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GPCR modulation of TRPM3 and other TRPM ion channels

Alkhatib, Omar

Awarding institution: King's College London

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GPCR modulation of TRPM3 and other TRPM ion channels

Thesis submitted for the degree of

Doctor of Philosophy

King’s College London

Omar Alkhatib

Wolfson Centre for Age-Related Diseases Institute of Psychiatry, Psychology & Neuroscience King’s College London

2016-2020

Abstract

The transient receptor potential (TRP) superfamily is composed of 28 non-selective cation channels, which have been implicated in a myriad of physiological processes ranging from temperature detection and sensory transduction to insulin release, gene transcription and ionic homeostasis. G-protein coupled receptors (GPCRs) are the largest family of signalling proteins involved in all physiological processes and are targeted by many established pharmacotherapies. TRP channels are often effectors activated or modulated by GPCR signalling. The aim of this thesis is to functionally characterise the TRP melastatin 3 (TRPM3) channel, its modulation by GPCRs and examine GPCR regulation of other members of the

TRPM subfamily, specifically TRPM2, TRPM7 and TRPM8 channels.

The ionic current characteristics produced by the two selective TRPM3 agonists, pregnenolone sulphate (PS) and CIM0216, were investigated using patch-clamp electrophysiology and Ca2+-imaging techniques in a heterologous expression system. The findings demonstrate that activation of TRPM3 by CIM0216 but not PS strongly desensitises in the presence of extracellular Ca2+. Application of CIM0216 results in a biphasic TRPM3 current waveform and in two open states in single-channel recordings. TRPM3 activation by both agonists was fully inhibited at acidic pH (pH 5) but unaffected by alkaline pH (pH 8).

Moreover, TRPM3 activity both in vitro and in vivo was found to be inhibited by increasing

DMSO concentrations.

Activation of TRPM3 in heterologous and sensory neurons was promiscuously and robustly inhibited by GPCRs coupled to any of Gαi, Gαq and Gαs subunits. Investigations of GPCR signalling cascades on TRPM3 demonstrated that GPCR-mediated inhibition of TRPM3 is due to the direct interaction of liberated Gβγ subunits with the channel. Behavioural

2 investigations in vivo demonstrated that activation of GPCRs that are targeted by analgesic drugs as well as by the proinflammatory mediators prostaglandin E2 and bradykinin inhibit

TRPM3-mediated nociception and inflammatory heat hypersensitivity.

The effect of GPCR activation was also examined on other members of the TRPM subfamily.

My results demonstrate that GPCRs coupled to Gαi/o-, Gαq- and Gαs do not appear to influence heat- or H2O2-evoked TRPM2 activity. In contrast, TRPM7 was activated reversibly and transiently following stimulation of Gαs-coupled receptors, whereas TRPM8 was exclusively inhibited by stimulation of Gαq-coupled receptors.

In conclusion, this thesis demonstrates that TRPM3 channels are thus far the only TRPM family member to be promiscuously inhibited by GPCRs that couple to Gαi/o, Gαq and Gαs.

Acknowledgements

First and foremost, I would like to thank David for his support and supervision throughout the past years. His extensive knowledge is truly amazing and something I aspire to have one day. I would also like to thank Stuart for his constant support from the beginning of my MSc degree and for opening great opportunities for me including introducing me to David and helping me acquire the PhD position.

Being a part of David’s lab has been a privilege and I have met truly great people. I would like to thank my good friend Nurjahan for our frequent discussions about life and our futures – they were great therapeutic sessions. I would like to thank Robson for his guidance and help with my PhD especially with the in vivo experiments – I will definitely miss our coffee/lunch breaks and your Portuguese language lessons! My experience at David’s lab would have been completely different without my friend Ulku whose humour and attitude brightened up my days. I would like to thank Clive for his help and for giving me access to his extensive in vivo expertise. I also want to thank Eva for her help with all-things molecular! I want to thank Nisha for her support and guidance throughout my PhD. Finally, I would like to thank Ivona for all her help with passaging cells and keeping my cell lines alive while I was gone!

I would like to thank my family for their support and love throughout my PhD. Lastly, I would like to thank Shireen for her support and ability to cope with my overbearing stress!

Table of Contents

Abstract ...... 2 Acknowledgements ...... 4 Table of Figures ...... 10 Chapter 1. General introduction ...... 13 1.1 Introduction to transient receptor potential channels ...... 14 1.1.1 Discovery ...... 14 1.1.2 Structure ...... 14 1.1.3 Sub-families ...... 17 1.1.4 Mammalian TRP channels ...... 18 1.1.4.1 TRPA ...... 18 1.1.4.2 TRPC ...... 18 1.1.4.3 TRPV ...... 19 1.1.4.4 TRPML ...... 21 1.1.4.5 TRPP ...... 21 1.1.5 TRPM sub-family ...... 22 1.1.5.1 TRPM1 ...... 22 1.1.5.2 TRPM2 ...... 24 1.1.5.3 TRPM3 ...... 26 1.1.5.4 TRPM4 ...... 27 1.1.5.5 TRPM5 ...... 27 1.1.5.6 TRPM6 ...... 28 1.1.5.7 TRPM7 ...... 28 1.1.5.8 TRPM8 ...... 29 1.2 Introduction to G-protein coupled receptors ...... 29 1.2.1 Discovery ...... 29 1.2.2 Families ...... 31 1.2.2.1 Glutamate family (class C) ...... 31 1.2.2.2 Rhodopsin family (class A) ...... 31 1.2.2.3 Adhesion family ...... 32 1.2.2.4 Frizzled family (class F) ...... 32 1.2.2.5 Secretin family (class B) ...... 32 1.2.3 GPCR signalling dynamics ...... 33 1.2.3.1 Gα- and Gβγ-mediated signalling ...... 34

1.2.3.2 GPCR kinases and arrestins ...... 35 1.3 GPCR regulation of ion channels ...... 39 1.3.1 GPCR regulation of GIRK channels ...... 41 1.3.2 GPCR regulation of VGCCs ...... 42

1.3.3 GPCR regulation of Kv7 channels ...... 43 1.3.4 GPCR regulation of TRP channels...... 43 1.3.4.1 TRPV1-4 ...... 43 1.3.4.2 TRPA1 ...... 45 1.3.4.3 TRPC1, 3, 4 and 5 ...... 45 1.4 Aims of this thesis ...... 46 Chapter 2. Materials and methods ...... 47 2.1 Cell culture ...... 48 2.1.1 Human embryonic kidney and Chinese hamster ovary cells ...... 48 2.1.1.1 TRPM3 HEK293 cell line ...... 48 2.1.1.2 TRPM2 HEK293 cell line ...... 48 2.1.1.3 pGLO HEK293 cell line ...... 49 2.1.1.4 Transiently transfected HEK293 and CHO cells ...... 49 2.1.2 Dorsal root ganglia neurons ...... 50 2.2 Imaging of intracellular calcium levels ...... 51 2.2.1 Fura-2 ...... 51 2.2.2 Microscope-based imaging of intracellular calcium levels ...... 51 2.3 Fluorometric measurement of intracellular calcium and cAMP levels ...... 52 2.4 Patch-clamp electrophysiology ...... 53 2.5 Behavioural assessment of pain responses ...... 54 2.6 Solutions and reagents...... 54 2.6.1 Solutions ...... 54 2.6.2 Reagents ...... 55 2.7 Patch-clamp and fluorometric assay data analysis ...... 56 2.8 Experimental design and statistical analyses ...... 57 Chapter 3. Characterisation of TRPM3 activity ...... 59 3.1 Introduction ...... 60 3.1.1 Introduction to TRPM3 ...... 60 3.1.2 TRPM3 gene and expression ...... 60 3.1.3 Activation of TRPM3 ...... 61 3.1.3.1 Pregnenolone sulphate and structurally related compounds ...... 61 3.1.3.2 Nifedipine ...... 63

3.1.3.3 D-erythro-sphingosine ...... 63 3.1.3.4 Clotrimazole and CIM0216...... 64 3.1.3.5 Hypotonicity ...... 65 3.1.4 Inhibition of TRPM3 ...... 65 3.1.4.1 Dihydropyridines ...... 65 3.1.4.2 Non-selective TRP inhibitors ...... 65 3.1.4.3 Monovalent and divalent cations ...... 66 3.1.4.4 Fenamates ...... 66 3.1.4.5 Progesterone and structurally related compounds ...... 67 3.1.4.6 Thiazolidinediones ...... 67 3.1.4.7 Flavonoids and ononetin...... 67 3.1.5 Permeation and biophysical properties of TRPM3 ...... 70 3.2 Aims of the present study ...... 74 3.3 Results ...... 75 3.3.1 Characterisation of heterologously-expressed TRPM3 ...... 75 3.3.1.1 Activation of TRPM3 by PS and CIM0216 in fluorometric assays ...... 75 3.3.1.2 Activation of TRPM3 by PS in whole-cell patch clamp electrophysiology ...... 77 3.3.1.3 Activation of TRPM3 by CIM0216 in whole-cell patch clamp electrophysiology ...... 80 3.3.1.4 TRPM3 desensitisation ...... 85 3.3.1.5 TRPM3 modulation by pH levels ...... 91 3.3.2 TRPM3 activation elicits nociceptive behaviour in vivo ...... 97 3.4 Discussion ...... 99 3.4.1 PS-induced activation of TRPM3 ...... 99 3.4.2 CIM0216-induced activation of TRPM3 ...... 101 3.4.3 Modulation of TRPM3 by acidic pH ...... 104 3.4.4 Activation of TRPM3 in vivo produces nociceptive behaviour ...... 105 3.5 Conclusion ...... 106 Chapter 4. Promiscuous GPCR inhibition of TRPM3 by Gβγ subunits ...... 107 4.1 Introduction ...... 108 4.1.1 GPCRs and analgesia ...... 108 4.1.1.1 Opioids ...... 108

4.1.1.2 GABAB ...... 109 4.1.1.3 NPY receptors ...... 110 4.1.1.4 Adenosine receptors ...... 111 4.1.2 GPCRs and inflammatory pain ...... 112 4.1.2.1 Prostaglandins ...... 112

4.1.2.2 Bradykinin ...... 114 4.1.3 TRPM3 in nociception and inflammatory hyperalgesia ...... 116 4.1.3.1 TRPM3 modulation by GPCRs ...... 117 4.2 Aims of the present study ...... 118 4.3 Results ...... 120

4.3.1 Gi/o-coupled GPCR regulation of TRPM3 activity ...... 120 4.3.1.1 Morphine, baclofen and PYY inhibit TRPM3-mediated PS-induced Ca2+ responses...... 120

4.3.1.2 Inhibition of TRPM3 is independent of cAMP and does not rely on Gαi proteins ...... 134 4.3.1.3 Beta-gamma subunits of Gi proteins mediate inhibition of TRPM3 ...... 136

4.3.1.4 Gi/o-coupled GPCRs modulate TRPM3-mediated nociceptive responses ...... 141

4.3.1.5 Adenosine inhibits TRPM3 in sensory neurons and in vivo through Gi/o-coupled A1 receptor activation ...... 143

4.3.2 Gq- and Gs-coupled GPCR regulation of TRPM3 ...... 146

4.3.2.1 Non Gi/o mediated inhibition of TRPM3 ...... 146

4.3.2.2 Gs-mediated inhibition of TRPM3 ...... 149

4.3.2.3 Gs-mediated inhibition of TRPM3 in sensory neurons ...... 152

4.3.2.4 PAR2 activation inhibits TRPM3 through promiscuous Gi/o- and Gq-mediated signalling ...... 156

4.3.2.5 Activation of endogenous Gq-coupled purinergic receptors Y inhibit TRPM3 activity in HEK293 cells ...... 160

4.3.2.6 Gq-mediated inhibition of TRPM3 channels by muscarinic M1 receptors ...... 163

4.3.2.7 Gq-mediated inhibition of TRPM3 in sensory neurons ...... 165

4.3.2.8 Gs- and Gq-mediated inhibition of TRPM3 is independent of PKA and PKC activity ...... 167 4.3.2.9 Gβγ subunits mediate inhibition of TRPM3 in heterologous and endogenous systems 169

4.3.2.10 Gs- and Gq-coupled GPCRs inhibit TRPM3 mediated nociception ...... 173 4.3.2.11 Identifying binding site of Gβγ protein on TRPM3 channels ...... 176 4.4 Discussion ...... 177

4.4.1 Gi/o-mediated inhibition of TRPM3 ...... 177

4.4.2 Gs- and Gq-mediated inhibition of TRPM3 ...... 182 4.5 Conclusion ...... 187 Chapter 5. GPCR regulation of TRPM2, TRPM7 and TRPM8 ...... 188 5.1 Introduction ...... 189 5.1.1 TRPM2 activation and modulation by GPCRs ...... 189 5.1.1.1 TRPM2 activation ...... 189 5.1.1.2 GPCR regulation of TRPM2 ...... 191 5.1.2 TRPM7 activation and modulation by GPCRs ...... 192

5.1.2.1 TRPM7 activation ...... 192 5.1.2.2 GPCR regulation of TRPM7 ...... 194 5.1.3 TRPM8 activation and modulation by GPCRs ...... 197 5.1.3.1 TRPM8 activation ...... 197 5.1.3.2 GPCR regulation of TRPM8 ...... 197 5.2 Aims of the present study ...... 201 5.3 Results ...... 202 5.3.1 GPCR modulation of TRPM2 ...... 202

5.3.1.1 GPCR modulation of H2O2-mediated activation of TRPM2 ...... 202 5.3.1.2 GPCR modulation of heat-mediated activation of TRPM2 ...... 204 5.3.2 GPCR modulation of TRPM7 ...... 207

5.3.2.1 Gi/o- and Gq-coupled GPCR modulation of TRPM7 ...... 207

5.3.2.2 Gs-coupled modulation of TRPM7 ...... 208 5.3.3 GPCR modulation of TRPM8 ...... 212

5.3.3.1 TRPM8 is solely regulated by Gαq proteins ...... 212 5.4 Discussion ...... 214 5.4.1 GPCR modulation of TRPM2 ...... 214 5.4.1.1 GPCR activation has no effect on TRPM2 activation by oxidative stress ...... 214 5.4.1.2 GPCR activation has no effect on heat-induced activation of TRPM2 ...... 214 5.4.2 GPCR modulation of TRPM7 ...... 215 5.4.3 GPCR modulation of TRPM8 ...... 216 5.5 Conclusion ...... 216 Chapter 6. General discussion, study limitations and future directions ...... 217 References ...... 224

Table of Figures

Figure 1-1 Structure of the TRPM8 channel ………………………………………………………………………………… 16

Figure 1-2 Phylogenetic tree of TRP channels ……………………………………………………………………………… 17

Figure 1-3 Mammalian TRPM subfamily phylogenetic tree …………………………………………………………. 22

Figure 1-4 mGluR6 regulation of TRPM1 activity on ON-bipolar cells ………………………………………….. 24

Figure 1-5 Differing levels of GPCR activity ……………………………………………………………………………..….. 34

Figure 1-6 Canonical G-protein and arrestin signalling pathways……………………………………..…..…….. 38

Figure 1-7 Mechanisms of GPCR regulation of ion channels ……………………………………………………..… 40

Figure 3-1 Gating of TRPM3 ………………………………………………………………………………………………………… 69

2+ Figure 3-2 PS elicits [Ca ]i-responses in TRPM3 expressing HEK293 cells ……………………………………. 76

2+ Figure 3-3 CIM0216 elicits [Ca ]i-responses in TRPM3 expressing HEK293 cells …………………………. 77

Figure 3-4 PS activates outwardly-rectifying currents in TRPM3 expressing HEK293 cells …………… 79

Figure 3-5 PS-mediated activation of TRPM3 leads to single channel openings in TRPM3 expressing HEK293 cells …………………………………………………………………………………………………………….. 80

Figure 3-6 CIM0216 activates double-rectifying currents in TRPM3 expressing HEK293 cells …….. 82

Figure 3-7 Biphasic CIM0216 activation of TRPM3 does not rely on binding site accessibility …….. 84

Figure 3-8 CIM0216-mediated activation of TRPM3 leads to two distinct channel openings in TRPM3 expressing HEK293 cells ………………………………………………………………………………………………. 85

Figure 3-9 Desensitisation of PS-induced inward currents is not rapidly reversible ……………………… 86

Figure 3-10 High concentrations of PS are required to desensitise TRPM3 inward currents ………… 87

Figure 3-11 PIP2 supplementation does not prevent desensitisation of PS-induced inward currents ………………………………………………………………………………………………………………………………………. 88

Figure 3-12 Desensitisation of PS-induced inward currents is not caused by Ca2+ entry ………………. 89

Figure 3-13 Desensitisation of CIM0216-induced inward currents is caused by Ca2+ entry …………… 90

Figure 3-14 TRPM3 activation by PS is inhibited at pH 5 ………………………………………………………………. 92

Figure 3-15 TRPM3 activation by CIM0216 is inhibited at pH 5 ………………………………………………….... 94

Figure 3-16 TRPM3 activation by PS is inhibited at pH 5 in whole-cell patch clamp recordings ……. 96

Figure 3-17 pH inhibition-response curves for TRPM3 activation by PS and CIM0216 …………………. 97

Figure 3-18 TRPM3 activation by PS and CIM0216 elicits nociceptive behaviour in mice that is

10 inhibited by high concentrations of DMSO …………………………………………………………………………………. 99

Figure 4-1 Morphine inhibits TRPM3 channels expressed on sensory neurons …………………………. 121

Figure 4-2 Activation of μ-opioid receptors inhibits TRPM3 ………………………………………………………. 125

Figure 4-3 Activation of other Gi-coupled receptors also inhibit TRPM3 ……………………………………. 128

Figure 4-4 CB1 activation can inhibit TRPM3 channels ………………………………………………………………. 131

2+ Figure 4-5 Morphine does not inhibit capsaicin evoked [Ca ]i-responses …………………………………..133

Figure 4-6 Opioid-mediated inhibition of TRPM3 is independent of cAMP and Gαi subunits ………135

Figure 4-7 βγ subunits mediate Gi/o inhibition of TRPM3 …………………………………………………………….137

Figure 4-8 Effect of Gβγ subunits on PS-evoked currents ……………………………………………………………140

Figure 4-9 Nociceptive responses to TRPM3 agonists are modulated by Gi/o GPCR ligands ………...142

Figure 4-10 Adenosine A1 receptors regulate TRPM3 activity …………………………………………………… 145

Figure 4-11 GPCR-mediated regulation of TRPM3 activity in HEK293 cells ………………………………… 148

Figure 4-12 Gs-coupled adenosine 2B receptor activation inhibits TRPM3-mediated responses in HEK293 cells ………………………………………………………………………………………………………… 151

Figure 4-13 Prostaglandin EP2 receptors inhibit TRPM3-mediated responses in sensory neurons 155

Figure 4-14 PAR2 receptor activation inhibits TRPM3-mediated responses through Gi/o- and

Gq/11-mediated signalling ………………………………………………………………………………………………………….. 159

Figure 4-15 Endogenous P2Y receptor activation inhibits TRPM3-mediated responses through

Gq/11-mediated signalling ………………………………………………………………………………………………………..… 162

Figure 4-16 Muscarinic M1 receptor induced inhibition of TRPM3 ………………………………………..…. 164

Figure 4-17 Bradykinin BK2 receptor activation inhibits TRPM3-mediated responses in sensory neurons ………………………………………………………………………………………………………………….…… 166

Figure 4-18 TRPM3 inhibition by butaprost and bradykinin is independent of PKA and PKC …..… 168

Figure 4-19 Gs- and Gq-coupled GPCR inhibition of TRPM3 is mediated by Gβγ protein ……………. 170

Figure 4-20 Gs- and Gq-coupled GPCR inhibition of TRPM3 in sensory neurons is reliant on

Gβγ protein ……………………………………………………………………………………………………………………………... 172

Figure 4-21 TRPM3 dependent antinociceptive and analgesic effects of Gs- and Gq-coupled GPCR activation prevents mouse nociceptive behaviour in response to TRPM3 agonists and reverses heat hyperalgesia in FCA-treated mice …………………………………………………………………………………………..… 175

Figure 4-22 TRPM3 peptide sequence does not sequester Gβγ protein …………………………….…….. 177

Figure 4-23 Schematic representation of promiscuous GPCR-mediated inhibition of TRPM3 ion channels ………………………………………………………………………………………………………………………………………. 187

Figure 5-1 GPCR activation does not regulate TRPM2 channel activation by H2O2 …………………….…. 203

Figure 5-2 GPCR activation does not regulate TRPM2 channel activation by heat ……………………….. 206

Figure 5-3 PAR2 activation does not regulate TRPM7 channel activity ………………………………………… 208

Figure 5-4 A2B activation potentiates TRPM7 channel activity …………………………………………..……….. 210

Figure 5-5 A2B-mediated potentiation of TRPM7 does not rely on PKA and Gβγ protein …………….. 210

Figure 5-6 TRPM8 activity is only regulated by Gq/11-mediated signalling …………………………………….. 212

Chapter 1. General introduction

1.1 Introduction to transient receptor potential channels

Transient receptor potential (TRP) channels form a heterogeneous superfamily of non- selective cation channels that act as cellular sensors for a wide spectrum of chemical and physical stimuli (Clapham, 2003; Clapham et al., 2001; Ramsey et al., 2006). TRP channels play crucial roles in various fundamental physiological processes including sensory transduction, cell survival, fertilisation and development. The superfamily is conserved in every metazoan organism that has been subject to sequence analysis (Montell, 2011)

1.1.1 Discovery

A mutation in the visual system of the fruit fly, Drosophila melanogaster, lead to the discovery of the first TRP channel (Cosens and Manning, 1969). Later studies showed that rhodopsin coupled to phospholipase C (PLC) activation opens cationic channels that evoke a depolarising membrane current in photoreceptor cells (Pedersen et al., 2005). The Drosophila TRP mutant exhibited a transient response to a continuous light stimulus rather than the wildtype sustained receptor potential (Cosens and Manning, 1969). The gene behind this transient receptor potential was hence named the trp gene and was cloned in 1989 (Montell and Rubin,

1989).

1.1.2 Structure

All TRP channels are membrane proteins with six transmembrane spanning regions (S1-S6) with a cation-permeable pore between the S5 and S6 region (Nilius and Owsianik, 2011).

Functional TRP channels exist as tetramers that can either be homo- or heteromultimeric.

The amino (N) and carboxy (C) termini of TRP channels are on the intracellular side of the membrane and vary in length between members of the subfamily and between different

14 splice variants of the same channel (Hellmich and Gaudet, 2014; Nilius and Owsianik, 2011).

Various TRP channels contain motifs such as the TRP motif, a conserved six amino acid sequence, calmodulin (CaM) binding domains and sites for phosphorylation and lipid interaction that are important for regulation of channel function (Nilius and Owsianik,

2011). Recently, the structure of the cold-sensing TRPM8 channel was determined using cryo-electron microscopy (Cryo-EM) (Figure 1-1) (Yin et al., 2018).

Figure 1-1 Structure of the TRPM8 channel

A Cryo-EM reconstruction. B Model of TRPM8 viewed from the extracellular side (left), from the membrane plane (middle) and from the cytosolic side (right).TMD: transmembrane channel domain, CD: cytosolic domain. C Topology diagram portraying the protein domains with secondary structure elements. HTH: helix-turn-helix, CTDH: C-terminal domain helix. Taken from (Yin et al., 2018).

