) may also contribute to the in vivo effects of kavalactones.

Nevertheless, a moderate reduction in behavioural response(s), together with evidence from electrophysiological and mutational studies will provide more definitive evidence that specific GABAAR subtypes do mediate kavalactone actions.

6.4. Concluding remarks

The findings from clinical trials on the efficacy of medically-prescribed kava extracts containing standardised, enriched kavalactones in the treatment of anxiety are encouraging. Besides alleviating the debilitating symptoms associated with anxiety, kava shows no tolerance, addictive and withdrawal tendencies [6]. Together with the mild-to- negligible side effects reported in these human studies [7, 8], the presumed bioactive constituent kavalactones are attractive novel anxiolytics. However, as with most natural products, the effective clinical use of kavalactones suffers from the lack of understanding of their biological targets. Our pharmacological study with kavain provides evidence that the naturally-abundant kavalactone directly enhances GABAAR function. These findings serve as a starting point for future target-validation studies.

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enantiomer on the α1β3γ2L GABAA receptor account for their in vivo effects. 6. Sarris J, Stough C, Teschke R, Wahid ZT, The Journal of physiology. Bousman CA, Murray G, et al. Kava for 2015;593(22):4943-61. the treatment of generalized anxiety disorder RCT: Analysis of adverse 2. Smith TE, Djang M, Velander AJ, Downey Reactions, liver Function, addiction, and CW, Carroll KA, van Alphen S. Versatile sexual effects. Phytotherapy Research. asymmetric synthesis of the kavalactones: 2013;27(11):1723-8. First synthesis of (+)-kavain. Organic Letters. 2004;6(14):2317-20. 7. Sarris J, LaPorte E, Schweitzer I. Kava: A comprehensive review of efficacy, safety, 3. Cirilli R, Ferretti R, Gallinella B, Bilia AR, and psychopharmacology. Australian and Vincieri FF, La Torre F. Enantioseparation New Zealand Journal of Psychiatry. of kavain on Chiralpak IA under normal- 2011;45(1):27-35. phase, polar organic and reversed-phase conditions. Journal of separation science. 8. Sarris J, Stough C, Bousman CA, Wahid 2008;31(12):2206-10. ZT, Murray G, Teschke R, et al. Kava in the treatment of generalized anxiety 4. Ramzan I, Tran V. Chemistry of kava And disorder: a double-blind, randomized, kavalactones. In: Singh YN, editor. Kava placebo-controlled study. Journal of from Ethnology to Pharmacology: CRC clinical psychopharmacology. Press; 2004. p. 76-103. 2013;33(5):643-8.