1.1.3 Sub-families

The TRP family consists of seven subfamilies: TRPA (Ankyrin), TRPC (canonical), TRPM

(melastatin), TRPML (mucolipin), TRPP (polycystin), TRPN (no mechanoreceptor potential C) and TRPV (vanilloid) (Figure 1-2) (Nilius and Owsianik, 2011).The TRPN sub-family has only been identified in lower vertebrates and invertebrates. Currently, 28 members in six TRP subfamilies are expressed in mammals, 27 of which are expressed in humans.

Figure 1-2 Phylogenetic tree of TRP channels

A phylogenetic tree of TRP channels based on sequence homology. Taken from (Nilius and Owsianik, 2011)

1.1.4 Mammalian TRP channels

1.1.4.1 TRPA

The TRPA subfamily consists only of TRPA1 which was discovered as an ankyrin-like transmembrane protein with a similar structure to other TRP channels (Jaquemar et al., 1999).

TRPA1 was initially thought to be a putative noxious cold sensor (Story et al., 2003) although some studies have found no effect of cold on TRPA1 (Bautista et al., 2006; Jordt et al., 2004).

The channel is activated by allyl isothiocyanate (mustard oil), cinnamaldehyde (cinnamon) and allicin (garlic) and can also be activated by products of oxidative stress and environmental irritants such as acrolein (Andersson et al., 2008; Bautista et al., 2006). Hence TRPA1 may be better described as a chemo-nociceptor.

1.1.4.2 TRPC

The TRPC subfamily were the first mammalian TRP channels studied and TRPC1 was discovered and cloned in 1995 based on its sequence homology to the trp gene in Drosophila

(Wes et al., 1995). The TRPC subfamily consists of seven members which are divided into four groups based on sequence homology: TRPC1, TRPC2, TRPC3/TRPC6/TRPC7 and TRPC4/TRPC5.

TRPC channels can form both homotetramers and heterotetramers within the TRP superfamily including various TRPC channels (Kim et al., 2014; Storch et al., 2012; Strübing et al., 2001) and TRPP2 (Kobori et al., 2009). TRPC1 is broadly expressed in mammalian tissues and is implicated in immune regulation as TRPC1 deletion resulted in a reduced production of

T helper type 2 cytokines and chemokines in the lungs of ovalbumin-sensitised mice (Yildirim et al., 2012). There is a debate whether TRPC1 homotetramers can form functional ion channels as currents are difficult to detect in heterologous overexpression systems (Myeong et al., 2016). However, TRPC1 coexpression with TRPC4 or TRPC5 to form heterotetramers

18 changes the double-rectifying current/voltage (I/V) curves of TRPC4 and TRPC5 homotetramers to outward-rectifying I/V curves (Myeong et al., 2016). Therefore, TRPC1 expression is believed to exert its physiological function by altering the activity of other channels. TRPC2 is a pseudogene in humans but is expressed in the vomeronasal organ of mice where it responds to pheromones and therefore controls sexual behaviour (Stowers et al., 2002). TRPC3, TRPC6 and TRPC7 are expressed in cardiac and smooth muscle cells and can all be directly activated by lipids such as diacylglycerol (DAG) (B. Beck et al., 2006; Hofmann et al., 1999). TRPC4 and TRPC5 are broadly expressed. TRPC4 has been shown to be important for endothelial-dependent regulation of vascular tone and neurotransmitter release from the thalamic interneurons (Freichel et al., 2004). TRPC5 has been shown to be involved in the transduction of mechanical, osmotic and cold stimuli (Gomis et al., 2008; Zimmermann et al.,

2011).

1.1.4.3 TRPV

The founding member of the TRPV family, TRPV1, was named based on its activation by the vanilloid capsaicin (Caterina et al., 1997). Initial studies with capsaicin demonstrated a thermoregulatory role for TRPV1 in rats and guinea pigs (Jancsó-Gábor et al., 1970a, 1970b;

Szolcsányi et al., 1971). Application of capsaicin in rat isolated dorsal root ganglion (DRG) neurons induced fluxes of ions demonstrating that TRPV1 can be studied in sensory neurons

(Wood et al., 1988). Furthermore, protons were shown to activate a subpopulation of neurons that was also responsive to capsaicin demonstrating that TRPV1 is activated in acidic environments (Bevan and Geppetti, 1994). Capsazepine was the first discovered and characterised antagonist of TRPV1, and thereby of any TRP channel (Bevan et al., 1992). Later, anandamide was identified as an endogenous TRPV1 activator, which when released,

19 activates TRPV1 on perivascular sensory nerves causing the release of calcitonin-gene-related peptide (CGRP) leading to vasodilation (Zygmunt et al., 1999). TRPV1 was also shown to be activated by temperatures higher than 42 °C and is physiologically important for thermal and chemical nociception, regulation of body temperature, behavioural responses to noxious heat and heat hyperalgesia following inflammation (Caterina et al., 2000; Davis et al., 2000). TRPV2 is expressed in sensory neurons and was thought of as a noxious heat sensor that detects temperatures >52 °C; however, TRPV2 knock-out mice displayed normal thermal nociception

(Park et al., 2011). TRPV3 is expressed in various tissues including the oral cavity, the gastrointestinal tract and in keratinocytes of skin (Peier et al., 2002b). TRPV3 was thought to detect innocuous warm temperatures between 31 and 39 °C and TRPV3 knock-out mice exhibited strong deficits in responses to innocuous and noxious heat (Moqrich et al., 2005).

However, TRPV3 knock-out in mice with a more homogeneous genetic background displayed no altered thermosensation (Huang et al., 2011). The same study found no effect of TRPV4 knock-out on thermosensation in mice. TRPV4 has been implicated in osmoregulation where

Trpv4-/- mice drank less water and became more hyperosmolar than wildtype mice and is involved in mechanotransduction (Gao et al., 2003; Liedtke and Friedman, 2003; Wu et al.,

2007). Mutations in TRPV4 are associated with hereditary disorders such as metatropic dysplasia (Camacho et al., 2010, p. 4), a skeletal dysplasia, and Charcot-Marie-Tooth disease type 2C (Landouré et al., 2010), a neuromuscular disorder. TRPV5 and TRPV6 are highly selective to Ca2+ and contribute to calcium homeostasis. Both channels form homo- and heterotetramers (Hoenderop et al., 2003) with TRPV5 being exclusively expressed in the kidney and TRPV6 having a wider distribution in pancreatic, placental and intestinal tissues.

Both channels are constitutively active but are inactivated by high concentrations of calcium, therefore preventing calcium poisoning of the cells (Vennekens et al., 2000; Yue et al., 2001).

1.1.4.4 TRPML

The TRPML family consists of the Ca2+-permeable TRPML1, TRPML2 and TRPML3 which are mainly localised to intracellular compartments such as endosomes and lysosomes (Karacsonyi et al., 2007; Song et al., 2006; Venkatachalam et al., 2006). Loss of function mutations in

TRPML1 results in the neurodegenerative disorder mucolipidosis IV (Bargal et al., 2000; Bassi et al., 2000; Sun et al., 2000). All TRPML channels are activated by phosphatidylinositol 3, 5- bisphosphate (PI(3,5)P2) which is found in high levels in endolysosomes; however, TRPML1 and Drosophila TRPML are both inhibited by PIP2 which is mainly found in plasma membrane

(Feng et al., 2014; Xiaoli Zhang et al., 2012). These findings agree with the notion that TRPMLs are mainly active in endolysosomoes rather than in plasma membrane. Mammalian cells lacking TRPMLs exhibit features of diminished autophagic flux including accumulation of autophagic vacuoles, p62 and polyubiquitinated proteins (Curcio-Morelli et al., 2010;

Vergarajauregui et al., 2008; Zeevi et al., 2009). However, some studies have found that knockdown of TRPML1 in mammalian cells does not significantly delay the delivery of endocytosed material to lysosomes (Miedel et al., 2008).

1.1.4.5 TRPP

The TRPP family consists of TRPP 2,3 and 5. Mutations in TRPP2 leads to the development of autosomal dominant polycystic kidney disease (ADPKD) (Mochizuki et al., 1996). ADPKD is a common, monogenic and progressive disorder. The disease leads to the enlargement of kidneys and the formation of fluid-filled cysts progressively leading to renal failure. TRPP2 has been found to form complexes with polycystin1, a larger protein with 11 transmembrane domains, playing physiological roles such as tubular morphogenesis and maintaining left-right symmetry (Qian et al., 1997; Zhu et al., 2011).

1.1.5 TRPM sub-family

The mammalian TRPM subfamily consists of eight structurally and functionally diverse members proposed to contribute to the detection and transduction of heat, cold, osmolarity, taste, pH, redox state and control of Mg2+ levels homeostasis and cell proliferation or death (Zholos, 2010). TRPM channels can be phylogenetically subdivided into four groups: TRPM1/3, TRPM6/7, TRPM4/5 and

TRPM2/8 (Figure 1-3) (Harteneck, 2005).

Figure 1-3 Mammalian TRPM subfamily phylogenetic tree

A phylogenetic tree of the mammalian TRPM subfamily. The subfamily can be divided into four groups based on sequence homology: TRPM1/3, TRPM6/7, TROM4/5 and TRPM2/8. Adapted from (Venkatachalam and Montell,

2007)

1.1.5.1 TRPM1

The founding member of the TRPM sub-family, TRPM1 is expressed in skin melanocytes and eye. The

TRPM1 transcript is alternatively spliced to produce TRPM1-S and TRPM1-L in humans. Initially named melastatin, TRPM1-S was the first mammalian TRPM subfamily cloned (Duncan et al., 1998).

Expression of TRPM1-S was initially correlated with pigmentation in melanoma cell lines and inversely correlated with the metastatic potential of melanoma cells as a tumour suppressor (Duncan et al.,

1998; Fang and Setaluri, 2000). However, the presence of mouse Trpm1-L and -S transcripts in the retina were later discovered (Koike et al., 2010). TRPM1-S interacts directly with TRPM1-L to suppress

22 the latter’s activity by inhibiting its translocation to the plasma membrane in vitro (Xu et al., 2001).

Additionally, TRPM3 channels were found to biochemically interact with TRPM1 channels and form functionally heteromultimeric channels in a heterologous expression system (Lambert et al., 2011).

TRPM1 plays a crucial role in visual contrast recognition in vertebrate vision. When light is absorbed, rod and cone photoreceptor cells transmit signals through glutamate release to depolarising ON- bipolar cells and hyperpolarising OFF-bipolar cells (DeVries and Baylor, 1993; Dowling, 1978). TRPM1 is negatively regulated by the G-protein coupled receptor (GPCR) metabotropic glutamate receptor 6

(mGluR6) (Koike et al., 2010; Xu et al., 2016, p. 1). This regulation is by the direct interaction of liberated Gαo and Gβγ proteins with TRPM1. In the dark, a decrease in photon absorption by photoreceptor cells increases the concentration of glutamate released around the synaptic ribbon area of a photoreceptor cell. This leads to the activation of mGluR6 resulting in the release of Gαo and

Gβγ proteins. Both G-proteins inhibit TRPM1 leading to a decrease in cationic conductance

(hyperpolarisation, Figure 1-4). In contrast, an increase of photon absorption during light stimulation decreases glutamate release. This in turn inactivates mGluR6 leading to the opening of constitutively active TRPM1 channels and the depolarisation of ON-bipolar cells.

Figure 1-4 mGluR6 regulation of TRPM1 activity on ON-bipolar cells

In dark conditions (left panel), glutamate is released from the presynaptic terminal which binds to and activates postsynaptic mGluR6s. Subsequent Gαo activation leads to direct inhibition of TRPM1 channels by Gαo and Gβγ proteins hence hyperpolarising postsynaptic terminals. Conversely, photon absorption in light conditions (right panel) reduces glutamate release from the presynaptic terminal.

Constitutively active TRPM1 channels therefore conduct Ca2+ into the postsynaptic terminal causing depolarisation. Modified from Irie and Furukawa, 2014.

1.1.5.2 TRPM2

TRPM2 mRNA is ubiquitously expressed in all tissues except for bone and cartilage therefore implicating TRPM2 in a variety of physiological processes. Point mutations in TRPM2 may

24 confer susceptibility to Western Pacific amyotrophic lateral sclerosis and parkinson’s disease

(Hermosura et al., 2008) A Nudix hydrolase domain in the carboxy-terminal tail of TRPM2 functions as an ADP-ribose pyrophosphatase where cyclic ADPR and nicotinamide-adenine dinucleotide (NAD) are thought to activate the channel (Perraud et al., 2001). TRPM2 activation promotes inflammation and immune responses though the production of cytokines

CXCL8, interleukin-6 and 10 and TNF-α (Wehrhahn et al., 2010; Yamamoto et al., 2008).

TRPM2 also plays a role in insulin secretion from pancreatic β-cells (Inamura et al., 2003;

Togashi et al., 2006; Uchida et al., 2011). TRPM2 has recently been discovered to be involved in phagosome maturation where it aids in bacterial clearance hence reducing mortality in a mouse model of E. coli sepsis (Zhang et al., 2017). Paradoxically, TRPM2 reduces the inflammatory response via cellular depolarisation and subsequent reduction of reactive oxygen species (ROS) production in phagocytes (Di et al., 2011). TRPM2 is also highly expressed in the central nervous system (CNS) where it has been found in dorsal root ganglion

(DRG) sensory neurons (Nazıroğlu et al., 2011), microglia (Fonfria et al., 2006; Kraft et al.,

2004) and in hippocampal neurons (Olah et al., 2009). TRPM2 has a physiological role in temperature sensation and thermoregulation which is consistent with it being activated by warm temperatures (Song et al., 2016; Togashi et al., 2006). A recent study demonstrated that TRPM2 channels expressed in the preoptic area of the hypothalamus play a role in sensing hyperthermia and are important for temperature homeostasis (Song et al., 2016).

Additionally, TRPM2 is involved in heat sensation in somatosensory and autonomic neurons

(Tan and McNaughton, 2016).

1.1.5.3 TRPM3

TRPM3 is unique to other TRP channels as the Trpm3 gene has several alternative splice sites producing over 20 different isoforms (Oberwinkler and Philipp, 2014). Due to the high similarity between these variants, it is difficult to produce specific primers and antibodies

(Held et al., 2015b). Therefore, not all the isoforms have been identified in cellular tissues, making their physiological relevance and function unknown. TRPM3 is expressed in a wide variety of tissues, such as in the brain including the cerebellum and posterior hypothalamus, spinal cord, sensory neurons, retina, testis, pituitary, kidney and adipose tissue (Lee et al.,

2003). Furthermore, it is expressed in bladder, pancreas, heart, ovaries and sperm cells

(Grimm et al., 2003). TRPM3 mRNA is also found in mouse dorsal root ganglion (DRG) and trigeminal ganglion (TG) neurons in a similar number of cells as TRPV1 (Vandewauw et al.,

2013; Vriens et al., 2011). Early studies indicate that TRPM3 may play a role in heat sensing as TRPM3-null mice displayed reduced heat nociception and demonstrated reduced sensation of elevated temperatures in various behavioural tests such as hot plate and thermal preference tests (Vriens et al., 2011). Furthermore, Trpm3-/- mice did not demonstrate any major deficits apart from altered behavioural thermosensation. A more recent study demonstrated that acute noxious heat sensing in mice relies on TRPM3, TRPV1 and TRPA1

(Vandewauw et al., 2018). Robust somatosensory heat responsiveness at the cellular and behavioural level is maintained if at least one of these three TRP channels is functional

(Vandewauw et al., 2018). Pharmacological inhibition and genetic KO of all three channels strongly prevents heat responses in both rapidly firing C and Aδ fibres that innervate the skin and isolated sensory neurons (Vandewauw et al., 2018). Triple KO mice lacked the acute withdrawal response to noxious heat required to avoid burn injury but demonstrated normal

26 nociceptive responses to cold or mechanical stimuli and a preference for moderate temperatures (Vandewauw et al., 2018).

1.1.5.4 TRPM4

In contrast to most functionally characterised TRP channels, TRPM4 is permeable only to monovalent cations (Fleig and Penner, 2004; Launay et al., 2002; Mathar et al., 2014; Song and Yuan, 2010). The channel is activated by intracellular Ca2+ although in a rapidly desensitising manner and phosphatidylinositol 4,5-biphosphate (PIP2) resensitises TRPM4

(Nilius et al., 2006, p. 4; Zhang et al., 2005). TRPM4 is expressed in a variety of tissues and is involved in several physiological processes including inflammation by mediating axonal and neuronal degeneration in experimental autoimmune encephalomyelitis and multiple sclerosis

(Schattling et al., 2012) and insulin secretion (Cheng et al., 2007). Recently, TRPM4 has been shown to be involved in the transduction of bitter, sweet and umami stimuli implicating it in the transduction of taste (Dutta Banik et al., 2018).

1.1.5.5 TRPM5

TRPM5, like TRPM4, is permeable only to monovalent cations. It is also activated by elevated intracellular Ca2+ levels and is voltage-sensitive (Hofmann et al., 2003; Liu and Liman, 2003;

Prawitt et al., 2003). TRPM5’s physiological significance was realised with the discovery of its expression being largely restricted to taste receptor cells (Pérez et al., 2002; Zhang et al.,

2003). The three modalities of bitter, sweet and umami are mediated by GPCRs that bind their respective tastant (Lindemann, 2001; Margolskee, 2002). These GPCRs activate the G protein gustducin and PLCβ2 consequently leading to an intracellular signalling cascade that results with an electrical response that is likely carried out by TRPM5. This is evidenced by TRPM5 being coexpressed with GPCRs responsible for bitter, sweet and umami tasting and with

27 gustducin and PLCβ2 (Pérez et al., 2002; Zhang et al., 2003). However, TRPM5 knockout (KO) mice demonstrated reduced, but not abolished, bitter, sweet and umami taste sensation

(Damak et al., 2006; Eddy et al., 2012; Ohkuri et al., 2009) suggesting that a TRPM5- independent mechanism is required for normal taste transduction. KO of both TRPM4 and

TRPM5 abolished the animal’s ability to detect these three modalities(Dutta Banik et al.,

2018).

1.1.5.6 TRPM6

TRPM6 is largely expressed in kidney and intestinal cells. TRPM6 along with TRPM7 are the only ion channels, of more than 300, that contain a fully functional C-terminal kinase. This kinase is part of the atypical protein kinase (APK) subfamily, also known as the α-kinases which share little sequence similarity to conventional protein kinases (Duan et al., 2018; Middelbeek et al., 2010; Yamaguchi et al., 2001). TRPM6 plays a critical role for systemic Mg2+ homeostasis and exists as a homodimer but can also heterodimerise with TRPM7. The naturally occurring

S141L TRPM6 missense mutation prevents TRPM6 from heterodimerising with TRPM7, disrupting epithelial magnesium absorption and causing the autosomal recessive disorder hypomagnesemia with secondary hypocalcemia in patients (Chubanov et al., 2004).

1.1.5.7 TRPM7

TRPM7 plays an indispensable role in cellular biology as it regulates homeostasis of divalent cations Mg2+, Ca2+ and Zn2+ (Zou et al., 2019). An example of its critical role is that TRPM7 is believed to be the most important cation channel involved in controlling intracellular Mg2+ levels in vascular cells. The channel is modulated by vasoactive factors including AngII, endothelin-1, aldosterone and bradykinin (Callera et al., 2009; Touyz et al., 2006; Valinsky et al., 2016; Yogi et al., 2011). Additionally, TRPM7 is crucial for embryogenesis and

28 development where global knockout of TRPM7 in mice results in embryonic lethality prior to embryonic day 7 (Jin et al., 2012).

1.1.5.8 TRPM8

TRPM8 was originally found in the testis and prostate tissue and was shown to be upregulated in prostate tumours (Tsavaler et al., 2001). This upregulation has now been identified in several other tumours (Yee et al., 2010). TRPM8 was soon found to be expressed in a subpopulation of small-diameter, cold-sensitive peripheral sensory neurons (McKemy et al.,

2002; Peier et al., 2002a). The channel plays a crucial role in the detection of cold temperatures (Bautista et al., 2007; Colburn et al., 2007; Dhaka et al., 2007). More recently,

TRPM8 has been found to act as a neuronal osmosensor that regulates eye blinking in mice

(Quallo et al., 2015).

1.2 Introduction to G-protein coupled receptors

GPCRs are the largest class of membrane proteins in the human genome and mediate most cellular responses to a wide variety of external stimuli. Around 800 GPCRs have been identified in humans, roughly half of which have sensory functions mediating olfaction

(∼400), taste (33), light perception (10) and pheromone signalling (5) (Mombaerts, 2004).

Approximately 350 non-sensory GPCRs mediate signalling by ligands ranging from small molecules, to peptides to large proteins. A high proportion of drugs in clinical usage target

GPCRs (Overington et al., 2006; Rask-Andersen et al., 2014).

1.2.1 Discovery

The pioneering work of Earl Sutherland in the 1950’s and 1960’s was one of the drivers that ultimately led to the unravelling of the molecular structure and mechanism of action of

GPCRs. While examining the mechanism of phosphorylase activation by glucagon and adrenaline, Sutherland established the first broken cell preparations in which he could observe hormonal effects. This experimental setup led him to discover cyclic AMP (cAMP) and the enzyme responsible for its formation, adenylate cyclase (AC) (Hardman et al., 1971;

Lefkowitz, 2000). Martin Rodbell and Lutz Birnbaumer would later observe in fat cells and liver membranes that both systems contained distinct types of receptors there were coupled to a common catalytic cyclase (Birnbaumer and Rodbell, 1969). They proposed an intermediate protein acts as a transducer that couples the receptors to the catalyst (Rodbell et al., 1971). Elliott Ross and Alfred Gilman confirmed that the transducer is the heterotrimeric G protein Gαs which couples the receptors to cyclase activation (Ross and

Gilman, 1977). A family of G proteins that all utilised GTP hydrolysis to couple receptors were then discovered (Cassel and Selinger, 1976). These G proteins were responsible for stimulating or inhibiting many effectors including adenylate cycle, PLC and ion channels

(Gilman, 1987). Gαs was first isolated by Gilman in 1980 (Northup et al., 1980) followed by the isolation of Gαi which functioned as an inhibitor of adenylate cyclase (Bokoch et al., 1984;

Codina et al., 1983). The development of radioligand-binding studies in the 1970’s permitted direct study of membrane receptors. Once a ligand was available it allowed for the purification of a receptor (Lefkowitz, 2000). The β2-adrenergic receptor was initially purified from amphibian erythrocytes (Shorr et al., 1981) followed by the β1-adrenergic receptor, which was purified from avian erythrocytes (Shorr et al., 1982). These discoveries subsequently allowed the clear physical separation of the receptors from adenylate cyclase

(Haga et al., 1977; Limbird and Lefkowitz, 1977) and helped demonstrate that the receptors could be found in agonist-driven complexes with a G protein (Limbird and Lefkowitz, 1978).

Cloning of the β2-adrenergic receptor and rhodopsin permitted the realisation that these two

30 receptors, with very different functions, share sequence homology and an alleged seven membrane-spanning topography and could possibly belong to the same gene family (Dixon et al., 1986). Although this realisation was initially startling, subsequent cloning of other

GPCRs confirmed the existence of this large family (Lefkowitz, 2000).

1.2.2 Families

GPCRs are classified by dividing them into six different classes based on sequence homology

(Kolakowski, 1994). These classes are: Class A (rhodopsin-like), Class B (secretin receptor family), Class C (metabotropic glutamate), Class D (fungal mating pheromone receptors), Class

E (cyclic AMP receptors) and Class F (frizzled/smoothened). Classes D and E are not found in vertebrates. Alternatively, GPCRs are classified using the GRAFS system, which consists of five main classes that overlap with classes A-F: Glutamate (G), Rhodopsin (R), Adhesion (A),

Frizzled/Taste2 (F) and Secretin (S) (Schiöth and Fredriksson, 2005).

1.2.2.1 Glutamate family (class C)

In humans, this family consists of the eight metabotropic glutamate receptors (mGluR), two

γ aminobutyric acid (GABA) receptors that exist in two splice variants A and B, taste receptors type 1 and a calcium sensing receptor (IUPHAR).

1.2.2.2 Rhodopsin family (class A)

The rhodopsin family is the largest with approximately 400 members and includes receptors for a wide variety of neurotransmitters, peptides, hormones and small molecules. Members of this family include opioid receptors (µ, δ, κ and nociception), neuropeptide Y, purinergic

2Y, bradykinin, prostaglandin E2, muscarinic acetylcholine, adenosine and protease-activated receptors (IUPHAR).

1.2.2.3 Adhesion family

The adhesion family consists of 33 members in humans with a broad distribution in embryonic, neuronal, reproductive tract and tumour cells (Schiöth and Fredriksson, 2005).

Adhesion GPCRs are unique in that they have an extended extracellular N-terminal. Most adhesion GPCRs possess a GPCR proteolytic site (GPS) motif in the N-terminal which can undergo autoproteolysis. The physiological role of this process is unknown; however, mutations in this domain in GPR56 can cause receptor misfolding and bilateral frontoparietal polymicrogyria, a congenital brain malformation resulting in irregularities on the surface of the cortex (Chiang et al., 2011; Jin et al., 2007; Ke et al., 2008; Piao et al., 2004). Another of example adhesion GPCRs is the latrophilin-1, a key target of α-latrotoxin, which is found in venom of the black widow spider (Krasnoperov et al., 1997; Lelianova et al., 1997).

1.2.2.4 Frizzled family (class F)

The frizzled family consists of 10 frizzled and smoothened proteins (IUPHAR). The frizzled receptors are activated by secreted lipoglycoproteins of the Wnt family, whilst the smoothened receptors are indirectly activated by the Hedgehog family of proteins acting on the transmembrane protein patched.

1.2.2.5 Secretin family (class B)

The secretin family consists of 15 members that are activated by polypeptide hormones and neuropeptides such as secretin, calcitonin gene-related peptide (CGRP), vasoactive intestinal peptide (VIP), glucagon, and glucagon-like peptide 1 (Harmar, 2001). These peptides are of clinical relevance in diabetes (glucagon and glucagon-like peptide 1), migraine and neuroinflammation (CGRP) and inflammation (VIP) (Archbold et al., 2011).

1.2.3 GPCR signalling dynamics

Classical signal transduction through GPCRs relies on receptor-mediated activation of heterotrimeric G proteins that consist of Gα, Gβ and Gγ subunits. GPCRs signal through the four major G-protein families Gs, Gi/o, Gq/11 and G12/13 (Downes and Gautam, 1999; Hilger et al., 2018; Simon et al., 1991). In the inactive GDP-bound state, Gα subunits associate with Gβγ dimers to form an inactive heterotrimer. Activation of GPCRs then promotes GDP dissociation from Gα, which is considered to be the rate-limiting step in G-protein activation (Higashijima et al., 1987). Free GTP then binds to the nucleotide-binding site of Gα promoting Gβγ dissociation. Gα proteins interact directly with effector molecules such as PLC, adenylyl cyclase and ion channels (Kristiansen, 2004; Milligan and Kostenis, 2006; Xu et al., 2016; Zhang et al., 2012). Gβγ can also target some of the same and additional effectors as Gα along with recruiting G-protein receptor kinases (GRKs) to the membrane to regulate GPCR activity (Khan et al., 2013; Smrcka, 2008; Xu et al., 2016). GPCR signalling is terminated when the intrinsic

GTPase activity of Gα subunits hydrolyses GTP to GDP, followed by the re-association of Gα with Gβγ dimers. This classical two-state model of activation proposes that GPCRs exist in an equilibrium between inactive and active states (Gether, 2000; Makita and Iiri, 2014; Sato et al., 2016). This explains why GPCRs are sometimes constitutively active in the absence of agonists. Therefore, agonists act by shifting the equilibrium to the active state, inverse agonists produce the opposite effect and neutral antagonists only compete with agonists without influencing the equilibrium or constitutive activity of GPCRs (Figure 1-5) (Sato et al.,

2016). Furthermore, agonists with different efficacies can produce different dynamics in the same GPCR-G-protein complex. Full agonists are able to stabilise complexes with lower

33 affinities for GDP in comparison to partial agonists (Gregorio et al., 2017; Roberts et al., 2004;

Seifert et al., 1999; Selley et al., 1997; Zhang et al., 2004).

Figure 1-5 Differing levels of GPCR activity

A simple schematic demonstrating the different levels of GPCR activity. GPCRs can exist in a constitutively active state with minimal activity levels. Neutral antagonists have no effect on constitutive activity and only prevent binding of agonists. Full and partial agonists can fully and partially activate GPCRs whereas inverse agonists decrease the activity of GPCRs.

1.2.3.1 Gα- and Gβγ-mediated signalling

The four subclasses of Gα proteins (Gs, Gi/o, Gq/11 and G12/13) target specific signalling cascades

(Wettschureck and Offermanns, 2005). Gαs, Gαi and Gαo all target adenylyl cyclase where Gαs stimulates and Gαi/o inhibits it. Stimulation of adenylyl cyclase convert ATP into cAMP.

Increased levels of cAMP leads to the activation of cAMP-regulated proteins including protein kinase A (PKA) and cyclic nucleotide-gated channels. Gαq/11 activates PLCβ isoforms which in turn hydrolysis PIP2 into inositol triphosphate (IP3) and membrane-bound diacylglycerol

(DAG). IP3 opens IP3 receptors on the endoplasmic reticulum (ER) membrane leading to

2+ Ca release whilst DAG activates protein kinase C (PKC). Gα12/13 activates Rho guanine- nucleotide-exchange factors (GEFs) which subsequently activate Rho. Gβγ regularly interacts with the same targets that are engaged by Gα subunits (Figure 1-6) (Khan et al., 2013).

1.2.3.2 GPCR kinases and arrestins

Rhodopsin phosphorylation following receptor activation was observed before the realisation that a GPCR family existed (Kühn, 1974; Kühn and Dreyer, 1972). Later studies discovered that rhodopsin kinase binds rhodopsin-containing membranes only after receptor activation by light (Kühn, 1978). Following these discoveries, another receptor kinase, β-adrenergic receptor kinase, was identified to specifically phosphorylate activated

β2-adrenergic receptors (Benovic et al., 1986b). The same kinase was also observed to phosphorylate rhodopsin following receptor activation (Benovic et al., 1986a). This kinase and the rhodopsin kinase were then cloned and sequenced and are now known as GPCR kinase 2 (GRK2) and GRK1. This indicates that a larger family of GRKs exists which currently consists of seven members (Benovic et al., 1989; Gurevich et al., 2012; Gurevich and

Gurevich, 2019).

Members of the GRK family are recruited to active GPCRs through various means. GRK1 and

7 are prenylated at their C-terminal and are therefore constitutively localised to the membrane (Gurevich and Gurevich, 2019). The pleckstrin homology (PH) domain of GRK2 and GRK3 binds Gβγ (Koch et al., 1993; Lodowski et al., 2003; Touhara et al., 1994).

Following Gβγ binding, GRK2 and GRK3 are recruited to the membrane where they phosphorylate GPCRs and stop receptor signalling (Haga and Haga, 1992; Li et al., 2003;

Pitcher et al., 1992). GRK4, 5 and 6 lack C-terminal prenylation and the PH domain and

35 instead associate with the plasma membrane through palmitoylation of their C-terminal cysteines and through an amphipathic helix interacting with the membrane phospholipids

(See Gurevich et al., 2012 for review). Phosphorylation of rhodopsin (Arshavsky et al., 1985) and β2-adrenergic receptors (Benovic et al., 1989; Sibley et al., 1986) by GRKs partially but not fully reduced G-protein signalling, suggesting that other factors were involved.

Binding of arrestins to active and phosphorylated GPCRs was demonstrated with visual arrestin-1 and non-visual arrestin-2 (Krasel et al., 2005; Wilden et al., 1986). Visual arrestin-

1 prevented coupling of phosphorylated rhodopsin to its cognate G protein transducin

(Wilden et al., 1986). It does so by competing with transducin for the activated and phosphorylated rhodopsin (Krupnick et al., 1997; Wilden, 1995). The addition of purified visual arrestin-1 significantly increased the desensitising effect of β2-adrenergic receptor phosphorylation by GRK2, suggesting that a non-visual homolog of arrestin-1 may be required for homologous desensitisation of non-rhodopsin GPCRs (Benovic et al., 1987). The first non-visual arrestin was cloned (Lohse et al., 1990) and termed β-arrestin for its preference to bind to β2-adrenergic receptors over rhodopsin (Lohse et al., 1992). Cloning of a second non-visual arrestin followed, which was termed β-arrestin 2 (Attramadal et al.,

1992). Later these arrestins were renamed in order of cloning with visual arrestin-1, β- arrestin 1 and 2 being termed arrestin 1, 2 and 3 (Sterne-Marr et al., 1993). Thereafter only one more arrestin, cone photoreceptor-specific arrestin-4, was identified in mammals (Craft et al., 1994; Murakami et al., 1993). Therefore hundreds of GPCRs expressed in mammals are targeted by four arrestins, where arrestin-1 and -4 are expressed in photoreceptor cells in the retina and bind photopigments, and arrestin-2 and -3 serve all other GPCRs (Gurevich and Gurevich, 2019).

Arrestins and G proteins compete to bind to the same inter-helical cavity on the cytoplasmic side of the GPCR (Carpenter et al., 2016; Kang et al., 2015; Liang et al., 2017; Rasmussen et al., 2011; Zhou et al., 2017). Unlike bound G proteins, which dissociate from GPCRs when

GTP is present, arrestins remain attached. Therefore, arrestins easily compete with G proteins and bind to active and phosphorylated GPCRs (Gurevich and Gurevich, 2004). In addition to binding to the inter-helical cavity, arrestins also tightly bind receptor-attached phosphatases (Zhou et al., 2017). Arrestins facilitate GPCR internalisation through coated pits. Arrestin-2 and -3 bind clathrin (Goodman et al., 1996) and its adapter AP2 (Laporte et al., 1999) through sites in their C-termini (Kim and Benovic, 2002) that are more accessible following GPCR binding (Zhou et al., 2017), similar to what occurs when arrestin-1 binds

GPCRs (Gurevich et al., 1994; Hanson et al., 2006; Vishnivetskiy et al., 2010, 2002).

As well as arresting GPCR signalling, arrestins serve as signal transducers themselves (Figure

1-6). Receptor-bound arrestins were found to promote Src-dependent activation of pro- proliferative mitogen activated protein kinases (MAPKs) ERK1 and 2 (Luttrell et al., 1999).

Arrestin-3 was then discovered to scaffold apoptosis signal-regulating kinase 1-MAP kinase kinases (MAP2Ks) 4/7-c-Jun N-terminal kinase 3 cascade following GPCR activation

(McDonald et al., 2000). Arrestins are able to scaffold several other three-tiered MAPK cascades (Luttrell et al., 2001; Peterson and Luttrell, 2017). MAPK cascades are conserved three-tier signalling modules which consists of upstream MAP kinase kinase kinases

(MAP3Ks), intermediate MAP2Ks and downstream MAPKs (Gurevich and Gurevich, 2019).

MAP3Ks and MAP2Ks activate their downstream target kinases by phosphorylating their activation loops (Tian and Harding, 2014). MAPKs phosphorylate a plethora of nuclear and cytoplasmic proteins to produce cellular responses. Arrestins, acting as scaffolding proteins, bring MAP kinases closer to each other and thereby facilitates signal transduction. Crucially,

37 signalling only occurs when upstream MAP3Ks are activated and arrestins have so far not been implicated in this step. Therefore for arrestins to facilitate signal transduction, MAP3Ks must be activated by other factors such as integrins (Stupack and Cheresh, 2002) and growth factor receptors (Garrington and Johnson, 1999).

Figure 1-6 Canonical G-protein and arrestin signalling pathways

A simplified schematic demonstrating canonical G-protein (left of dotted line) and arrestin (right of dotted line) signalling pathways. Gαi/o inhibits whereas Gαs stimulates adenylyl cyclase (AC) which then converts ATP into cAMP subsequently activating PKA. Gαq/11 activates PLC which converts PIP2 into IP3 and DAG. IP3 activates IP3 receptors in the endoplasmic reticulum and leads to the efflux of Ca2+ from Ca2+ stores. DAG activates PKC.

Gα12/13 activation activates guanosine exchange factors which then activates Rho kinase. Gβγ can activate all these signalling pathways. GRKs bind to and phosphorylate active GPCRs leading to receptor desensitisation.

Arrestins then bind phosphorylated GPCRs and become activated leading to several signalling events including

38 activation of SRC and MAP kinases. Arrestin binding also acts to stop GPCR-mediated signalling by further desensitising the receptor and internalising it, which results in receptor recycling or degradation.

1.3 GPCR regulation of ion channels

GPCR activity regulates a variety of ion channels whether by direct G protein interactions and through G-protein-dependent secondary signalling pathways or by non-G-protein-dependant signalling (Figure 1-7). G protein-inwardly rectifying potassium channels (GIRKs), voltage- gated calcium channels (VGCCs), KV7 and TRP channels are all examples of ion channels that are regulated by GPCR activity (Salzer et al., 2019).

Figure 1-7 Mechanisms of GPCR regulation of ion channels

Schematic of GPCR-mediated regulation of ion channels. A G-proteins have been shown to directly bind and regulate ion channels. G-proteins can also indirectly regulate ion channels through downstream signalling

2+ cascades. PIP2 hydrolysis can regulate ion channels directly or via IP3 and DAG. IP3 receptor (IP3R) -mediated Ca release from the endoplasmic reticulum (ER) causes Ca2+ ions to bind and activate CaM which in turn binds and regulates ion channels. DAG-mediated activation of PKC also modulates ion channels. cAMP can directly regulate

40 ion channels or regulate them through activating PKA. B GPCRs also regulate ion channels through G-protein independent arrestin signalling. Arrestin activation can regulate ion channels by scaffolding proteins to the plasma membrane and by recruiting or internalising them.

1.3.1 GPCR regulation of GIRK channels

GIRK channels are homo- or heterotetrameric channels formed of four different subunits (Kir

3.1-3.4). GIRKs are promiscuously activated by Gi/o-, Gq- and Gs-coupled GPCRs through liberated Gβγ subunits (Dascal, 1997; Dolphin, 2003; Yamada et al., 1998) although Gi/o- coupled GPCRs predominantly activate GIRKs (Touhara and MacKinnon, 2018). This is observed in sinoatrial node pacemaker cells where GIRK channels are activated by stimulated

Gi/o-coupled muscarinic acetylcholine receptor M2 (M2Rs) following release of acetylcholine from the vagus nerve (Krapivinsky et al., 1995; Logothetis et al., 1987; Sakmann et al., 1983;

Soejima and Noma, 1984; Wickman et al., 1994). Open GIRK channels hyperpolarise the cell membrane by outwardly conducting K+ ions thereby lengthening the interval between cardiac action potentials. On the other hand, stimulation of Gs-coupled β-adrenergic receptors increases heart rate partly by PKA-mediated stimulation of L-type Ca2+ channels (Bean et al.,

1984; DiFrancesco and Tortora, 1991; Yue et al., 1990) and ryanodine receptors (Ullrich et al.,

2012; Valdivia et al., 1995; Xiao et al., 2005). Activation of β-adrenergic receptors does not open GIRK channels even though Gβγ is also liberated by this receptor (Digby et al., 2008;

Hein et al., 2006). Later biochemical studies demonstrated that activation of M2Rs catalyse

Gβγ subunit release at higher rates than β2-adrenergic receptors do, resulting in higher Gβγ concentrations that activate GIRK channels (Touhara and MacKinnon, 2018). This increased rate of Gβγ release following M2R activation leads to a faster G-protein association rate with

GIRK which accounts for the specificity of M2Rs with GIRK (Touhara and MacKinnon, 2018).

Additionally, PIP2 is necessary for GIRK function and its depletion prevents GIRK activation by

Gβγ subunits indicating it as a cofactor for GPCR regulation of GIRK channels. (Huang et al.,

1998; Sui et al., 1998). GIRK channel activation by Gβγ or ethanol increases the channel’s affinity for PIP2 resulting in an increase in the channel open time (Bodhinathan and Slesinger,

2013; Huang et al., 1998; Petit-Jacques et al., 1999).

1.3.2 GPCR regulation of VGCCs

VGCCs mediate Ca2+ influx in response to membrane depolarisation and regulate many physiological processes including muscle contraction, gene expression and neurotransmission

(Catterall et al., 2005). N, P/Q and R-type VGCCs are all directly inhibited by binding of Gβγ subunits (Hille, 1994; Ikeda, 1996). VGCCs are promiscuously inhibited by Gi/o-, Gq- and Gs- coupled GPCRs (Ewald et al., 1989; Golard et al., 1994; Shapiro and Hille, 1993; Zhu and Ikeda,

1994). Like GIRK channels, VGCCs are predominantly modulated by Gαi/o-coupled GPCRs (Holz et al., 1986; McFadzean et al., 1989; Scott and Dolphin, 1986). Furthermore, N-type VGCCs are inhibited by Gq/11-coupled muscarinic M1 receptor (M1R) activation by PIP2 depletion

(Gamper et al., 2004). The N-type CaV2.2 channel is inhibited following GPCR stimulation through two pathways in sympathetic neurons (Hille, 1994). The quicker pathway is voltage dependent, insensitive to intracellular Ca2+ chelation and membrane delimited. It is initiated following the activation of Gi/o-coupled GPCRs, can be relieved by applying large positive pulses (Bean, 1989; Lipscombe et al., 1989; Zamponi and Snutch, 1998) and is described as direct voltage-dependent binding of Gβγ protein to Cav2.2 and P/Q-type Cav2.1 VGCCs

(Herlitze et al., 1996; Ikeda, 1996). The slower pathway is voltage independent, sensitive to

2+ Ca chelation and insensitive to PTX and therefore not mediated by Gi/o-coupled receptors.

This pathway is initiated following stimulation of Gq-coupled receptors (Bannister et al., 2002;

Bernheim et al., 1991; Mathie et al., 1992; Shapiro et al., 2000). PIP2 depletion inhibits both

N-type and L-type VGCC activity through voltage-independent pathways and promotes the susceptibility of the channels to voltage-dependent inactivation where the resynthesis of PIP2 from PIP is required for VGCC channel recovery (Suh et al., 2010).

1.3.3 GPCR regulation of Kv7 channels

Currents passing through KV7 channels are termed M-currents as they were originally described as currents that are inhibited by activating muscarinic acetylcholine receptors

(Brown and Adams, 1980). Activation of Gq-coupled GPCRs inhibits KV7 channels through depletion of membrane PIP2 (Suh and Hille, 2002). Additionally, KV7 channels are inhibited by

2+ IP3-mediated increases in intracellular Ca levels and subsequent calmodulin binding

(Gamper and Shapiro, 2003; Selyanko and Brown, 1996).

1.3.4 GPCR regulation of TRP channels

1.3.4.1 TRPV1-4

TRPV1 channels are sensitised by Gq- and Gs-coupled receptors where inflammatory mediators such as bradykinin and prostaglandin are well known to sensitise TRPV1 in neurons

(Cesare and McNaughton, 1996; Reichling and Levine, 1997). Every step in the Gq-mediated pathway can affect TRPV1 channel function (Rohacs, 2013). PIP2 in the plasma membrane decreases TRPV1 activity by stabilising the channel in its resting state and interfering with agonist binding but at the same time, PIP2 may function as a positive co-factor that when bound to TRPV1 in the closed state primes the channel for activation (Gao et al., 2016). PKC

43 activation sensitised TRPV1 by phosphorylating two serine residues on the C-terminus (Julius,

2013). Finally, a rise in cytosolic Ca2+ leads to rapid desensitisation of TRPV1 in response to prolonged activation (Rohacs, 2013). This desensitisation is due to calmodulin binding to the

N-terminal ankyrin repeat domain which is conserved in TRPV3 and TRPV4 (Phelps et al.,

2010). PKA-mediated phosphorylation of TRPV1 following Gs-coupled GPCR activation sensitises TRPV1 to its agonists and reduces Ca2+-mediated desensitisation (Bao et al., 2015).

Activation of various GPCRs including the Gq-coupled bradykinin B2 (BK2) and Gs coupled prostaglandin E2 EP2 receptors by inflammatory modulators have been shown to sensitise

TRPV1 (Basbaum et al., 2009; Cesare and McNaughton, 1996; Lopshire and Nicol, 1998;

Schnizler et al., 2008; Tang et al., 2004). Conversely, activation of Gi/o-coupled µ-opioid receptors reduces PKA activity and subsequently decreases TRPV1 channel activity (Endres-

Becker et al., 2007; Vetter et al., 2006). Arrestin signalling also affects TRPV1 activity where

β-arrestin 2 has been shown to be involved in TRPV1 desensitisation (Por et al., 2012). β- arrestin 2 scaffolding of phosphodiesterase PDE4D5 to the plasma membrane led to TRPV1 desensitisation and siRNA-mediated knockdown of β-arrestin 2 in primary cultures produced significantly greater initial and repeated capsaicin-evoked responses (Por et al., 2012).

Moreover, genetic KO of β-arrestin-2 in mice led to significantly reduced desensitisation of

TRPV1 in primary cultures (Por et al., 2012).

2+ TRPV2 Ca -dependent desensitisation has been linked to decreases in membrane PIP2 levels

(Mercado et al., 2010) whereas TRPV3 has been shown to be potentiated by PIP2 hydrolysis following activation of muscarinic M1 acetylcholine receptors (M1R) (Doerner et al., 2011).

TRPV4, like TRPV1, is phosphorylated and sensitised by PKA and PKC (Peng et al., 2010; Xu et al., 2003). TRPV4 is also negatively modulated by arrestins, specifically β-arrestin 1 (Shukla et al., 2010). Angiotensin receptor (AT1R) and TRPV4 form a complex in the plasma membrane

(Shukla et al., 2010). AT1R stimulation recruits β-arrestin 1 to the complex leading to the ubiquitination of TRPV4 and subsequent receptor internalisation and functional downregulation of the channel (Shukla et al., 2010).

1.3.4.2 TRPA1

TRPA1 channels are sensitised by Gq- and Gs- coupled GPCRs. Activation of Gq-coupled protease activated receptor 2 (PAR2) receptors increased TRPA1 ionic currents in DRGs neurons through PIP2 hydrolysis (Dai, 2016; Dai et al., 2007). PKC-mediated phosphorylation was also shown to sensitise TRPA1 following bradykinin B1 (BK1) receptor activation (Meotti et al., 2017). Activation of the Gs-coupled adenosine 2 A (A2A) receptors sensitised TRPA1 currents in oesophageal C-fibres through PKA activation (Brozmanova et al., 2016).

1.3.4.3 TRPC1, 3, 4 and 5

TRPC channels are activated by Gq-coupled GPCRs where the exact mechanism remains unknown (Schaefer et al., 2000). Several mechanisms have been proposed including sensitisation by increases in intracellular Ca2+ levels (Blair et al., 2009; Gross et al., 2009) and activation of TRPC4 (Otsuguro et al., 2008) and TRPC5 (Trebak et al., 2009) following PIP2 hydrolysis. Paradoxically, TRPC5 can also be inhibited by PIP2 (Kim et al., 2008; Trebak et al.,

2009). Additionally, activation of PKA following Gαs activation inhibited both channels (Sung et al., 2011). Both TRPC4 and TRPC5 have been shown to be directly activated by Gαi subunits

(Jeon et al., 2012, 2008). Recently, Gαq proteins have been shown to directly interact and activate TRPC4 and TRPC5 whilst simultaneously inhibiting TRPC1/4 and TRPC4/5 heterotetramers through PLC-mediated PIP2 hydrolysis (Myeong et al., 2018). Arrestin signalling has been shown to recruit TRPC3 to the plasma membrane (Liu et al., 2017).

Activation of AT1R with the β-arrestin 1 biased agonist, TRV120027, promoted the recruitment of TRPC3 or PLCγ to the AT1R-β-arrestin 1 signalling complex (Liu et al., 2017).

TRPC3 activation was prevented when the C-terminal of β-arrestin 1 was replaced with that of β-arrestin 2’s or by using a TAT-p1 peptide that blocks the interaction between β-arrestin

1 and PLCγ (Liu et al., 2017). Therefore, arrestin signalling is not only limited to receptor desensitisation and internalisation.

1.4 Aims of this thesis

The aim of this thesis is to characterise TRPM3 activity so that we could determine how to accurately examine its role in somatosensation. Many ion channels including TRP channels are regulated by GPCRs and therefore the second aim of this thesis is to explore GPCR modulation of TRPM3 channels in heterologous and native systems. The final aim of this thesis is to explore whether GPCRs can modulate other members of the TRPM family such as TRPM2,

TRPM7 and TRPM8.

Chapter 2. Materials and methods

2.1 Cell culture

2.1.1 Human embryonic kidney and Chinese hamster ovary cells

Human embryonic kidney 293 (HEK293 cells) were grown in DMEM supplemented with penicillin (100 U/ml), streptomycin (100 µg/ml) and foetal bovine serum (FBS) (10%). Chinse hamster ovary (CHO) cells were grown in MEM AQ media with the same supplements noted for HEK293 cell growth. HEK293 and CHO cells stably expressing mouse TRPM3 were grown in the additional presence of G418 (0.5 mg/ml). 12-24 hours before experimentation, cells were plated onto either 96-well black-walled plates (Costar) at high density (~80%) or glass

13mm cover slips at low densities (~20%), which were both precoated in poly-D-lysine (10

µg/ml). Cultures were maintained at 37 °C in a humidified incubator gassed with 5% CO2.

2.1.1.1 TRPM3 HEK293 cell line

HEK293 cells were transfected with a TRPM3α2 plasmid (pcDNA3.1) DNA (provided by Dr

Stephan Philipp; University of Saarland, Homburg, Germany) using lipofectamine 2000 transfection reagent (Invitrogen). Transfected cells were then exposed to culture medium supplemented with G418 (0.5 mg/ml) 24 hours after transfection to select for stable transfectants. Single clones were identified using limited dilution and expansion. Expression of TRPM3 in the clones was functionally assessed by testing pregnenolone sulphate sensitivity in a Ca2+-imaging experiment.

2.1.1.2 TRPM2 HEK293 cell line

HEK293 cells were transfected with a GFP-mTRPM2 plasmid (pcDNA3.1) DNA (provided by

Prof Peter McNaughton; King’s College London, London, United Kingdom) using lipofectamine

2000 transfection reagent (Invitrogen). Transfected cells were then exposed to culture

48 medium supplemented with G418 (0.5 mg/ml) 24 hours after transfection to select for stable transfectants. Transfected cells were then sorted using flow cytometry and selected according to GFP expression. Expression of TRPM2 was confirmed by fluorescence-activated cell sorting using the BD FACSCanto II and the BD FACSAria III and was functionally assessed by testing

2+ H2O2 sensitivity in Ca -measuring experiments.

2.1.1.3 pGLO HEK293 cell line

HEK293 cells stably expressing pGloSensor-22F (Promega, Southampton, UK) were generated by Evangelia Semizoglou, using the clonal selection method mentioned earlier.

2.1.1.4 Transiently transfected HEK293 and CHO cells

TRPM3 HEK293 and CHO cells were transiently transfected with plasmids encoding, mouse

GFP-tagged MOR, rat GFP-tagged cannabinoid receptor (CB1), human pEYFP-N1-Adenosine

2B receptor (a gift from Robert Tarran Addgene plasmid # 37202), pRK5 BARK1 minigene (a gift from Robert Lefkowitz Addgene plasmid # 14695 http://n2t.net/addgene:14695;

RRID:Addgene_14695), muscarinic M1 receptor (a gift from David Julius), mouse RFP-FKBP12-

5ptpase enzyme and PM-FRB-CFP (gifts from Tamas Balla: Addgene plasmid # 67515 http://n2t.net/addgene:67515; RRID:Addgene_67515 and plasmid # 67517 http://n2t.net/addgene:67517; RRID:Addgene_67517, respectively). Transiently transfected cells were experimented on 1-2 days post transfection and then discarded.

HEK293 cells were transiently transfected with either mTRPM7 (pcDNA4, a gift from Andrew

Scharenberg, Addgene plasmid # 45482; http://n2t.net/addgene:45482; RRID:

Addgene_45482) or rTRPM8 (pcDNA3.1) using lipofectamine 2000 transfection reagent

(Invitrogen). mTRPM7 and rTRPM8-transfected cells were functionally tested the following day in Ca2+-imaging experiments and discarded afterwards.

2.1.2 Dorsal root ganglia neurons

Dorsal root ganglion (DRG) neurons were prepared from adult C57Bl/6 mice. Animals were culled by cervical dislocation as approved by the UK Home Office, and spinal ganglia were removed from all levels of the spinal cord using aseptic methods. Ganglia were incubated in

0.25% collagenase in serum-free MEM containing 1% penicillin and streptomycin for 3 hours at 37 °C in a humidified incubator gassed with 5% CO2 in air. Afterwards, the cells were incubated with 0.25% trypsin for 20 minutes. The ganglia were then dissociated mechanically by trituration with flame polished Pasteur pipettes to obtain a suspension of single cells.

Trypsin was removed by addition of 10 ml MEM (containing 10% FBS) followed by centrifugation at 1000 revolutions/min for 10 minutes. The pellet which contained the ganglia was resuspended in MEM containing 1% penicillin and streptomycin, 10% FBS and 0.05%

DNase. The cell suspension was then centrifuged through a 2ml cushion of sterile 15% bovine albumin in MEM at 1000 revolutions/min for 10 minutes. The pellet containing the neurons was thereafter re-suspended in an appropriate volume of MEM containing 10% FBS, 50 ng/ml

NGF and 10 µM cytosine arabinoside to prevent/reduce the growth of non-neuronal cells. The neurons were then plated at a high density (~80%) onto the centre of sterile 13mm glass coverslips previously coated with 10 µg/ml poly-D-lysine for imaging of intracellular calcium levels or at low density (~20%) for electrophysiological experiments. Cultures were maintained at 37 °C in a humidified incubator gassed with 5% CO2 in air and cells were studied

12-24 hours after dissociation.

2.2 Imaging of intracellular calcium levels

2.2.1 Fura-2

Fura-2, a UV excitable ratiometric calcium indicator dye was used to measure changes in

2+ 2+ [Ca ]i. The excitation spectrum for Fura-2 changes upon Ca -binding, emission measured at

>510 nm increases when the dye is excited at 340 nm and decreases at 380 nm excitation.

Cells were loaded with the acetoxymethyl (AM) ester version of the dye which allows Fura-2 to pass across cell membranes by passive diffusion. Intracellular esterases then cleave the ester bonds once the dye is inside the cell, yielding a relatively membrane-impermeant acidic form of the dye (Grynkiewicz et al., 1985). For all calcium imaging experiments the cells or neurons were loaded with Fura-2 for 1-2 hour(s) prior to experimentation. Fura-2 was loaded in physiological extracellular solution supplemented with 0.01% pluronic acid and 1 mM probenecid. Pluronic acid promotes permeation of Fura-2 through the cell membrane and probenecid helps to prevent Fura-2 from being exported from the cell by inhibiting organic- anion transporters in the plasma membrane.

2.2.2 Microscope-based imaging of intracellular calcium levels

Cells or neurons were plated onto 13 mm glass coverslips which formed the base of the perfusion chamber (volume ~1ml). The chamber was mounted on to the stage of an inverted microscope (Nikon Diaphot) and cells were viewed using a 10x Fluor objective with a numerical aperture of 0.5.

Test solutions were applied to cells by local microperfusion of solution through a fine tube placed very close to the cells being studied. The temperature of the superfusate was controlled using a Peltier device connected to a power supply (Marlow Industries, model

SE5010) with temperature measured at the orifice of the inflow tube. Cells and neurons were perfused with solutions supplied from one of eight reservoirs. In experiments using neuronal preparations, neurons were distinguished from non-neuronal cells by a final depolarising challenge with a solution containing 50 mM KCl, which evoked a calcium influx through voltage-gated calcium channels.

Fura-2 signals were measured using RatioMaster Fluorescence Microscopy System (PTI). Cells were excited by light generated by a xenon-arc lamp which was passed alternately through one of two monochromonators (DeltaRam high speed monochromator, PTI) to transmit light of the pre-selected wavelengths (340 nm and 380 nm, ± 2 nm). The emitted light was filtered by a long pass optical filter (>510 nm) and captured by a cooled CCD camera (PTI CoolOne).

Exposure length was equal for each excitation wavelength and determined by the user to ensure adequate signal without saturation of the camera (typically 100-400 ms).

PTI ImageMaster software served as the user interface during the experiments to monitor the fluorescence emission intensity ratios at 340 nm/380 nm excitation and was also used to select individual cells/neurons of interest for analysis. The intensity time-base data was exported into Microsoft Excel (Microsoft) for further analysis and then into Origin (Origin Pro, version 9.1) for graphical representation of the results.

2.3 Fluorometric measurement of intracellular calcium and cAMP levels

To monitor intracellular cAMP levels, we used the pGLOSensor-22F plasmid, which is a biosensor for cAMP that is able to respond rapidly and reversibly to changes in the intracellular concentration of cAMP (Binkowski et al., 2009). Untransfected and pGLOSensor-

22F HEK293 cells were plated in poly-d-lysine-coated 96-well black-walled plates (Corning

2+ Costar) 1–2 d before experimentation. For [Ca ]i measurements, cells were loaded with fura-

2 using the protocol described above.

When measuring changes in intracellular cAMP levels, pGLOSensor-22F-expressing HEK293 cells were loaded with GloSensor (Promega) diluted in physiological saline solution for 45 min before experimentation. In both assays, wells were injected with compounds and responses read using the FlexStation 3 Multi-Mode Microplate Reader (Molecular Devices) using appropriate excitation and emission wavelengths (340/380 excitation, 520 emission for fura-

2 and luminescence for Glosensor).

Experiments were performed using triplicate columns so that multiple (2-4) experiments were carried out per 96-well plate. Cell confluency was typically greater than 80% at the time of experimentation.

In the experimental chapters, n refers to the number of independent experiments performed

(each from a different cell passage). In each individual, a minimum of 3 replicate samples per treatment were used.

2.4 Patch-clamp electrophysiology

Cells and neurons were studied under voltage-clamp conditions using an Axopatch 200B amplifier and pClamp 10.0 software (Molecular Devices). Recordings were performed at varying voltages using borosilicate electrodes (2.5 – 5 MΩ) filled with a solution (see below).

In some experiments, compounds or peptides were also included in the intracellular solution.

Clampfit 10.7 software (Molecular Devices) was used to analyse acquired data. The data was then exported to Microsoft Excel (Microsoft) for further analyses and then into Origin (Origin

Pro, version 9.1) for graphical representation of the results.

2.5 Behavioural assessment of pain responses

Male and female C57Bl6 mice were kept in a climatically controlled environment with ad libitum access to food and water and were acclimatized in the procedure room for 1 hour before the experiments. All behavioural experiments were approved by the King’s College

London Animal Welfare and Ethical Review Board and conducted under the UK Home Office

Project Licence (PF0C9D185). Most behavioural experiments consisted of administration of compounds by intraplantar (i.pl.) injections into the hind paw of mice using Luer-syringe

(Hamilton Co.) fitted with a 26-guage x 3/8 inch needle. Mice were habituated to the experimental Perspex chambers for an hour before the experiment and placed in the chambers immediately after injection of the compounds. The duration of pain-related behaviours (licking, biting, flinching, shaking and elevating the injected paw) were recorded using a digital stop-watch. Total pain response times over the first 2 min were used for analysis as the pain behaviours were largely restricted to this period.

2.6 Solutions and reagents

2.6.1 Solutions

Whole-cell patch clamp and Ca2+-imaging experiments were conducted in a physiological extracellular solution consisting of (in mM) 140 NaCl, 5 KCl, 10 glucose, 10 HEPES, 2 CaCl2, 1

2+ MgCl2 buffered to pH 7.4 (NaOH). For Ca -free experiments, CaCl2 was omitted from the buffer and 1 mM EGTA was added.

Extracellular solutions for pH of 5 and 6 replaced HEPES with 20 mM MES. Solutions for pH of

8 replaced HEPES with 20 mM AMPSO.

Whole-cell patch clamp intracellular solutions consisted of (in mM) 140 CsCl, 10 EGTA, 10

HEPES, 2 MgATP and 2 Na2ATP and was buffered to pH 7.3 (CsOH). In I/V and G/V experiments, 140 mM CsCl was decreased to 70 mM and the solution was supplemented with

70 mM NMDG-Cl. Intracellular solutions for single-channel recordings excluded MgATP and

Na2ATP.

In Ca2+-imaging experiment, the same extracellular solution as patch-clamp experiments was

2+ used. In TRPM7 transfected HEK293 cells, MgCl2 was removed to prevent Mg -mediated block of TRPM7. For experiments using TRPM8-transfected HEK293 cells, solutions were warmed to 28 °C to prevent cold activation of TRPM8. For heat-activation experiments using untransfected or TRPM2-transfected HEK293 cells, solutions were heated up to 50 °C. The temperature of the superfusate was controlled using a Peltier device connected to a power supply (Marlow Industries, model SE5010) with temperature measured at the orifice of the inflow tube.

2.6.2 Reagents

Master stock solutions were aliquoted and stored at -20 °C. Dilutions from these aliquots into extracellular solutions were made on the day of the experiment.

Stock solutions of pregnenolone sulphate, capsaicin, forskolin (Sigma-Aldrich, Poole, UK), bisindolylmaleimide VIII, butaprost (Cayman Chemical, Ann Arbor, USA), WIN 55212–2,

AM251, Gallein, BIIE 0246, BAY 60-6583, CIM0216, PSB 603, KT 5720, naltriben, menthol, forskolin (Tocris Bioscience; Bristol, UK) and SSR240612 (Sanofi Aventis, Surrey, UK) were made in DMSO. GRK2i, carbachol, 2-Furoyl-LIGRLO-amide, adenosine, bradykinin, HOE 140

(Tocris Bioscience), morphine sulphate and naloxone (Sigma-Aldrich) were made in H2O. PGE2

(Sigma-Aldrich) and CAY10598 (Cayman Chemical) were made in ethanol. DAMGO (Tocris

Bioscience; Bristol, UK), SB205607 (Tocris Bioscience; Bristol, UK), U50488 (Tocris Bioscience;

Bristol, UK), PYY (ABGent, San Diego, CA) and H2O2 (Sigma-Aldrich; St Louis, MO) were made in physiological extracellular solution. A stock solution of 8-bromo cAMP was made in H2O titrated with NaOH. Stock solutions of L-AP4 (Tocris Bioscience, Bristol, UK) and (RS)-Baclofen

(Tocris Bioscience, Bristol, UK) were made in molar equivalent solutions of NaOH. Pertussis toxin (0.2 mg/ml; Sigma-Aldrich) was diluted in cell media. GTPS and GDPβS (Sigma-Aldrich) were added to the intracellular solution. CFA was acquired from Sigma-Aldrich.

2.7 Patch-clamp and fluorometric assay data analysis

Electrophysiological data were analysed using ClampFit (Axon Instruments). Origin 9.2

(OriginLab Corporation) and GraphPhad Prism (GraphPad Software, Inc.) were used for statistical analyses and data display.

• For kinetic calculations, the following function was used: 푛 −푡 휏 푓(푡) = ∑ 퐴푖푒 푖 + 퐶 푖=1 Where,

A1 = amplitude and τ1 = time constant of the exponential component and C = y-axis offset.

• Current/Voltage (I/V) plots generated from ramps were converted into conductance/voltage (G/V) plots using the following equation: 퐼 퐺 = 푉(푉푚−푉푟푒푣) Where,

G = conductance, I = current, Vm = membrane potential and Vrev = reversal potential.

• To calculate voltage dependence shifts, G/V plots were fitted with a Boltzmann function:

퐴2 − 퐴1 푦 = + 퐴1 1 + 푒(푉− 푉50)/푑푥 Where,

A1= bottom asymptote, A2 = top asymptote, V50 = centre and dx = time constant.

• To calculate the EC50, IC50 and Hill slope in fluorometric assays, a dose-response curve with variable slope was used:

퐴2 − 퐴1 푦 = 퐴 + 1 1 + 10(퐿표푔푥0−푥)푝 Where,

A1 = bottom asymptote, A2 = top asymptote, Logx0 = centre and p = Hill slope.

A2 was fixed at 0 as baseline was subtracted from the fitted data.

2.8 Experimental design and statistical analyses

Data are presented as box plots showing the mean (square symbol), median (horizontal line), and SEM (box). Data in Gi/o-coupled GPCR-mediated inhibition of TRPM3 (Chapter 4) that were published in the journal eLife are presented as box and whisker plots showing the mean

(square symbol), median (horizontal line), interquartile range (box) and 5% and 95% percentile points (whiskers). For imaging and electrophysiology experiments, n indicates the number of agonist-responding neurons or cells; for multiwell experiments, n indicates the number of independent experiments performed (each in triplicate wells); and in behavioural experiments, n values indicate the number of animals in each group. A priori power calculations were not performed, but our sample sizes are similar to, or greater than, those generally used in the field. Normality of data was tested using the Shapiro–Wilk Test.

Normally distributed data were analysed using an independent-samples t test. Differences in normally distributed data means between three groups or more were analysed using two-

57 way and repeated-measures ANOVA, followed by Tukey’s or Dunnett’s multiple-comparisons post hoc test. Differences in non-normally distributed data means between two groups were analysed using a Mann–Whitney U test. Differences in non-normally distributed data means between three groups or more were analysed using a Kruskal–Wallis test, followed by a

Dunn’s multiple-comparisons post hoc test. All statistical analyses were made using Prism version 7 (GraphPad).

Chapter 3. Characterisation of TRPM3 activity

3.1 Introduction

3.1.1 Introduction to TRPM3

Functional studies of TRPM3 channels were made possible by the discovery of pregnenolone sulphate (PS) as an activator of TRPM3 (Wagner et al., 2008). More pharmacological compounds have since been discovered to modulate TRPM3 channel activity further advancing our understanding of channel function and characteristics (summarised in Figure

3-1).

3.1.2 TRPM3 gene and expression

Alternative splicing of the TRPM3 gene transcript leads to the expression of a large number of isoforms (Oberwinkler and Philipp, 2014). Six splice variants of the human TRPM3 gene were identified (a-f) (Lee et al., 2003). The same study showed that overexpressed TRPM3a variant localised to the intracellular compartments and the plasmalemmal compartment in

HEK293 cells (Lee et al., 2003). Another human variant was also discovered, TRPM31325, which is composed of 1325 amino acid residues and possesses a shorter C-terminal and longer N- terminal than previously described variants and was also activated by cell swelling (Grimm et al., 2003).

In mice, five splice variants (TRPM3α1-5), were initially described which were between 1699-

1721 amino acids (Oberwinkler et al., 2005). From residue 156 onwards, the mouse TRPM3 protein variants and the human TRPM3 protein variants share high sequence homology of

~97%. Each discovered mouse variant has an analogous human variant (mouse TRPM3α1 : human TRPM3c, mTRPM3α2 : hTRPM3a, mTRPM3α3 : hTRPM3b, mTRPM3α4 : hTRPM3e and

60 mTRPM3α5 : hTRPM3d). Therefore TRPM3 splice events are highly conserved in mice and humans (Oberwinkler et al., 2005).

Like many other TRP channels, TRPM3 is expressed in a wide variety of tissues. It is found in the brain, spinal cord, sensory neurons, eye, testis, pituitary, kidney and adipose tissue (Lee et al., 2003). Furthermore, it is expressed in blood vessels, bladder, pancreas, heart, ovaries and sperm cells (Grimm et al., 2003). In situ hybridisation and RT-qPCR experiments detected

TRPM3 mRNA in mouse DRG and TG neurons at levels comparable to TRPV1 (Vandewauw et al., 2013; Vriens et al., 2011).

3.1.3 Activation of TRPM3

3.1.3.1 Pregnenolone sulphate and structurally related compounds

The neurosteroid PS is one of the first agonists of TRPM3 to be identified (Wagner et al.,

2008). PS levels measured in human serum demonstrate similar concentrations in both males and females and decreases from the µM range at birth to the nM range during adulthood (de

Peretti and Mappus, 1983). PS activates TRPM3 in a micromolar range and application of it

2+ leads to increases in [Ca ]i, which were reliant on extracellular calcium and were absent in untransfected cells (Wagner et al., 2008). Furthermore, activation of TRPM3 by PS generated outwardly rectifying currents. PS is capable of activating TRPM3 in <100ms with an EC50 range of 12-23 µM indicating an extracellular binding site. Utilising inside-out patch-clamp, this was proven as application of PS onto the intracellular side of the membrane had no effect (Wagner et al., 2008). Furthermore, application of PS on pancreatic islet and INS1 cells, which endogenously express TRPM3, evoked responses similar to those in HEK293 cells expressing

TRPM3. Therefore PS can be used as a pharmacological probe for TRPM3 expression (Wagner et al., 2008). However, PS has also been observed to reversibly insert into the plasma

61 membrane and altering membrane capacitance (Mennerick et al., 2008) therefore PS may be activating TRPM3 through an indirect mechanism. To test whether PS activates TRPM3 by altering membrane capacitance, a synthetic enantiomer of PS, ent-PS, that is able to alter membrane capacitance identically to the naturally occuring enentiomer of PS. Ent-PS was significantly less effective at evoking TRPM3-mediated responses in TRPM3-expressing

HEK293 cells than PS (Drews et al., 2014). Therefore TRPM3 is believed to possess chirally- selective sites for direct PS binding. However, PS is not a specific ligand for TRPM3 as it can also modulate Kir2.3 potassium channels (Kobayashi et al., 2009), N-methyl-D-aspartate

(NMDA) (Cameron et al., 2012; Jang et al., 2004; Kostakis et al., 2011) and GABAA receptors

(Akk et al., 2001; Shen et al., 2000).

Compounds with similar structures to PS such as pregnenolone, DHEA and DHEA sulphate were also shown to be activators of TRPM3 channels (Wagner et al., 2008). These compounds were less effecacious than PS and were later shown to be partial agonists of TRPM3 (Majeed et al., 2010).

Steroids that are closely related to PS including pregnenolone glucuronidate, pregnenolone hemisuccinate, epipregnenolone sulphate and epiallopregenanole sulphate also activate

TRPM3 (Drews et al., 2014). The double bond between C5-C6 present in PS is not required for

TRPM3 activation as TRPM3 is activated by epipregnenolone sulphate. However, the β- orientation of the sulphate group at the C3 position is important since a compound that lacks the C5-C6 double bond and has the sulphate group in α-orientation was incapable at activating TRPM3 (Drews et al., 2014). Along with the β-orientation, it is crucial for the group at the C3 position to carry a negative charge since compounds with neutral groups at the C3 position such as pregnenolone methyl ether and pregnenolone acetate were unable to

62 stimulate TRPM3. Conversely, compounds containing hemisuccinate and glucuronidate in the

C3 position were also able to activate TRPM3 (Drews et al., 2014).

3.1.3.2 Nifedipine

Electrophysiological investigation of TRPM3 demonstrate that nifedipine, a dihydropyridine calcium channel blocker, is also a TRPM3 agonist. Nifedipine was able to activate TRPM3- mediated responses in a concentration-dependent manner in TRPM3-transfected HEK293 cells (Wagner et al., 2008). Like PS, nifedipine was able to evoke endogenousTRPM3 responses in INS1 and pancreatic islet cells. Co-application of maximally effective concentration of nifedipine and PS to TRPM3-transfected Chinese hamster ovary (CHO) cells resulted in a supra-additive effect demonstrating that nifedipine and PS activate TRPM3 by binding to different sites (Drews et al., 2014). However, nifedipine has also been shown to

2+ activate TRPA1 in DRG and TG neurons as [Ca ]i responses following nifedipine treatment were abrogated in TRPA1 KO mice (Fajardo et al., 2008) and were not significantly altered in

TRPM3 KO mice (Vriens et al., 2011).

3.1.3.3 D-erythro-sphingosine

D-erythro-sphingosine (DeSPH) was purported to be a TRPM3 agonist as application of it

2+ evoked robust increases in [Ca ]i in TRPM3-transfect HEK293 cells but not in untransfected cells (Grimm et al., 2005). Additionally, dihydro-D-erythro-sphingosine and N,N-dimethyl-D- erythro-sphingosine also activate TRPM3 albeit less effectively (Grimm et al., 2005). In another study, short applications of DeSPH did not activate endogenous TRPM3 in INS1 cells or TRPM3-transfected HEK293 cells (Wagner et al., 2008). Prolonged exposure of these cells to DeSPH did lead to large currents though this was also observed in untransfected HEK293 cells which suggest a TRPM3-independent mechanism. However, prior to the generation of

63 the large non-specific currents, a small outwardly rectifying current was observed in TRPM3- transfected HEK293 cells that was not observed in untransfected cells (Wagner et al., 2008).

This small current may be due to activation of TRPM3 channels, but nevertheless the non- specific effects of DeSPH limits its use for functional studies of TRPM3 channels.

3.1.3.4 Clotrimazole and CIM0216

A search for TRPM3 modulators lead to the observation that the antifungal compound, clotrimazole, was found to enhance PS-activated TRPM3 currents (Vriens et al., 2014).

Clotrimazole also induced an inwardly rectifying current component at negative voltages thereby making TRPM3 currents double-rectifying. Application of clotrimazole alone had a modest effect on TRPM3 and activation of the inwardly rectifying currents by PS and clotrimazole is stimulant dependent as clotrimazole along with heat does not produce them

(Vriens et al., 2014). Furthermore, clotrimazole exerts its effect by binding to an intracellular site on TRPM3. This was demonstrated when clotrimazole included in the extracellular pipette solution in inside-out patches did not lead to a double-rectifying current in TRPM3- transfected HEK293 cells suggesting that clotrimazole was being washed away prior to it acting on TRPM3. In contrast, inclusion of clotrimazole in the intracellular solutions during whole-cell recordings elicited a double-rectifying PS-evoked current (Vriens et al., 2014), demonstrating that clotrimazole binds to an intracellular site on TRPM3.

The selective TRPM3 agonist CIM0216 also activates currents with double rectification in

TRPM3 channels in a manner that mimics the effects of co-application of PS and clotrimazole

(Held et al., 2015a). Co-application of clotrimazole and CIM0216 inhibited CIM0216-mediated currents in TRPM3-transfected HEK293 cells suggesting a shared binding site for both

2+ compounds. In contrast, CIM0216 potentiates PS-evoked increases in [Ca ]i in a

64 concentration-dependent manner (Held et al., 2015a). In contrast to clotrimazole, CIM0216 responses are potentiated at higher temperatures suggesting a synergistic effect of heat and

CIM0216. Finally, i.pl. injections of CIM0216 in the hind paws of mice evoked pain-like behaviour that was significantly reduced in TRPM3 KO mice (Held et al., 2015a).

3.1.3.5 Hypotonicity

TRPM3 can also be activated by decreases in osmolarity. TRPM3-transfected HEK293 cells

2+ treated with a hypotonic solution (200 mosmol/litre) had increased [Ca ]i levels as opposed to untransfected cells (Grimm et al., 2003). This response was abolished in calcium-free condition and was reversed by washing with an isotonic solution. This study was completed prior to the discovery of TRPM3 agonists and any synergy between agonists and osmolarity has not been tested yet.

3.1.4 Inhibition of TRPM3

3.1.4.1 Dihydropyridines

Although nifedipine activates TRPM3, other dihydropyridines inhibit the channel. PS-evoked responses in TRPM3-transfected HEK293 cells were inhibited by co-application of 50 µM nimodipine (Drews et al., 2014). Nicardipine and nitrendipine also produced similar inhibitory effects at 50 µM (Drews et al., 2014). These concentrations for all three compounds are much higher than what is required to inhibit L-type Ca2+ channels (approximately 1 µM) (Bean, 1984)

3.1.4.2 Non-selective TRP inhibitors

2-Aminoethoxydiphenyl borate (2-APB) is a non-selective inhibitor of many TRP channels including TRPM3, however, it can activate TRPV1-4 channels (Colton and Zhu, 2007). 2-APB can also activate and inhibit store-operated Ca2+ entry, making it a complex compound to

65 utilise (DeHaven et al., 2008). Other non-selective TRP inhibitors include the trivalent Gd3+ and La3+ ions inhibit many TRP channels including TRPM3 (Grimm et al., 2003; Lee et al., 2003;

Xu et al., 2005).

3.1.4.3 Monovalent and divalent cations

Functional studies on the TRPM3α1 and TRPM3α2 variants revealed that both are inhibited by intracellular magnesium ions in a concentration-dependent manner (Oberwinkler et al.,

2005). This inhibition is not unique to TRPM3 however as TRPM6 (Voets et al., 2004b) and

TRPM7 (Nadler et al., 2001) are also inhibited by intracellular magnesium ions I therefore used intracellular solutions with very low Mg2+ concentrations of approximately 80 µM by buffering Mg2+ with ATP. TRPM3α2 is also inhibited by extracellular sodium, potassium, magnesium and calcium ions in a concentration-dependent manner whereas TRPM3α1 is only inhibited by extracellular magnesium and calcium ions (Oberwinkler et al., 2005).

3.1.4.4 Fenamates

Mefenamic acid, a fenamate non-steroidal anti-inflammatory (NSAID) drug, is a potent and selective inhibitor of TRPM3 with an IC50 of 6.6 µM (Klose et al., 2011). Mefenamic acid inhibited PS-evoked currents in TRPM3-transfected HEK293 cells in a pH dependent manner where its inhibition was strongest at pH 6 and weakest at pH 8 (Klose et al., 2011). Mefenamic acid inhibits TRPM3 by acting on the extracellular side of the channel as dialysis of the compound through the patch pipette did not inhibit PS-evoked currents (Klose et al., 2011).

This inhibition was also demonstrated in INS1 cells where mefenamic acid inhibited PS-evoked calcium influx and insulin secretion. Other fenamates such as DCDPC, meclofenamic acid, tolfenamic acid and niflumic acid also inhibit TRPM3 but in a non-selective manner where they also block TRPM2, TRPV4, and TRPC6 (Klose et al., 2011).

3.1.4.5 Progesterone and structurally related compounds

TRPM3 is also inhibited by the endogenous steroid progesterone, progesterone metabolites,

17β-oestradiol and dihydrotestosterone (Majeed et al., 2012). Progesterone inhibited constitutive TRPM3 activity and PS- and nifedipine-evoked responses in TRPM3-transfected

HEK293 cells whereas progesterone metabolites and 17β-oestradiol only minimally inhibited

TRPM3. Dihydrotestosterone inhibited PS-evoked responses and acted as a competitive antagonist for the PS binding site (Majeed et al., 2012).

3.1.4.6 Thiazolidinediones

The PPAR-γ agonist rosiglitazone inhibits TRPM3 activity following PS and nifedipine activation (Majeed et al., 2011). This inhibition was not mediated by PPAR-γ as the endogenous PPAR-γ agonist, 15d-PGJ2, had no effect on TRPM3 activity. Other thiazolidinedione drugs such as troglitazone and pioglitazone also inhibited TRPM3 activity

(Majeed et al., 2011).

3.1.4.7 Flavonoids and ononetin

The deoxybenzoin compound, ononetin, along with the citrus fruit flavonoids eriodictyol, hesperetin and naringenin have all been identified as potent TRPM3 antagonists (Straub et al., 2013). TRPM3 inhibition by ononetin and naringenin was shown to be reversible in patch- clamp experiments. Both compounds were shown to inhibit PS-, nifedipine- and heat-evoked responses in TRPM3-transfected HEK293 cells and DRG neurons (Straub et al., 2013). Like mefenamic acid, ononetin and naringenin are believed to bind to an extracellular site on

TRPM3 as intracellular application of either had no effect on PS-evoked currents. Ononetin is presumed to act as a competitive antagonist to PS as higher concentrations of PS result in a

67 rightward shift in ononetin inhibition. In contrast, naringenin acts in a non-competitive manner (Straub et al., 2013). Ononetin was also able to inhibit PS-mediated TRPM1 activation and activated TRPA1 whilst naringenin partially inhibited TRPV1 and TRPM7 and both activated and inhibited TRPM8 (Straub et al., 2013).

Another flavanone, isosakuranetin, derived from blood oranges and grapefruits has also been shown to inhibit TRPM3 (Straub et al., 2013). Isosakuranetin is believed to bind to an intracellular site on TRPM3 and inhibits TRPM3-mediated currents in TRPM3-transfected

HEK293 cells and partially inhibits TRPM3-mediated heat-evoked currents (Straub et al.,

2013). However, isosakuranetin also partially inhibits TRPM1-mediated PS responses and partially activates TRPA1 (Straub et al., 2013).

Figure 3-1 Gating of TRPM3

TRPM3 channels can be activated by pregnenolone sulphate (PS), nifedipine (nif), heat and hypotonicity (hypo) with PS being the most widely used pharmacological tool for functional studies of TRPM3. These agonists activate the canonical ion permeation pathway through the poor loop between TM5 and TM6. More recently,

PS coapplication with clotrimazole (clt) or CIM0216 (CIM) application alone have been proposed to open an alternative ion permeation pathway in TM4 of TRPM3 which is more permeable to Na+ ions. TRPM3 is also inhibited by compounds such as ononetin (Ono) and isorakuranetin (isosak) and by extracellular cations including the pore blocker La3+.

3.1.5 Permeation and biophysical properties of TRPM3

Only three TRPM3 variants have been studied electrophysiologically. The human TRPM31325 variant was shown to have greater permeability for Ca2+ ions over both Na+ ions (permeability

+ ratio of 1.57 ± 0.31 for PCa/PNa) and Cs ions (1.14 ± 0.10 for PCa/PCs) (Grimm et al., 2003). The permeability of endogenous TRPM3α2 in pancreatic β-cells is also higher for divalent over monovalent cations where the order of conductivity for divalent cations is Ca2+ > Zn2+ > Mg2+

> Ni2+ (Wagner et al., 2010). Functional characterisation of heterologous TRPM3α1 and

TRPM3α2 variants in HEK293 cells demonstrated that TRPM3α2 is 10 times more permeable for Ca2+ ions and 100 times more permeable for Mg2+ ions (Oberwinkler et al., 2005). This difference in permeability is due to alternative splicing in exon 24, which adds an amino acid domain consisting of 12 amino acids in the assumed pore-region between the fifth and sixth

TM domains. Furthermore, an alanine residue in TRPM3α2 is replaced with a proline residue in TRPM3α1. The study found that this amino acid domain is only found in the hTRPM3c variant and is absent from all other variants (Oberwinkler et al., 2005). This amino acid domain consists of positively charged amino acid residues, which through electrostatic repulsion, may decrease the permeability of TRPM3α1 for cations (Oberwinkler et al., 2005). Almost all

TRPM3 studies are conducted on the TRPM3α2 variant.

More recently, a second permeation pathway through TRPM3 has been proposed. The double-rectifying current caused by binding of PS and clotrimazole is proposed to be carried by different ionic species flowing through two distinct pores (Vriens et al., 2014). The central pore in the postulated pore-region, which mainly conducts divalent cations, and an alternative permeation pathway located in TM4, which mainly conducts Na+ ions. Several observations have been utilised to justify the claim for an alternative permeation pathway.

First, clotrimazole changed the relative contribution of Na+ and Ca2+ ions to the PS-activated

2+ 2+ inward current. This is evidenced by the modest increase in [Ca ]i in Ca imaging experiments

(1.4 fold potentiation) when compared to > 10 fold potentiation of inward currents in patch- clamp experiments (Vriens et al., 2014). Na+ imaging experiments also revealed a very strong

+ 2+ potentiation of [Na ]i (20.4 fold). Additionally, combined Ca -imaging and patch-clamp recordings demonstrated that Ca2+ contributed to roughly 40% of PS-induced inward currents whereas it only contributed to 20% in PS + clotrimazole-induced currents (Vriens et al., 2014).

The inwardly rectifying current was also insensitive to La3+ block where the cation only inhibited outward currents and was abolished by a mutation in the TM4 domain that substituted tryptophan for arginine (W982R). Finally, coapplication of PS and clotrimazole gave rise to two distinct types of single-channel currents with clotrimazole producing distinct openings with lower current amplitudes and longer openings (Vriens et al., 2014). This alternative permeation pathway is likened to the omega current in the Shaker potassium channel which occurs with the substitution of the first S4 arginine with smaller amino acids

(Tombola et al., 2005). This mutation leads to a new permeation pathway through S4.

CIM0216 activates TRPM3 channels in a manner similar to coapplication of PS and clotrimazole evidenced by the opening of both permeation pathways (Held et al., 2015a).

Similar lines of evidence were used to demonstrate CIM0216 opening of the alternative permeation pathway including the insensitivity of inwardly-rectifying currents to La3+ block and greater permeation of Na+ ions (Held et al., 2015a). Although no TRPM3 single-channel recordings were conducted with CIM0216.

Alignment of an amino acid region (consisting of 15 amino acids) in the S4 region of Shaker potassium channels, which includes voltage-sensing arginines, with the entire TRPM3

71 sequence revealed a high degree of similarity in the TM4 region of TRPM3 (40% identity, 60% similarity) (Held et al., 2018). A single gating charge arginine is conserved in the TM4 of TRPM3 at the position corresponding to the second arginine (R2) of the Shaker potassium channel

(Held et al., 2018). Other non-positively charged amino acids in TRPM3 correspond to the R1,

R3 and R4 positions in Shaker potassium channels (Held et al., 2018). The W982R mutation in

TRPM3 which inhibited the alternative permeation pathway substituted an arginine that corresponds to the R1 position in Shaker potassium channels (Vriens et al., 2014). Substitution of an aspartate at position D988 in TRPM3 equivalent to R3 of the Shaker potassium channel by arginine or lysine (D988R, D988K) demonstrated normal activation by PS but fully lacked the inward-rectification with clotrimazole coapplication (Held et al., 2018). Conversely, substituting aspartate with glutamate (D988E) thereby conserving the negative charge at this position did not alter current rectification in with both PS and PS + clotrimazole. However,

CIM0216 application did not activate inward- and outward-rectifying currents in D988R and

D988K TRPM3 channels but did activate D988E TRPM3 producing wild-type-like TRPM3 currents (Held et al., 2018). These results suggest that altering the charge at the D988 position of TRPM3 prevents currents through the alternative permeation pathway but preserves ionic flux through the central pore. Furthermore, substitution of glycine by larger amino acids such as arginine or alanine (G991R and G991A) in a position corresponding with the R4 site in

Shaker potassium channels decreased inward currents following PS + clotrimazole and

CIM0216 application but had no effect on outward currents (Held et al., 2018). Mutations in non-gating charge residues in S4 only had a minor effect on the properties and opening of the alternative permeation pathway in TRPM3 (Held et al., 2018). Other TM regions also affect the alternative permeation pathway. This was demonstrated by mutating the aromatic tyrosine residues in the TM1 region with the non-aromatic threonine (Y878T, Y882T and

Y885T). These mutations eliminated clotrimazole-induced inward currents and CIM0216 activation of TRPM3 channels (Held et al., 2018). Furthermore, neutralising negative residues in TM3 (glutamate with glutamine, E941Q and aspartate with asparagine, D964N) also reduced CIM0216- and clotrimazole-induced inward currents (Held et al., 2018). However, some of the mutagenesis results could be caused by changes to agonist binding sites or by allosteric effects on the central pore. The lack of effect of CIM0216 in some of these mutations suggests an impediment to CIM0216 binding to TRPM3 (Held et al., 2018).

The different ionic permeabilities and current-voltage profiles that CIM0216 and clotrimazole cause when compared to PS in TRPM3 could however be due to the opening of the same central pore but in a different configuration. Other TRP channels have been shown to display different activation profiles by different agonists with TRPA1 activation by the scorpion toxin

(WaTx) being the most recent example (Lin King et al., 2019). WaTx activates TRPA1 by accessing the same intracellular site that reactive electrophiles modify (Lin King et al., 2019).

The toxin stabilises TRPA1 in a biophysically distinct active state that has lower Ca2+ permeability and prolonged channel openings (Lin King et al., 2019). This decrease in Ca2+ permeability leads to WaTx eliciting acute pain and pain hypersensitivity without triggering efferent release of neuropeptides and neurogenic inflammation usually caused by noxious electrophiles (Lin King et al., 2019).

Another example is TRPV1 pore dilation following prolonged exposure to chemical agonists which leads to a three-fold greater permeation to Ca2+ over Na+ ions (Chung et al., 2008).

TRPV1 pore dilation also allowed for the permeation of large organic cations like N-methyl-D- glucamine and large molecules including dyes such as FM1-43 and YO-PRO1 (Chung et al.,

2008). This effect was observed after repetitive or prolonged activation of TRPV1 with

73 capsaicin or protons but not with heat (Chung et al., 2008). Therefore, activation of TRP channels by different agonists or modalities can produce varying activation profiles through a single channel pore.

3.2 Aims of the present study

The aim of the study was to characterise TRPM3α2 function by PS and CIM0216 in a TRPM3-

HEK293 cell line by examining agonist activation and desensitisation rates, current-voltage

(I/V) and conductance-voltage (G/V) relationships and single-channel characteristics. The effects of changes to proton concentrations (pH) on TRPM3 activation by PS and CIM0216 were also examined. Finally, the effect of PS and CIM0216 coadministration in mice is tested.

I performed all experiments included in this chapter but received Dr. Robson Da Costa’s help with the in vivo experiments.

3.3 Results

3.3.1 Characterisation of heterologously-expressed TRPM3

A stable mTRPM3α2 (referred to as TRPM3 hereafter) HEK293 cell line was generated to characterise TRPM3 activation by pregnenolone sulphate (PS) and CIM0216. The aim of the current study was to functionally characterise the properties of TRPM3.

3.3.1.1 Activation of TRPM3 by PS and CIM0216 in fluorometric assays

Since TRPM3 channels are permeable to divalent cations including Ca2+, fluorometric Ca2+ measurements were done. The endogenous neurosteroid PS is one of the most well- characterised TRPM3 agonists (Wagner et al., 2008). The properties of PS as a TRPM3 agonist

2+ were initially examined in a mTRPM3 HEK293 cells where changes in [Ca ]i were measured.

2+ Application of PS evoked concentration-dependent increases in [Ca ]i with a mean EC50 value of 6.59 ± 0.97 µM and a Hill slope of 1.05 ± 0.08 (Figure 3-2). In contrast, untransfected

2+ HEK293 cells did not respond to stimulation by PS, demonstrating that PS evoked [Ca ]I increases by activating TRPM3 (Figure 3-2).

2+ Figure 3-2 PS elicits [Ca ]i-responses in TRPM3 expressing HEK293 cells

A Timecourse of PS-evoked Ca2+-responses in TRPM3 overexpressing HEK293 cells. B Concentration response

2+ curve of [Ca ]i responses in TRPM3-expressing and untransfected HEK293 cells following application of PS, data points are mean ± SEM of triplicate wells (n = 4 independent experiments).

Next, we tested the effect of the synthetic TRPM3 agonist CIM0216 (Held et al., 2015a) on

2+ mTRPM3 HEK293. CIM0216 evoked concentration-dependent increases in [Ca ]i in TRPM3

HEK293 cells with a mean EC50 value of 1.53 ± 0.29 µM (Figure 3-3). In contrast, application of

2+ CIM0216 to untransfected HEK293 cells did not alter [Ca ]i demonstrating that

2+ CIM0216-evoked increases in [Ca ]i is through TRPM3 activation (Figure 3-3). The Hill slope produced by CIM0216 activation was 2.08 ± 0.3 suggesting cooperative agonism and a different mode of TRPM3 activation and thus likely a separate site of action than PS.

2+ Figure 3-3 CIM0216 elicits [Ca ]i-responses in TRPM3 expressing HEK293 cells

2+ Concentration response curve of [Ca ]i responses in TRPM3-expressing and untransfected HEK293 cells following application of CIM0216, data points are mean ± SEM of triplicate wells (n = 6 independent experiments).

3.3.1.2 Activation of TRPM3 by PS in whole-cell patch clamp electrophysiology

2+ Since TRPM3 activation by PS elicited increases in [Ca ]i in a fluorometric assay, we next tested whether it activates TRPM3 in whole-cell patch-clamp experiments. Application of PS

(100 µM) in untransfected HEK293 cells caused no change in membrane current. In contrast,

PS (100 µM) application to TRPM3 HEK293 cells elicited, strong, inward currents that were outwardly rectifying and reached peak conductance in τ= 0.59 ± 0.05 s (n = 8; Figure 3-4 A-C).

These currents quickly desensitised, and a desensitised-current state was reached τ = 7.73 ±

1.22 s after peak-current development (n = 8; Figure 3-4 A). PS-evoked inward currents typically desensitised by 56.8 ± 4.5 % (n = 12; Figure 3-4 A).

Figure 3-4 PS activates outwardly-rectifying currents in TRPM3 expressing HEK293 cells

A Example trace of PS (100 µM)-evoked inward current in whole-cell voltage clamp recordings (-60 mV) in TRPM3

HEK293 cells. B Current-voltage relationships measured at times corresponding to time points 1–3 in A. C

Conductance-voltage relationships measured at time corresponding to time points 2 and 3 in A.

We next characterised TRPM3 single channel conductance following PS activation by performing the inside-out patch-clamp configuration on excised membrane patches. PS only activates TRPM3 through its extracellular site and does not permeate the cell membrane

(Wagner et al., 2008), therefore, PS was included in the patch pipette solution. PS (30 µM) elicited single openings with current amplitudes of 4.53 ± 0.57 pA and conductance values of

56.7 ± 6.6 pS at +80 mV holding potential (n = 3; Figure 3-5 A and B). These values agree with previously published data on PS-evoked single-channel openings in TRPM3 channels (Vriens et al., 2014).

A B

Figure 3-5 PS-mediated activation of TRPM3 leads to single channel openings in TRPM3 expressing HEK293 cells

A Example trace of PS (30 µM)-evoked outward currents in single channel voltage clamp recordings (+80 mV) of inside-out membrane patches from TRPM3 HEK293 cells. O = open and C = closed B Gaussian fit of TRPM3 channel opening events in A with clear peaks demonstrating closed and open current values.

3.3.1.3 Activation of TRPM3 by CIM0216 in whole-cell patch clamp electrophysiology

CIM0216 application (10 µM) to untransfected HEK293 cells did not produce membrane currents (Figure 3-6 A). Interestingly, CIM0216 application (10 µM) on TRPM3 HEK293 cells caused a biphasic current that was composed of initially small, outward-rectifying currents that peaked in τ = 4.4 ± 0.4 s (n = 6) and were followed by strong, double-rectifying currents that reached peak conductance in τ = 14.9 ± 0.9 s (n = 6; Figure 3-6 A-C). Peak currents also desensitised and reached a desensitised-current state in τ = 14 ± 2.6 s after peak-current development (n = 5). CIM0216-evoked inward currents typically desensitised by 47.1 ± 3 % (n

= 6).

CIM0216-evoked outward conductance that plateaued at high voltages, making it possible to use Boltzmann fits to identify the half maximal voltage activation (V50) of CIM0216 at the different stages of TRPM3 activation (Figure 3-6 C). The initial CIM0216-induced current failed to produce outward conductance plots that plateaued in almost all cells tested (4 out of 5); therefore, it was difficult to obtain accurate V50 values for this phase of TRPM3 activation. In contrast, the peak and desensitised stages of CIM0216-mediated activation of TRPM3 produced conductance plots that plateaued at high voltages. Boltzmann fits of these two stages of activation reveal that although CIM0216-mediated activation of TRPM3 desensitised, maximum conductance was reached at lower voltages (V50 of peak CIM0216 conductance: 60.4 ± 2.3 mV; desensitised conductance: 53 ± 2 mV; Figure 3-6 D). This data suggests that TRPM3 desensitisation during CIM0216 application is not caused by loss of affinity for CIM0216.

Figure 3-6 CIM0216 activates double-rectifying currents in TRPM3 expressing HEK293 cells

A Example trace of CIM0216 (10 µM)-evoked inward current in whole-cell voltage clamp recordings (-60 mV) in

TRPM3 HEK293 cells. B Current-voltage relationships measured at times corresponding to time points 1–4 in A.

C Conductance-voltage relationships measured at times corresponding to time points 2-4 in A fitted using the

Boltzmann V50 equation. D V50 values of Boltzmann fits for outward conductance-voltage relationships of 3 and

4 in C (n = 5, both). *p<0.05 paired two-tailed t-test.

CIM0216 has been proposed to activate TRPM3 channels in a similar manner to PS by opening the canonical pore but by also opening a proposed alternative ion permeation pathway where

CIM0216 competes with the intracellular binding site of clotrimazole (Held et al., 2015a).

Therefore, CIM0216 may act on TRPM3 by initially binding to the extracellular site of TRPM3 and eliciting the observed small, outward-rectifying current then crosses the cellular membrane and binds to an intracellular site on TRPM3 thereby evoking the larger, double- rectifying current. Additionally, there may be an enzymatic- or protein-mediated interaction separate from TRPM3 that produces the second phase of CIM0216 activation. To test this, we performed cell-detached inside-out patch-clamp experiments. CIM0216 was applied on the intracellular side of TRPM3 which evoked macro currents that demonstrated the same biphasic activation profile observed in whole-cell patch clamp (Figure 3-7 A). Indeed, both phases of the generated current developed over similar timespans to what was observed in whole-cell patch-clamp experiments where the initial current generated by CIM0216 in inside-out patches developed in τ = 4.55 ± 0.6 s and the second-phase current developed in τ

= 18 ± 3.1 s (Figure 3-7 B). Therefore, the biphasic activation of TRPM3 by CIM0216 is likely to be solely caused by CIM0216 interaction with TRPM3 and is not due to CIM0216 accessibility to binding sites. These observations strongly indicate that CIM0216 exclusively interacts with

TRPM3 at sites accessible form the intracellular side of the membrane.

Figure 3-7 Biphasic CIM0216 activation of TRPM3 does not rely on binding site accessibility

A Example trace of CIM0216 (1 µM)-evoked outward macro currents in inside-out voltage clamp recordings (+60 mV) of membrane patches from TRPM3 HEK293 cells. B Scatter plot representing the rate of activation of the initial and peak CIM0216 currents in both whole-cell (initial n = 6, peak n = 6) and inside-out (initial n = 5, peak n = 3) patch clamps configurations.

We also examined single-channel conductance of TRPM3 caused by CIM0216 in the cell- detached inside-out patch-clamp configuration. CIM0216 application (1 µM) to the intracellular side of TRPM3 elicited two distinct channel openings with current amplitudes of

2.1 ± 0.33 pA for the smaller amplitudes and 3.54 ± 0.42 pA for the larger amplitudes (Figure

3-8 A and B). The conductance values of each were 26.2 ± 4.1 pS and 44.2 ± 5.2 pS at +80 mV holding potentials (n = 5).

Figure 3-8 CIM0216-mediated activation of TRPM3 leads to two distinct channel openings in TRPM3 expressing HEK293 cells

A Example trace of CIM0216 (1 µM)-evoked outward currents in single channel voltage clamp recordings (+80 mV) of inside-out membrane patches from TRPM3 HEK293 cells. O1 = open state 1, O2 = open state 2 and C = closed B Gaussian fit of TRPM3 channel opening events in A with clear peaks demonstrating closed and open current values.

3.3.1.4 TRPM3 desensitisation

PS- and CIM0216-evoked inward currents in TRPM3 HEK293 cells strongly desensitised during sustained application of either agonist, we therefore wanted to explore the mechanisms responsible.

Initially, we examined whether subsequent application of PS elicited desensitised currents. PS

(100 µM) was applied twice with an intermediate washout period. The PS-evoked current following initial application desensitised as observed previously whereas subsequent PS application only produced desensitised currents (Figure 3-9). When normalised to the desensitised-current state of the initial PS application, subsequent challenges with PS evoked

85 current responses of similar amplitude to the desensitised plateau of the first challenge, and these were not subject to further tachyphylaxis (92.8 ± 5.7 %; n = 6)

Figure 3-9 Desensitisation of PS-induced inward currents is not rapidly reversible

Example trace of two subsequent PS (100 µM)-evoked inward current in whole-cell voltage clamp recordings (-

60 mV) in TRPM3 HEK293 cells.

To test whether PS-evoked currents desensitised due to a loss of affinity of TRPM3 for PS, we examined the effect of two applications of 30 µM PS followed by two applications of 100 µM

PS. Application of 30 µM PS in most instances produced modest currents that only slightly desensitised by 20.6 ± 6 % (Figure 3-10 A and B). This was also evident in the following 30 µM

PS application since the current generated was of similar size to the initial PS application and desensitised minimally by 12.3 ± 3.3 % (Figure 3-10 A and B). However, subsequent application of 100 µM PS generated a strong response similar to that described above which declined by 42.4 ± 3.9 % (Figure 3-10 A and B). Accordingly, the second application of 100 µM

PS produced already desensitised currents that declined to a lesser extent of 21.7 ± 4.6 %

(Figure 3-10 A and B). These data suggest that high concentrations of PS elicit a pronounced

86 activation of TRPM3 which desensitises during activation and is not achievable with the use of low concentrations.

Figure 3-10 High concentrations of PS are required to desensitise TRPM3 inward currents

A Example trace of -evoked inward currents following multiple PS applications (30 and 100 µM) in whole-cell voltage clamp recordings (-60 mV) in TRPM3 HEK293 cells. B Scatter plot representing level of desensitisation following application of different PS applications in A (n = 10 for all applications). *p<0.05, **p<0.01 and

***p<0.001 when compared to first application of 100 µM PS, one-way ANOVA followed by Dunn’s multiple comparisons test.

Ca2+ entry through TRP channels has been shown to activate PLCδ which subsequently hydrolyses membrane PIP2 thereby limiting TRP channel activity in a negative-feedback mechanism (Yudin et al., 2011). We therefore included bovine brain-derived PIP2 (10 µM) in the patch pipette and dialysed TRPM3 HEK293 cells for >5 min to ensure that PIP2 has sufficient access to TRPM3. This technique has been shown to work in previously published studies (Borbiro et al., 2015; Lukacs et al., 2013, 2007) although there have been issues

87 reported when brain-derived PIP2 is used due to the precipitation and formation of lipid droplets in aqueous buffers (Tóth et al., 2015). PS application (100 µM) in Ca2+-containing solutions along with PIP2 dialysis generated robust inward currents that desensitised to a similar extent observed with control solutions by 51.9 ± 5.3 (Figure 3-11 A and B).

Figure 3-11 PIP2 supplementation does not prevent desensitisation of PS-induced inward currents

A Example trace of two subsequent PS (100 µM)-evoked inward current (-60 mV) in TRPM3 HEK293 cells dialysed with bovine-brain derived PIP2 (10 µM) through the patch pipette. B Comparison of the level of desensitisation in control (n = 11) and PIP2-treated cells (n = 14).

2+ Increased [Ca ]i desensitises many TRP channels including TRPM8 (Chuang et al., 2004; Yudin et al., 2011) and TRPA1 (Wang et al., 2008). To determine whether Ca2+ entry was responsible for TRPM3 desensitisation, PS was applied in Ca2+ free solutions that were supplemented with

EGTA to chelate free Ca2+ ions. Application of PS (100 µM) in these conditions also caused strong inward currents that desensitised during application to a similar extent seen in Ca2+-

88 containing solutions by 55.6 ± 7.7 % (Figure 3-12 A and B). Furthermore, subsequent application of PS in Ca2+-free solutions generated currents that were 89.7 ± 5.4 % the size of the desensitised current-state during the first PS application thereby suggesting no further desensitisation occurs (n = 4). The rate of desensitisation was similar, and the desensitised- current state was reached in τ = 7.63 ± 1.1 s (n = 5). Altogether, these results indicate that

TRPM3 desensitisation during PS-mediated activation is not due to Ca2+ entry.

Figure 3-12 Desensitisation of PS-induced inward currents is not caused by Ca2+ entry

A Example trace of two subsequent PS (100 µM)-evoked inward current in whole-cell voltage clamp recordings

(-60 mV) in TRPM3 HEK293 cells in Ca2+-free extracellular solutions. B Scatter plot representing the level of desensitisation in control (n = 11) and Ca2+-free conditions (n = 7).

We next examined the effect of extracellular Ca2+ ions on CIM0216-induced inward currents in TRPM3 HEK293 cells. Application of CIM0216 (10 µM) in Ca2+-free solutions also generated a biphasic current that contained an initial, small current with a similar rate of activation of

5.91 ± 1.51 τ (Figure 3-13 A and B) that was observed in experiments with Ca2+-containing

89 solutions. This initial current was also followed by a much larger current, however, unlike the current observed in Ca2+-containing solutions. This second phase of current responses underwent modest desensitisation by 2.93 ± 1.4 % (Figure 3-13 A and D) but remained at a steady-state for prolonged periods of exposure. Furthermore, the second phase current took much longer to plateau than it did in Ca2+-containing solutions (τ = 42.6 ± 4.2 s; Figure 3-13 A and C). This data indicates that TRPM3 desensitisation during CIM0216 activation is due to

Ca2+ entry.

Figure 3-13 Desensitisation of CIM0216-induced inward currents is caused by Ca2+ entry

A Example trace of CIM0216 (10 µM)-evoked inward current in whole-cell voltage clamp recordings (-60 mV) in

TRPM3 HEK293 cells in Ca2+-free extracellular solutions. B Scatter plot representing the rate of activation of the initial phase of CIM0216-induced currents in control and Ca2+-free conditions (n = 6, both) C Scatter plot representing the rate of activation of the peak phase of CIM0216-induced currents in control (n = 6) and Ca2+- free conditions (n = 8) D Scatter plot representing the level of desensitisation in control (n = 6) and Ca2+-free conditions (n = 9). ***p<0.001 two-tailed t-test for C and D.

3.3.1.5 TRPM3 modulation by pH levels

Several TRP channels including TRPV1 (Bevan and Geppetti, 1994; McLatchie and Bevan,

2001), TRPM2 (Du et al., 2009), TRPM7 (Liu et al., 2005) and TRPM8 (Andersson et al., 2004) are activated or inhibited by extracellular and intracellular pH changes. To date, there are no studies demonstrating the effect of H+ on TRPM3 activity. We therefore investigated the effects of pH changes on TRPM3 activity by PS and CIM0216 using both Ca2+ fluorometric assays and whole cell-patch-clamp electrophysiology.

PS concentration-response curves were generated in 96-well plates in extracellular solutions

2+ with pH values of 5, 6, 7.4 and 8 (Figure 3-14 A-E). PS application increased [Ca ]i at pH 6, 7.4 and 8 with an EC50 value of 8.56 ± 1.46 µM (n = 3), 6.6 ± 0.97 µM (n = 4) and 3.32 ± 0.64 µM

(n = 3), respectively. However, PS responses were completely absent in solutions with a pH of

Figure 3-14 TRPM3 activation by PS is inhibited at pH 5

A-D Time courses of PS concentration response curves in different pH environemtns. E Concentration response

2+ curves of [Ca ]i responses in TRPM3-expressing HEK293 cells following application of PS in solutions with different pH levels mean ± SEM.

2+ CIM0216 application mostly had no effect on [Ca ]i in pH 5 solutions in fluorometric experiments where only application of 10 µM CIM0216 was able to mildly activate TRPM3

(Figure 3-15 A). CIM0216 concentration-response curves generated in solutions with pH values of 6, 7.4 and 8 had an EC50 value of 1.87 ± 0.28 µM (n = 3), 1.53 ± 0.29 µM (n = 6) and

1.1 ± 0.21 µM (n = 3), respectively (Figure 3-15 A-E).

Figure 3-15 TRPM3 activation by CIM0216 is inhibited at pH 5

A-D Time courses of CIM0216 concentration response curves in different pH environments. E Concentration

2+ response curves of [Ca ]i responses in TRPM3-expressing HEK293 cells following application of CIM0216 in solutions with different pH levels mean ± SEM.

Whole-cell patch-clamp experiments were then used to confirm these results. Application of pH 5 evoked small and transient inward currents that are consistent with endogenously expressed acid-sensing ion channels (Gunthorpe et al., 2001). This transient current desensitised fully and did not influence the analysis of PS-evoked currents (Figure 3-16 A).

Subsequent application of PS (100 µM) which normally generates large inward currents had no effect in the acidic solution (n = 3). As a control, the same cell was superfused with control extracellular solution (pH 7.4) and application of PS generated strong inward currents (Figure

3-16 A). The effect of pH 8 was also examined in whole-cell patch-clamp experiments. Here,

PS applications (100 µM) generated strong inward currents that displayed similar characteristics to those obtained in control solutions (pH 7.4) and desensitised at a similar rate (τ = 8.5 ± 1.61 s, Figure 3-16 B and C) but desensitised to a larger extent by 79.4 ± 3.6 %

(Figure 3-16 D).

Figure 3-16 TRPM3 activation by PS is inhibited at pH 5 in whole-cell patch clamp recordings

A Example trace of PS (100 µM)-evoked inward current inhibition by solutions with pH 5 but not physiological pH 7.4 in whole-cell voltage clamp recordings (-60 mV) in TRPM3 HEK293 cells. B Example trace of PS (100 µM)- evoked inward currents in solutions with pH 8 in whole-cell voltage clamp recordings (-60 mV) in TRPM3 HEK293 cells. C Scatter plot representing rate of desensitisation of PS-induced inward currents in physiological pH 7.4 (n

= 8) and basic pH 8 (n = 6). D Scatter plot representing level of desensitisation of PS-induced inward currents in physiological pH 7.4 (n = 12) and basic pH 8 (n = 7).

We next performed inhibitory dose-response curves to investigate the relationship between

TRPM3 activity and different concentrations of [H+] ions. Concentrations of PS (20 µM) and

CIM0216 (1 µM) that sub-maximally activate TRPM3 were used to test the effect of changes

+ 2+ in [H ]. Activation of TRPM3 by PS and CIM0216 at pH 8 produced the strongest [Ca ]i-

96 response whereas the response was abolished at pH 5 ([H+] = 10 µM; Figure 3-17 A and B).

This inhibition by acidic environments produced a similar IC50 for TRPM3 activation by both

PS (pH 6.52 ± 0.18) and CIM0216 (pH 6.53 ± 0.06) and a similar Hill slope of -1.02 ± 0.02 for

PS and -0.83 ± 0.3. These results suggest that extracellular protons regulate TRPM3 channels through a mechanism that is essential for channel activity which is shared by agonism by both

PS and CIM0216.

Figure 3-17 pH inhibition-response curves for TRPM3 activation by PS and CIM0216

A and B Inhibitory dose-response curves of PS (20 µM) and CIM0216 (1 µM) by different concentrations of [H+]

(n = 3, both).

3.3.2 TRPM3 activation elicits nociceptive behaviour in vivo

Next, we examined whether administration of PS and CIM0216 elicits nociceptive behaviour in mice. Sole PS or CIM0216 administration through intraplantar (i.pl.) injections in the hind paw of mice has been shown to provoke nociceptive behaviour where this behaviour was

97 absent in TRPM3 KO animals indicating that the agonists are acting through TRPM3 (Held et al., 2015a; Vriens et al., 2011). Through our group’s previous work, administration of either agonist (5 nmole PS and 0.5 nmole CIM0216) on its own produced minimal and variable nociceptive behaviour. Furthermore, CIM0216 has been shown to potentiate PS-mediated responses in TRPM3 HEK293 cells. Therefore, we examined the effect of coadministering PS and CIM0216 by i.pl. injections in the hind paw of mice. PS (5 nmole) and CIM0216 (0.5 nmole) were prepared from stock solutions dissolved in DMSO. When we applied these agonists, we observed minimal pain-like behaviour in mice that lasted for 5.4 ± 2.2 s (Figure 3-18 A). These results contradicted with what is reported in the literature (Held et al., 2015a; Vriens et al.,

2011). One reason behind this disparity could be the DMSO content of the diluted solutions that were injected into the animals. Indeed, when we increased the concentration of the stock solution and diluted less of it to generate 5 nmole PS and 0.5 nmole CIM0216, and therefore lowering total DMSO content from 3.9 % to 0.54 %, coadministration of PS and CIM0216 generated robust and reproducible nociceptive behaviour in mice that lasted for 18.6 ± 2.3 s

(Figure 3-18 A). To confirm this result, a fixed concentration of PS (20 µM) was dissolved in solutions with varying DMSO concentrations and applied to TRPM3 HEK293 cells to generate

2+ a DMSO concentration-response curve. DMSO inhibited PS-evoked increases in [Ca ]i in a concentration dependent manner with an IC50 value of 0.25 ± 0.07 M, which corresponds to

1.74 ± 0.51% DMSO (n = 3; Figure 3-18 B). These results demonstrate that TRPM3 activity is strongly inhibited by DMSO concentrations that are commonly used experimentally.

Figure 3-18 TRPM3 activation by PS and CIM0216 elicits nociceptive behaviour in mice that is inhibited by high concentrations of DMSO

A Scatter plot representing duration of pain responses in mice given intraplantar injections of PS and CIM0216

(5 and 0.5 nmole) in the hind paw in solutions with low (n = 28) and high (n = 17) DMSO concentrations. B

2+ inhibitory dose response curve of different DMSO concentrations on PS (20 µM)-elicited [Ca ]i-responses in

TRPM3 expressing HEK293 cells.

3.4 Discussion

3.4.1 PS-induced activation of TRPM3

2+ Our results demonstrate that PS-induced activation of TRPM3 produces robust [Ca ]i- responses and outwardly-rectifying currents in TRPM3-expressing HEK293 cells. Furthermore, single-channel recordings with PS application in excised inside-out membrane patches demonstrate single TRPM3 channel openings. These results agree with published literature on PS (Vriens et al., 2014).

We also demonstrate that application of a high concentration of PS (100 µM) produces strong inward currents that desensitise during PS application. Current-voltage plots of the peak and desensitised currents during PS application produced outwardly-rectifying currents. When converted to conductance-voltage plots, both states of activation failed to plateau at high voltages and therefore could not be fitted with Boltzmann curves. In contrast, application of lower concentrations of PS (30 µM) produced inward currents that produced steady-state currents that did not strongly desensitise.

The observations obtained with patch-clamp indicate that TRPM3 activity desensitises, which apparently contrasts with the results from experiments with fura-2. It is likely that these apparent differences are explained by inherent differences between the two methodologies.

The Ca2+ signal is not a quantitative measure of the degree of channel activity over time, but

2+ 2+ only measures [Ca ]i. The cells ability to extrude and sequester Ca are critical for the time course of any Ca2+-transient. Large Ca2+ transients are likely to saturate fura-2 which has a Kd of about 100 nM and is saturated at micromolar levels of Ca2+ (Grynkiewicz et al., 1985), and depending on the intracellular fura-2 concentration, it may even buffer the signal at concentrations around the Kd.

Other TRP channels such as TRPM8 desensitise following Ca2+ entry (Yudin et al., 2011). This

2+ is due to Ca -mediated activation of PLCδ which hydrolyses membrane PIP2 thereby inhibiting TRPM8 activity (Yudin et al., 2011). This desensitisation was alleviated by loading

2+ cells with the Ca chelator BAPTA-AM, inclusion of PIP2 and MgATP in the patch pipette

(Yudin et al., 2011). Here, dialysis of TRPM3-expressing cells with bovine-brain derived PIP2 had no effect on desensitisation during PS application. Exclusion of Ca2+ from the extracellular solution also had no effect on TRPM3 desensitisation. Furthermore, the intracellular solution

100 used here contained MgATP and Na2ATP, which provide substrate for phosphoinositide 4-

2+ kinases to synthesise PIP2 and would limit the intracellular concentration of free Mg

(Zakharian et al., 2011). The intracellular solution also contained the Ca2+ chelator EGTA

(10mM), which limits the intracellular concentrations of free Ca2+. These results indicate that

TRPM3 desensitisation during PS-induced activation is not caused by Ca2+ entry or a decrease in membrane PIP2 levels.

One obstacle to examining desensitisation in a wide range of PS concentrations using patch- clamp electrophysiology is that higher concentrations of PS require higher DMSO content to remain soluble in solution. Concentrations of 200 µM PS and above precipitated in solutions with DMSO content of <2 %. Inhibitory concentration response curves with DMSO demonstrate that at these percentages, DMSO inhibits TRPM3 activity in HEK293 cells, therefore, experiments conducted with high PS concentrations were not feasible.

Nevertheless, the observation that 100 µM but not 30 µM PS induced currents that desensitised suggests that high concentrations of PS supra-activate TRPM3 but are followed by tachyphylaxis that may be due to a loss of affinity for PS. However, this is difficult to support since V50 values could not be calculated for the peak and desensitised conductance states and the limiting factor of insolubility with the use of higher concentrations of PS.

3.4.2 CIM0216-induced activation of TRPM3

2+ CIM0216-induced activation of TRPM3 also produced robust [Ca ]i-responses in TRPM3- expressing HEK293 cells. In whole-cell patch-clamp recordings, CIM0216 application consistently produced an initial, small and outwardly-rectifying current that has not been reported previously. This initial current phase was thereafter followed by a much larger, double rectifying current that developed over a longer time course. CIM0216-induced inward

101 currents strongly desensitised during CIM0216 application in a similar manner to that observed in PS-induced currents.

The initial and peak current-states during CIM0216 application were suggest that CIM0216 may be binding to several binding sites on TRPM3 or that it induces a slow conformational change in the channel. CIM0216 was previously reported to compete with the intracellular binding site for clotrimazole (Held et al., 2015a), therefore, the initial peak induced by

CIM0216 could be the result of CIM0216 initially binding to an extracellular site on TRPM3.

CIM0216 could then cross the cellular membrane and bind to an intracellular site on TRPM3 thereby generating the much larger, double rectifying current. Furthermore, the timecourse for the generation of the peak current may indicate that CIM0216 interacts with membrane proteins or kinases in the cytosol to produce this effect. CIM0216-induced macroscopic currents in excised, inside-out patches shared a similar pattern of activation to those observed in whole-cell patch-clamp experiments. Furthermore, the timecourse for the development of both initial and peak currents was the same in both configurations. These results indicate that the dual activation observed with CIM0216 is not due to binding site accessibility and is unlikely to depend on cytosolic proteins or factors.

Here, I have demonstrated the first time single-channel recordings with CIM0216. Application of CIM0216 onto the intracellular side of TRPM3 produced single-channel currents with two distinct conductance states. These distinct single channel conductances resemble those obtained with co-application of PS and clotrimazole, although the authors did not specify the conductance of the new open state caused by clotrimazole (Vriens et al., 2014).

In contrast to TRPM3 desensitisation during PS application, desensitisation of CIM0216 evoked currents was Ca2+-dependent. In Ca2+-free solutions, CIM0216 application also elicited

102 an initial, small increase in inward currents that was followed by the generation of a much larger current. The initial current developed in a similar timecourse to that in Ca2+-containing solutions; however, the peak current took much longer to plateau than it did in control solutions. This suggests that, in control solutions, CIM0216-induced currents may not reach maximal amplitude, since the current develops concurrently with an ongoing Ca2+-dependent.

After conducting the Ca2+-free experiments with PS and CIM0216 application, a study was published showing that PS-mediated desensitisation was due to Ca2+ activation of calmodulin

(CaM) (Przibilla et al., 2018). CaM is a Ca2+-binding protein that undergoes conformational changes after binding up to four Ca2+ ions (Chattopadhyaya et al., 1992). CaM can bind to and adapt the activity of target proteins such as TRP channels in accordance to changes in local

2+ [Ca ]i (Trost et al., 1999; Zhu, 2005). By sequence analysis of CaM-binding proteins and the amino acid sequence of TRPM3, four CaM binding regions have been experimentally identified (Holakovska et al., 2012) but any modulatory effect of CaM binding on TRPM3 activity was unexplored. In the recent study, five CaM binding sites on the N-terminal of

TRPM3 with different binding affinities in dependence of Ca2+ have been discovered (Przibilla et al., 2018). CaM binding to the amino terminus of TRPM3 was found to be mostly Ca2+ dependent whereas CaM binding to other parts of TRPM3 was Ca2+-independent (Przibilla et al., 2018). However, CaM binding sites on TRPM3 were identified by using isolated amino acid fragments of TRPM3 that do not accurately represent native full-length TRPM3 channels, which may bind CaM less frequently. Our results partly contradict the findings of this study as

TRPM3 desensitisation during PS application was not altered in Ca2+-free solutions. However,

TRPM3 desensitisation during CIM0216-induced activation was completely absent under

2+ Ca -free conditions. Additionally, the decrease in V50 values of CIM0216-induced activation

103 from peak to desensitised conductance states suggests that the observed desensitisation is not due to a change in TRPM3 affinity for CIM0216 and is possibly due to another factor such as CaM binding. The recent findings on CaM-mediated modulation of TRPM3 and our results suggest that TRPM3 modulation by Ca2+ entry is a complex matter that requires further investigation.

3.4.3 Modulation of TRPM3 by acidic pH

TRPM3 activation by both PS and CIM0216 was completely inhibited by pH 5. Application of

2+ PS or CIM0216 produced increases in [Ca ]i (within 10-30 seconds); however, pH 5 inhibited

Ca2+-responses completely and both were inhibited with similar concentrations of [H+] and had similar Hill slopes of approximately -1. This suggests that the same amino acid residue is being titrated by acidic pH and is therefore responsible for the observed TRPM3 inhibition.

This inhibition relies on changes to extracellular [H+] rather than intracellular [H+] since earlier studies indicate that it takes 2-5 min for extracellular acidic solutions to produce intracellular acidosis (Andersson et al., 2004). Furthermore, patch-clamp experiments with TRPM3- expressing HEK293 cells were conducted with intracellular solutions with a pH of 7.3 (buffered with 10 mM HEPES), which would delay and limit the intracellular acidosis produced by exposure to low extracellular pH.

Other TRPM channels such as TRPM2, TRPM5 and TRPM8 are also inhibited by acidic pH

(Andersson et al., 2004; Du et al., 2009; Liu et al., 2005). TRPM2 has been shown to be blocked

+ by external [H ] with an IC50 of pHo of 5.3, whereas internal protons inhibit TRPM2 with an

IC50 of pHi = 6.7 (Du et al., 2009). Extracellular proton block of TRPM2 is due to three titratable residues at the outer vestibule of the channel pore: H958, D964 and E994, whereas intracellular pH sensitivity is due to D933 which is located at the C-terminal of the S4-S5 linker

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(Du et al., 2009). TRPM5 is also inhibited by acidic pHo with an IC50 of pH = 6.2 (Liu et al., 2005).

Double mutation of a glutamate residue (E830) in the S3-S4 linker region and a histidine

(H934) in the S5-S6 linker region of TRPM5 abolished TRPM5 sensitivity to external protons

(Liu et al., 2005). Additionally, TRPM8 is inhibited by acidic pHi (Andersson et al., 2004).

However, reduced pHi only TRPM8 activation by cold or icilin, whereas menthol-induced responses were unaffected (Andersson et al., 2004). In contrast, TRPM6 and TRPM7 activity is increased in acidic pH environments (Li et al., 2006). These studies along with our results suggest a shared characteristic of regulation by [H+] across the TRPM family.

3.4.4 Activation of TRPM3 in vivo produces nociceptive behaviour

Finally, we demonstrate that co-administration of PS and CIM0216 through intraplantar injections in mouse hindpaws elicits robust pain-like behaviour. Interestingly, this response is essentially abolished if PS and CIM0216 are diluted in solutions with a relatively high

DMSO concentrations. TRPM3 is inhibited in hyperosmotic solutions (Grimm et al., 2003) and high concentrations of DMSO will increase the osmolality of solutions (Runckel and

Swanson, 1980). However, DMSO is able to cross cell membranes where this feature is critical to protect cell death by crystallising when frozen for cryopreservation (Gurtovenko and Anwar, 2007; Muldrew and McGann, 1990; Notman et al., 2006). Therefore, DMSO crossing the cell membrane will also increase the osmolality of the cell cytosol and in turn equalise the osmolality in and surrounding the cells that express TRPM3; however, this may not be an immediate effect and could occur slowly. Altogether, these results indicate that co-application of PS and CIM0216 in a low DMSO content solution is a reliable method to measure the effect of TRPM3 activation in vivo.

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3.5 Conclusion

PS and CIM0216 evoke TRPM3 activity with functionally distinct properties. Desensitisation of currents produced by PS is Ca2+-independent, whereas CIM0216 evoked current responses undergo Ca2+-dependent tachyphylaxis. Both PS- and CIM0216- evoked currents are inhibited by acidic pH. Furthermore, co-administration of PS and CIM0216 in mice produces robust pain-like behaviour indicative of TRPM3 activity, which can be utilised to assess the effect of

TRPM3 activation or inhibition in vivo.

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Chapter 4. Promiscuous GPCR inhibition of TRPM3 by Gβγ subunits

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4.1 Introduction

GPCR signalling plays an important role in sensory transduction, vision, olfaction and gustation. Activation of Gi/o-coupled receptors such as the µ-opioid receptor usually leads to analgesia and activation of GABAB receptors alleviates pain of muscle spasm in palliative care where these receptors have been targeted by various therapeutics over the years. On the contrary, activation of Gq/11- and Gs-coupled receptors such as the bradykinin B2 receptors and the prostaglandin E2 receptor 2 mostly leads to the sensitisation of nociceptors in cases of inflammation and produce contraction in other tissues including airway smooth muscles.

GPCRs mediate these effects through various means one of which is by regulating ion channel activity.

4.1.1 GPCRs and analgesia

4.1.1.1 Opioids

The main target treatment for moderate and severe pain remains to be µ-opioid receptor

(MOR) agonists such as morphine and fentanyl. Intrathecal and systemic administration of these agonists in animals and humans produces strong analgesia (Chen and Pan, 2006;

Onofrio and Yaksh, 1990; Wigdor and Wilcox, 1987). Agonists of MOR, δ (DOR), κ (KOR) and opioid receptor-like 1 (ORL1) inhibit neuronal activity through several mechanisms means.

The first is through the inhibition of VGCC in DRG neurons which limits the release of excitatory neurotransmitters (Acosta and López, 1999; Beedle et al., 2004; Kohno et al., 1999;

Moises et al., 1994; Pan et al., 2002; Wu et al., 2004). Synaptic release of glutamate, a major excitatory neurotransmitter e.g. mainly relies on the activity of P/Q-type and N-type VGCC that are found on glutamatergic afferent terminals (Luebke et al., 1993; Reid et al., 1997;

108

Wheeler et al., 1996). GPCR-mediated inhibition of VGCCs includes a rapid, membrane- delimited inhibition mediated by direct interactions with Gβγ subunits in a voltage-dependent manner. Gβγ protein inhibition of the VGCC channel encoded by exon 37b, CaV2.2 e[37b], is relieved by strong membrane depolarisation (Raingo et al., 2007). A slower, voltage- independent modulation of VGCCs by second messenger molecules follows. Prolonged activation of ORL1-opioid receptors in DRGs by nociceptin stimulates Gαq/11 subunits subsequently leading to PKC-dependent internalisation of the ORL1-CaV2.2 complex (Altier et al., 2006)

Opioid receptors also inhibit neuronal excitability is through activation of GIRK channels in postsynaptic neurons in the spinal cord (Marker et al., 2006; Schneider et al., 1998). MOR activation leads to significant hyperpolarisation of spinal superficial dorsal horn neurons via activation of GIRK channels (Marker et al., 2006; Schneider et al., 1998). MOR has been found to be coexpressed with GIRK1 and GIRK2 in a subset of lamina II excitatory neurons (Marker et al., 2006, 2005) where GIRK1 and GIRK2 KO mice exhibited diminished antinociceptive effects of intrathecally administered morphine (Marker et al., 2004; Mitrovic et al., 2003).

4.1.1.2 GABAB

GABA is the predominant inhibitory neurotransmitter in the central nervous system. GABA activates ionotropic GABAA receptor and the GPCRs GABAB. GABAB is present in primary afferent neurons, DRG neurons, spinal cord and brain (Price et al., 1987, 1984; Towers et al.,

2000). Capsaicin-induced degeneration of primary afferent fibres in neonatal rats halves

GABAB receptor density indicating that at least 50% of GABAB receptors are present in TRPV1- expressing primary afferent terminals (Price et al., 1987, 1984). Activation of GABAB1 receptor subunits has no effect on normal TRPV1 activity but does revert its sensitised state during

109 inflammation therefore blocking pathological but not acute TRPV1 pain signals (Hanack et al.,

2015).

Activation of GABAB receptors by intrathecal injections of baclofen produces antinociceptive effects in animal models of acute pain (Dirig and Yaksh, 1995; Hammond and Drower, 1984).

This is also seen in humans where intrathecal administration of baclofen provides analgesia in patients suffering from chronic and neuropathic pain (see Slonimski et al., 2004 for review).

Additionally, intrathecal infusion of baclofen has been shown to reduce musculoskeletal pain and pain caused by spinal cord injury and stroke (Becker et al., 2000; Loubser and Akman,

1996; Taira et al., 1994).

GABAB signalling produces analgesia by similar mechanisms to opioid receptors. GABAB activation has been shown to inhibit VGCCs in DRG neurons (Dolphin and Scott, 1986; Menon-

Johansson et al., 1993). Application of baclofen or GABA shortens action potential duration in

DRGs by reducing N-type VGCC current (Green and Cottrell, 1988). Furthermore, GABAB activation leads to the activation of GIRK channels (Marker et al., 2006). Activation of presynaptic GABAB also limits the release of glutamate from primary afferent (Ataka et al.,

2000; Iyadomi et al., 2000; Wang et al., 2007).

4.1.1.3 NPY receptors

NPY receptors Y1 and Y2 (NPY1 and NPY2) are each expressed in roughly 20% of sensory neurons and their agonist PYY is an important co-transmitter in sympathetic neurons

(Brumovsky et al., 2005; Ji et al., 1994; Taylor et al., 2014; Zhang et al., 1997, 1994). Activation of NPY2 receptors in primary afferent terminals in lamina II by NPY 13-36 and ahx 5-24 NPY reduces glutamate release from spinal synaptosomes (Martire et al., 2000). NPY1 and NPY2 receptors have been shown to couple to GIRK channels and VGCCs in thalamic brain slices

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+ (Sun et al., 2001b, 2001a). NPY1 receptor activation promotes K currents through GIRK channels at somatic and dendritic locations whereas NPY2 receptor activation inhibits N- and

P/Q-type VGCCs at inhibitory GABAergic presynaptic terminals (Sun et al., 2001b, 2001a).

4.1.1.4 Adenosine receptors

Adenosine receptor involvement in nociception has been studied extensively. Of the adenosine receptors, the A1 and A3 subtypes are Gi/o-coupled receptors and the A2A and A2B subtypes are Gs-coupled. A1 receptors are expressed in nociceptive neurons, spinal cord and brain (Lima et al., 2010; Reppert et al., 1991; Schulte et al., 2003). A1 receptor activation limits thermal hyperalgesia and mechanical allodynia in humans and rodents (Ekblom et al., 1995;

Sylvén et al., 1996). Furthermore, A1 receptor activation reduces hyperalgesia caused by a surgical incision in rats (Zahn et al., 2007). Additionally, administration of adenosine prior to and during major surgeries provided prolonged postoperative pain relief and decreased the need for postoperative opioid analgesics (Apan et al., 2003; Fukunaga et al., 2003; Segerdahl et al., 1997, 1996, 1995). Interestingly, the A1 receptor has been shown to be solely responsible for antinociceptive effects of acupuncture in mice (Goldman et al., 2010). This study was able to replicate the effect of acupuncture by the direct injection of the selective

A1 receptor agonist CCPA. Moreover, inhibition of AMP deaminase and adenosine deaminase, which are enzymes responsible for the degradation of extracellular adenosine, potentiated acupuncture-elicited increases in adenosine and prolonged its antinociceptive effect

(Goldman et al., 2010). These results suggest that A1 receptor agonists may be a novel analgesic therapy.

The A3 receptor has also been implicated in pain states. Activation of A3 receptors by systemic application of the selective agonist IB-MECA produced antinociceptive effects during the

111 second phase of the formalin test (Yoon et al., 2005). This effect did not extend to protective nociceptive responses such as those to noxious thermal or mechanical stimuli (Yoon et al.,

2005). Furthermore, intrathecal administration of the A3 receptor antagonist MRS1220 reversed the antinociceptive effect of adenosine in the second phase of the formalin test

(Yoon et al., 2006). These results clearly indicate an important role for spinal A3 receptors in the analgesic effects of adenosine. The analgesic effect of A3 receptor activation has also been observed in the periphery, where intraplantar administration of another selective A3 agonist,

MRS5698, reversed mechanical allodynia in the chronic constriction injury model of neuropathic pain in rats in a dose-dependent manner (Little et al., 2015).

4.1.2 GPCRs and inflammatory pain

4.1.2.1 Prostaglandins

Prostaglandins are lipid-derived autacoids generated through the sequential actions of cyclooxygenase and prostaglandin synthase. Well-known prostaglandins include PGD2, PGE2

PGI2 and PGF2a that act on eight prostanoid receptors which propagate several features of inflammation including oedema and pain. Blocking the synthesis of prostaglandins with non- steroidal anti-inflammatory drugs (NSAIDs) that inhibit COX-2 is an effective anti- inflammatory, antipyretic and pain-relieving treatment (see Vane, 1971; Vane and Botting,

1997 for review). Prostaglandins are important for other physiological functions including

PGI2’s role in vasodilation (Frölich, 1990) along with its role with PGE2, PGF2a and PGD2 in gastric mucosal defence (Peskar, 1977; Wallace, 2008).

PGE2 activates four PGE2 receptors, EP1 (Gq/11-coupled), EP2 and EP4 (Gs-coupled) and EP3

(Gi/o-coupled) which are all expressed in subsets of DRG neurons (Natura et al., 2013).

Coupling of EP receptors to various G proteins accounts for the variety of PGE2’s effects. PGE2-

112 induced activation of EP1 and EP4 sensitises TRPV1 by PKC and PKA mediated phosphorylation in HEK293 cells and DRG neurons (Bhave et al., 2002; Distler et al., 2003;

Moriyama et al., 2005). Phosphorylation of TRPV1 by PKC and PKA relies on the formation of a complex between these kinases, the scaffolding protein AKAP79/150 and TRPV1 (Zhang et al., 2008). In contrast, activation of EP3 receptors by intrathecal injection of the selective agonist ONO-AE-248 produced antinociceptive effects in rats with inflamed knee joints

(Natura et al., 2013). The author’s suggest that the antinociceptive effect of EP3 receptor activation may be an endogenous pain control mechanism to protect against inflammatory pain (Natura et al., 2013). However, nociceptive agonists that cause pain in the periphery applied intrathecally can dampen input to the dorsal horn through presynaptic inhibition and thereby produce antinociception. An example of this is the ability of TRPV1 activators to produce spinal analgesia (Dickenson et al., 1990; Eimerl and Papir-Kricheli, 1987) and that spinal TRPA1 activation by the metabolites of acetaminophen produces antinociception

(Andersson et al., 2011).

Several mechanisms in concert facilitate the sensitising and hyperalgesic effect of PGE2 including its enhancement of tetrodotoxin-resistant (TTX-R) sodium currents in DRG cells.

Nociceptive DRG neurons express two slowly-inactivating TTX-R sodium channels, the Nav1.8 and Nav1.9 channels (Rush et al., 1998). Nav1.8 is the major transient TTX-R channel whereas

Nav1.9 accounts for persistent TTX-R sodium currents (Meves, 2006). Application of PGE2 to

DRG neurons enhanced the TTX-resistant sodium current by increasing the slope of the activation curve and shifting it to more negative potentials, which results in an increase in the excitability of sensory neurons. This effect is observed in colonic and cutaneous DRG neurons and in pulmonary DRG neurons (Kwong and Lee, 2005). Prolonged application of PGE2 (1 hour)

113 increased the persistent TTX-R sodium current that is attributed to Nav1.9. This effect leads to depolarisation and augmented firing of spikes (Rush and Waxman, 2004).

4.1.2.2 Bradykinin

Bradykinin is a peptide that is predominantly generated by the enzymatic action of kallikreins on kininogen precursors (Howl and Payne, 2003) and is an inflammatory mediator that exerts its effect by activating the two Gq-coupled bradykinin receptors BK1 and BK2. Bradykinin preferentially binds BK2 receptors whereas its metabolite, des-ARG9-bradykinin, selectively binds BK1 receptors (Regoli et al., 1993). BK2 receptors are constitutively expressed in a variety of cells including nociceptive primary afferent neurons, macrophages and smooth muscle, whereas BK1 receptor expression is upregulated following injury or inflammation

(Regoli et al., 1981; Segond von Banchet et al., 1996). Therefore, in uninflamed tissues, bradykinin exerts its effect mostly through BK2 receptor activation and application of selective BK1 receptor agonists typically does not sensitise nociceptors in normal condition.

Various studies demonstrated this through utilising selective BK2 receptor antagonists such as HOE 140 and WIN 64338 and with BK2 receptor KO mice.

BK1 and BK2 receptors have been shown to contribute to the development of pain and hyperalgesia in the periphery and in the spinal cord (Ferreira et al., 2002; Fox et al., 2003;

Pesquero et al., 2000; Quintão et al., 2008). BK2 antagonist administration in rats and mice relieved the early phase (0-5 minutes) of formalin-induced nociception (Chichorro et al., 2004;

Corrêa and Calixto, 1993; de Campos et al., 1999; De Campos et al., 1996). Similarly, BK2 receptor antagonists alleviated capsaicin-evoked paw licking in mice and acute abdominal writhing responses to acetylcholine-, acetic acid- and kaolin intraperitoneal injections (de

Campos et al., 1999; De Campos et al., 1996; Heapy et al., 1993; Ikeda et al., 2001).

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Administration of the BK1 receptor antagonist des-Arg9,Leu8-bradykinin also alleviated early phase formalin-induced nociception in mice and rats (Corrêa and Calixto, 1993; Shibata et al.,

1989; Sufka and Roach, 1996). This data suggests that BK1 receptor expression is induced rapidly upon tissue injury or that BK1 receptor activation, even though BK1 receptor expression is low, can still exert prominent effects (Ferreira et al., 2008). Furthermore, administration of selective BK1 receptor agonists excited and sensitised cutaneous nociceptor in naïve monkeys and human (Eisenbarth et al., 2004; Khan et al., 1992), caused nocifensive behaviour in the mouse paw (Jesse et al., 2009; Porreca et al., 2006) and induced mechanical hyperalgesia in the rat paw (Poole et al., 1999).

Bradykinin receptor activation affects TRP channel activity where BK2-mediated nocifensive responses in the mouse paw are mediated by TRPV1 receptor stimulation (Chuang et al.,

2001; Ferreira et al., 2004; Shin et al., 2002). Bradykinin application to rat DRGs sensitised heat-activated currents. This sensitisation was found to be reproducible by application of the

PKC activator PMA and was inhibited by application of the nonselective protein kinase inhibitor staurosporine (Cesare and McNaughton, 1996). A later study confirmed that PKCε is responsible for the sensitisation of heat-evoked currents (Cesare et al., 1999). Application of

PMA to DRG neurons and TRPV1-expressing oocytes sensitised TRPV1 currents (Premkumar and Ahern, 2000). This effect was also observed with bradykinin-induced activation of BK2 receptors where inhibition of PKC prevented BK2-mediated sensitisation of TRPV1

(Premkumar and Ahern, 2000). In TRPV1/BK2-transfected HEK293 cells, application of bradykinin lowered the temperature threshold for TRPV1 activation to levels below physiological body temperatures in a concentration-dependent manner through PKC activation (Sugiura et al., 2002), and in capsaicin-sensitive DRG neurons (Sugiura et al., 2002).

Furthermore, PKC activation by PMA sensitised TRPV1 receptors to protons, anandamide and

115 capsaicin in TRPV1-transfected HEK293 cells, oocytes and DRG neurons (Crandall et al., 2002;

Vellani et al., 2001). This sensitisation was abolished by inhibiting the interaction of

AKAP79/150 with TRPV1 and by downregulating the expression of AKAP79 using siRNA (Zhang et al., 2008). Therefore, both bradykinin and PGE2 sensitise TRPV1 through PKC.

BK2 also decreases the excitability of Kv7 channels as demonstrated in small nociceptive rat

DRG neurons (Liu et al., 2010). Application of bradykinin caused a concentration-dependent inhibition of M currents in DRG neurons (Liu et al., 2010). Pretreatment with HOE 140 and the

PLC inhibitor edelfosin abolished M current inhibition. Furthermore, BK2-mediated M current

2+ inhibition was diminished by disrupting intracellular Ca signalling through the use of IP3 receptor antagonists, sarco-/endoplasmic reticulum Ca2+-ATPase inhibitor and intracellular

Ca2+ chelation (Liu et al., 2010). Inhibiting PKC with BIM had no affect and therefore these results indicate that bradykinin-mediated M current inhibition relies on PLC- and IP3-mediated increase of intracellular Ca2+ (Liu et al., 2010). Furthermore, intraplantar administration of the

Kv7 channel activator retigabine reduced bradykinin-induced pain responses in mice and reversed bradykinin inhibition of M current in DRG neuron patch-clamp experiments (Liu et al., 2010).

4.1.3 TRPM3 in nociception and inflammatory hyperalgesia

TRPM3’s role in nociception was demonstrated where intraplantar injection of PS evoked nocifensive responses in wildtype mice but not in TRPM3 KO mice suggesting that TRPM3 activation promotes pain (Vriens et al., 2011). The same study demonstrated TRPM3 activation by heat and that TRPM3 KO mice exhibited strong deficits in their avoidance response to noxious heat. Interestingly, mice lacking TRPM3 also failed to develop heat hyperalgesia associated with inflammation in a Freund’s Complete Adjuvant inflammatory

116 model (Vriens et al., 2011). Additionally, administration of TRPM3 antagonists isosakuranetin and hesperetin reduced the sensitivity of mice to noxious heat (Straub et al., 2013).

Altogether these results demonstrate that TRPM3 plays a role in nociception and inflammatory pain.

4.1.3.1 TRPM3 modulation by GPCRs

GPCR modulation of TRPM3 was first studied in TRPM31325-transfected HEK293 cells that were overexpressing Gq-coupled histamine H1 receptor and endogenously expressing muscarinic receptors (Grimm et al., 2003). Activation of H1R and muscarinic receptors with carbachol did

2+ not affect Mn entry through TRPM31325 (Grimm et al., 2003). Conversely, depletion of intracellular Ca2+ stores in hTRPM3a-transfected HEK293 cells by treatment with the Ca2+-

ATPase inhibitor thapsigargin or carbachol increased Ca2+ entry by roughly 40% (Lee et al.,

2003). This effect was not examined further.

More recently, TRPM3 activity has been shown to be positively modulated by membrane phosphatidylinositol phosphates (PIPs) (Badheka et al., 2015; Tóth et al., 2015). Application of either PIP2 or ATP to the intracellular side of excised TRPM3-HEK293 expressing cells promotes recovery of TRPM3 from desensitisation (Tóth et al., 2015). The effect of ATP is thought to be mediated by the activation of phosphatidylinositol kinases (PI-Ks) which leads to the resynthesis of PIPs in the plasma membrane (Tóth et al., 2015). Multiple PIPs were shown to directly enhance TRPM3 activity in membrane-delimited inside-out patches with the following rank order of potency PI(3,4,5)P3 > PI(3,5)P2 > PIP2 and PI(3,4)P2 >>PI(4)P (Tóth et al., 2015). TRPM3 activity was rapidly and reversibly inhibited by membrane PIP depletion.

Additionally, activation of the Gq/11-coupled muscarinic M1 receptor rapidly and reversibly inhibited TRPM3 channels (Badheka et al., 2015; Tóth et al., 2015). Therefore, only Gq-coupled

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GPCR modulation of TRPM3 activity was investigated. Additionally, these studies were conducted in heterologous expression systems and may not reflect what occurs in more relevant physiological environments such as in DRG neurons or in vivo.

We have recently demonstrated, along with two other groups, that activation of Gi/o-coupled

GPCRs inhibits TRPM3 both in vitro and in vivo (Badheka et al., 2017; Dembla et al., 2017;

Quallo et al., 2017). This inhibition is independent of Gαi/o subunits and any downstream signalling processes and is instead due to the direct interaction of Gβγ subunits with TRPM3.

Furthermore, TRPM3 was also inhibited by activation of heterologously expressed Gq-coupled

GPCRs in recombinant systems in a Gβγ-dependent manner (Badheka et al., 2017). Gαq subunits were also shown to not interact with TRPM3 channels (Dembla et al., 2017).

However, TRPM3 modulation by Gq/11-coupled GPCRs in native systems and by Gs-coupled

GPCRs was not examined.

4.2 Aims of the present study

The first aim of the study was to examine whether TRPM3 channel activity can be modulated by Gi/o-coupled GPCR activation and if so by what mechanism. The work presented for this aim is currently published in the journal eLife (Quallo et al., 2017). Dr. Talisia Quallo performed all imaging experiments, I performed the electrophysiological experiments and

Mr. Clive Gentry performed the in vivo experiments. For the adenosine study, I performed all the experiments and collaborated with Dr. Robson Da Costa’s in some of the imaging and in vivo experiments.

Since we demonstrated that TRPM3 activity is inhibited by Gi/o-coupled GPCR activation by

Gβγ proteins, we examined whether activation of Gq/11- and Gs-coupled GPCRs would also inhibit TRPM3 through Gβγ. Most results for this aim have been accepted for publication in

118 the Journal of Neuroscience (Alkhatib et al., 2019). I performed all electrophysiological experiments and all imaging experiments in HEK293 cells, Dr. Da Costa performed all the DRG imaging experiments and both Dr. Da Costa and I performed the in vivo behavioural experiments with TRPM3 and TRPV1 agonist application. Mr. Gentry performed the in vivo experiments in mice with chronic inflammation and Dr. Quallo obtained preliminary data on

PGE2-mediated inhibition of TRPM3.

These experimental aims were investigated using patch-clamp electrophysiology of TRPM3- transfected CHO and HEK293 cells and mouse DRG neurons, single cell Ca2+-measurements of

TRPM3-transfected HEK293 cells and mouse DRG neurons and cell population Ca2+ and cAMP measurements in HEK293 cells. The behavioural responses of wildtype and Trpm3-/-mice were also assessed to examine the effect of GPCR activation on TRPM3-mediated nociception.

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4.3 Results

4.3.1 Gi/o-coupled GPCR regulation of TRPM3 activity

4.3.1.1 Morphine, baclofen and PYY inhibit TRPM3-mediated PS-induced Ca2+ responses

We first examined the effects of the prototypical opioid receptor agonist, morphine, to test whether Gi/o-coupled GPCR activation can modulate native TRPM3 channels in DRG neurons.

Two consecutive applications of a submaximal concentration of pregnenolone sulphate (PS,

2+ 20 µM) were used to investigate the effect of morphine on TRPM3-mediated [Ca ]i-responses in isolated mouse DRG neurons. The first application of PS was used to identify TRPM3 expressing neurons whereas the second was to assess the effect of pharmacological treatments. The response amplitude of the second PS challenge is referred to as R (relative %

2+ response). In control experiments, the second PS challenge evoked [Ca ]i-responses (R) that were 63 ± 2 % of the first PS response amplitude (Figure 4-1 A). Morphine application (10 µM) for 2 min prior to and during the second PS challenge significantly reduced R to 12 ± 1 %

(Figure 4-1 B and C). In most PS-responsive neurons (55%, n = 115/209), application of

2+ morphine completely abolished PS-evoked [Ca ]i responses (R < 5 %).

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Figure 4-1 Morphine inhibits TRPM3 channels expressed on sensory neurons.

2+ + A Traces of DRG [Ca ]i responses evoked by two sequential PS challenges (20 µM) followed by high K (50 mM

KCl). B Effect of treatment with 10 µM morphine for 2 min before and during the second PS challenge.

F(340/380) indicates fura-2 emission ratio. The lower panels in A and B are pseudocolour images illustrating the change in F340/380 ratio in response to PS and KCl. The bar indicates the colours corresponding to various

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F340/380 values. C Box and whisker plots and data points showing the amplitudes for responses to the second

PS challenge in A and B, ***p<0.001; Mann-Whitney U test (control, n = 174; morphine, n = 209). D Effect of treatment with morphine (10 µM, n = 323), morphine (10 µM) and naloxone (1 µM, n = 110) and morphine (10

2+ µM) following an incubation with pertussis toxin (200 ng/ml for 2.5 h-18h, n = 253) on [Ca ]i responses evoked by the second PS (20 µM) challenge using the protocol in (A and B). Control group, n = 188. E Traces displaying

2+ neuronal [Ca ]i responses to two PS (5 µM) challenges in the absence and presence of naloxone (1 µM), followed by a 50 µM PS challenge and high K+ (50 mM KCl). ***p<0.001, compared to control. ††† p<0.001, compared to morphine (10 µM), Kruskal-Wallis. F Whole cell recording illustrating that morphine reversibly inhibits PS-evoked outward membrane currents in DRG neurons (+40 mV). G Inhibitory effect of morphine on whole cell outward current in a CHO cell co-expressing TRPM3 and µ-opioid receptor (+60 mV). H Current-voltage relationships for another TRPM3/µ-opioid receptor expressing CHO cell measured at times corresponding to time points 1–5 in panel G.

We utilised the opioid receptor antagonist naloxone to confirm that morphine was inhibiting

TRPM3 through a receptor-mediated mechanism. Naloxone (1 µM) fully reversed morphine

(10 µM)-mediated inhibition since the relative amplitude (R) produced by the second PS challenge in the presence of morphine + naloxone was 73 ± 3 % (Figure 4-1 D) which was similar to untreated control neurons (R = 64 ± 3 %). Therefore, morphine inhibits TRPM3 by activating opioid receptors expressed in DRG neurons.

We then investigated the role of Gi/o subunits by treating DRG neurons with pertussis toxin

(PTX), which acts by preventing the coupling of Gαi/o proteins to their cognate GPCR by catalysing ADP-ribosylation of Gαi/o proteins. Incubation with PTX (200 ng/ml, for 2.5-18 hours) significantly reduced the inhibitory effect of morphine in a large proportion of neurons

(morphine: R = 16 ± 2 %; morphine + PTX: R = 46 ± 2 %, Figure 4-1 D). These results suggest that activation of opioid receptor and subsequent Gαi/o protein signalling is able to inhibit the

122 activity of TRPM3 channels in sensory neurons. Opioid receptors like many Gi/o coupled GPCRs may possess constitutive activity (Rosenbaum et al., 2009). We therefore examined whether such tonic activity modulates endogenous TRPM3 activity in DRG neurons by using two

2+ consecutive challenges of a low concentration of PS (5 µM), which evoked [Ca ]i-responses in only a small percentage of neurons. In experiments where cells were treated with naloxone during the second PS challenge, the number of responding neurons increased significantly (p

< 0.001, Fisher’s) from 3.6 % (n = 36/999) to 9 % (90/999, Figure 4-1 E). In contrast, the number of PS responders fell from 3.5 % (n = 40/1153) to 1.8 % (n = 21/1153) in the absence of naloxone. These results suggest that constitutive activity of opioid receptors exerts a tonic inhibition of TRPM3 in DRG neurons.

2+ To confirm the observed opioid receptor-mediated inhibition of PS-evoked [Ca ]i-responses we used patch-clamp recordings. PS activation of TRPM3 produces marked outward current rectification as shown in chapter 3 (Wagner et al., 2008) and we therefore examined DRG neurons at a holding potential of +40 mV. Application of PS (50 µM) readily evoked membrane currents in a subset of isolated DRG neurons (Figure 4-1 F). Application of morphine (10 µM) during PS stimulation produced a rapid, near-complete and reversible inhibition of the PS- evoked current. We next studied the effect of morphine on PS-evoked currents in CHO cells co-expressing TRPM3 and the µ-opioid receptor (MOR). PS-evoked (100 µM) currents were reversibly inhibited by co-application of morphine (10 µM; inhibition 89 ± 4 %; n = 9; Figure

4-1 G). Examination of the current-voltage relationships before and during PS application and during morphine treatment (Figure 4-1 H) demonstrated that the response to PS was greatly inhibited by morphine at all voltages studied.

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Morphine is a non-selective opioid receptor agonist and DRG neurons express all three naloxone-sensitive opioid receptor subtypes (µ, κ and δ). We therefore tested the effects of

2+ subtype selective opioid receptor agonists on PS-evoked [Ca ]i responses in DRG neurons to determine which receptor subtype is important for morphine-induced inhibition of TRPM3.

Treatment with the δ-opioid receptor agonist, SB205607 (20 nM), had no effect (SB205607:

R = 52 ± 3 %; Figure 4-2 A). Application of the selective κ opioid receptor agonist, U50488 (20 nM), did not inhibit PS-evoked responses, however, it produced a modest but significant increase (U50488: R = 72 ± 3 %; control: R = 57 ± 3%, Figure 4-2 A). In contrast, application of

2+ the MOR agonist DAMGO (20 nM) significantly inhibited PS-evoked [Ca ]i-responses to a similar extent to observed morphine-mediated inhibition (DAMGO: R = 18 ± 2 %, Figure 4-2

A-C). These results indicate that morphine-mediated inhibition of TRPM3 is through MOR activation and suggests that MOR is co-expressed with TRPM3. This is in agreement with previous observations of MOR but not δ-opioid receptors are expressed in heat nociceptors

(Scherrer et al., 2009).

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Figure 4-2 Activation of μ-opioid receptors inhibits TRPM3

A Effect of the selective μ opioid receptor agonist DAMGO, δ opioid receptor agonist SB205607 and κ opioid

2+ receptor agonist U50488 (each at 20 nM) on [Ca ]i-responses evoked by PS (20 µM, second application, see Figure 4-1 for protocol). *p<0.05, ***p<0.001; Kruskal-Wallis (control, n = 152; DAMGO, n = 184; SB205607,

2+ n = 200; U50488, n = 140). B Traces displaying the effect of DAMGO (20 nM) on neuronal [Ca ]i responses to stimulation with PS (20 µM). F(340/380) indicates fura-2 emission ratio. C Pseudocolour images illustrating the change in F340/380 ratio in response to PS and KCl with the colours corresponding to various F340/380 values indicated by the bar.

Since activation of the Gαi/o-coupled MOR inhibited TRPM3, we next examined whether other

Gi/o-coupled GPCRs also can inhibit native TRPM3 channels in isolated DRG neurons. The metabotropic Gi/o-coupled GABAB1 and GABAB2 have been found to be highly expressed in peripheral sensory neurons (60-90%; (Charles et al., 2001; Cuny et al., 2012; Engle et al., 2012)

125 where activation of GABAB1 was shown to modulate activity of TRPV1 channels (Hanack et al.,

2015). Application of the selective GABAB agonist baclofen (100 µM) significantly inhibited the amplitude of the second PS response to R = 12 ± 2 % compared to R = 58 ± 6 % in control

2+ experiments (Figure 4-3 A and B) and abolished (R < 5 %) the PS-evoked [Ca ]i responses in

67 % of neurons (n = 130/194).

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Figure 4-3 Activation of other Gi-coupled receptors also inhibit TRPM3.

2+ A and B Effect of (RS)-Baclofen (100 µM) on the relative [Ca ]i-response amplitude evoked by PS (20 µM). C and

2+ D Effect of PYY (100 nM) on the relative [Ca ]i-response amplitude evoked by PS (20 µM). Plots illustrate the response amplitudes evoked by the second PS challenge (% of first PS amplitude) in control experiments and experiments where neurons were perfused with B (RS)-Baclofen (control, n = 138; Baclofen, n = 194) and D PYY

2+ (control, n = 289; PYY, n = 217). ***p<0.001; Mann-Whitney U test E Traces displaying [Ca ]i responses to two sequential 5 µM PS challenges, followed by a 50 µM PS challenge and depolarisation with high K+ (50 mM KCl).

Cells were exposed to 10 µM BIIE 0246 for 2 min before and during the second PS challenge. F-H Traces displaying

2+ DRG [Ca ]i responses to sequential 20 µM PS challenges (indicated by black bars) in the presence and absence of morphine (10 µM), baclofen (100 µM) and PYY (100 nM), followed by depolarisation with high K+ (50 mM KCl).

A group of cells were inhibited by some GPCR agonists but not others. F(340/380) indicates fura-2 emission ratio. I

Venn diagram illustrating the pattern of inhibition of PS responses by morphine, baclofen and PYY, n = 210 PS and KCl responsive cells. Responses in 31 cells were not inhibited by any of the GPCR agonists (second response amplitude >15% of first PS response amplitude).

Gi/o-coupled NPY receptors Y1 and Y2 are also expressed in 15-20 % of sensory neurons

(Brumovsky et al., 2005; Ji et al., 1994; Taylor et al., 2014; Zhang et al., 1997, 1994). Y1 receptors are mainly expressed in small diameter neurons whereas the Y2 receptor is predominantly expressed in medium and large diameter neurons. We therefore examined whether activation of NPY receptors by the agonist peptide YY (PYY) can also inhibit neuronal

TRPM3-mediated responses. Similar to the effect of morphine and baclofen application,

2+ treatment with PYY (100 nM) inhibited PS-evoked [Ca ]i-responses where it abolished the evoked increase in 57 % (n = 123/217, R < 5 %) of neurons. Following PYY treatment, the relative amplitude of the second PS response was reduced from 66 ± 3 % in control experiments to 11 ± 1 % Figure 4-3 C and D). Like MOR, Y2 receptors have been reported to

128 possess constitutive activity (Chen et al., 2000) and we therefore examined whether tonic Y2 receptor activity inhibits TRPM3 channel activity. Similar to the experiments conducted with naloxone-mediated potentiation of PS-evoked (5 µM) responses, treatment with the Y2 receptor inverse agonist BIIE 0246 (10 µM) significantly (p < 0.05, Fisher’s) increased the number of responding neurons from 1.1 % (n = 5/450) to 4.2 % (n = 19/450; Figure 4-3 E).

Conversely, the number of PS (5 µM) responders in control cells fell from 3.4 % (n = 15/440) to 1.8 % (n = 8/440). These results indicate that constitutive activity of Y2 receptors can tonically inhibit TRPM3.

Next, we examined whether opioids, baclofen and PYY target distinct populations of TRPM3- expressing neurons. Isolated DRG neurons were sequentially treated with morphine (10 µM), baclofen (100 µM) and PYY (100 nM) and challenged with PS (20 µM) in the presence and absence of these compounds (Figure 4-3 F-I). A high proportion of PS-sensitive neurons (43

%, n = 91/210; Figure 4-I) were inhibited by all three agonists (R < 15 %). A small percentage of neurons was inhibited by morphine and baclofen but not PYY (7 %, n = 14/210) and larger percentage of neurons was inhibited by morphine and PYY but not baclofen (13 %, n =

27/210). Additionally, baclofen and PYY but not morphine inhibited 10 % (n = 21/210) of neurons. Small populations of neurons were inhibited by application of only one agonist

(morphine: 7%, n = 15/210; baclofen: 4%, n = 9/210; PYY, 4%, n = 9/210) and a subpopulation of neurons were not inhibited by any of these compounds (15 %, n = 31/210). These results suggest that morphine, baclofen and PYY target overlapping populations of TRPM3- expressing neurons.

Not all Gi/o-coupled GPCRs are able to effectively inhibit TRPM3 activity. Activation of group

III metabotropic glutamate receptors (mGLUR4/6/7/8), which are present in DRG neurons

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(Carlton and Hargett, 2007; Govea et al., 2012), by L-AP4 (5 µM) had no effect on PS-evoked responses (R = 69 ± 3 %, n = 100) when compared to control (R = 61 ± 7 %, n =20). Cannabinoid

CB1 receptors are also expressed in a subset of DRG neurons (Agarwal et al., 2007; Veress et al., 2013) where application of the CB1 receptor agonist, WIN 55212-2 (1 µM) slightly but significantly inhibited the amplitude of PS-mediated responses (R = 51 ± 3 %; Figure 4-4 A) when compared to control (R = 62 ± 2 %). This effect suggests that activation of WIN 55212-2 inhibited PS-evoked responses in some but not in all DRG neurons. This was confirmed when

DRG neurons were challenged with 3 applications of PS (20 µM) with WIN 552212-2 (1 µM) present prior to and during the second application and WIN 552212-2 (1 µM) + AM251 (0.5

µM), a CB1 receptor antagonist, present during the third PS application (Figure 4-4 B). This protocol allowed for the detection of neurons whose PS-evoked responses were inhibited by

WIN 55212-2 and were restored when AM251 was co-administered with WIN 55212-2. WIN

55212-2 did not have an inhibitory effect in most neurons, however a small proportion of neurons (5 %, n = 8/156) demonstrated an AM251-reversible WIN 552212-2 inhibition of PS- mediated responses (Figure 4-4 B).

To examine this minor effect of CB1 receptor activation on DRG TRPM3 PS-mediated responses, we repeated the experiments in CHO cells expressing TRPM3 and transiently transfected with the CB1 receptor. Since not all cells were transfected with CB1 receptors, we compared the population responses to two consecutive PS applications. PS responses during the two PS applications (20 µM) were relatively stable in control cells not transfected with

CB1, however, WIN 552212-2 (1 µM) application inhibited the PS response in a sub-population of cells transfected with CB1. Additionally, the second PS response in CB1-transfected cells was altered by AM251 (Figure 4-4 C and D). These findings suggest that CB1 receptors can regulate TRPM3, however, there is little co-expression of TRPM3 and CB1 in DRG neurons.

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Figure 4-4 CB1 activation can inhibit TRPM3 channels

2+ A Effect of CB1 receptor agonist WIN 55212–2 (1 µM) on DRG [Ca ]i-responses evoked by PS (20 µM, second application). ***p<0.001; Mann-Whitney U test (control, n = 213; WIN 55212–2, n = 218). B Traces showing DRG

2+ [Ca ]i responses to three PS (20 µM) challenges in the absence and presence of WIN 55212–2 (1 µM) and WIN

55212–2 (1 µM) plus AM251 (0.5 µM) followed by high K+ (50 mM KCl). Many PS responses were unaffected by

WIN 55212–2 (upper panel) and some showed an inhibition that was reversed by co-application of the

2+ antagonist AM251 with WIN 55212–2 (bottom panel). C Traces displaying [Ca ]i responses to sequential 20 µM

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PS challenges (indicated by black bars) in CHO cells co-expressing TRPM3 and CB1 receptors in the absence (top) and presence (middle) of 1 µM WIN 552212–2 during the second PS challenge. Middle traces show cells where

2+ WIN 552212–2 inhibited the PS responses. Bottom: [Ca ]i responses to sequential applications of 5 µM PS with

2+ 0.5 µM present during the second PS application. D Plots showing the change in [Ca ]i response amplitudes

(second – first response). Note the increased number of negative values (inhibition) and the increased number of positive values (potentiation) in the presence of the agonist WIN 552212–2 and antagonist AM251, respectively.

Next, we investigated whether morphine is able to affect other TRP channel activity in DRG neurons by examining its effect on TRPV1 responses to capsaicin. Since capsaicin-mediated responses readily desensitise, we performed the experiments in the presence of cyclosporin

(1 µM) which decreases TRPV1 desensitisation (Docherty et al., 1996). DRG neurons were stimulated with multiple applications of capsaicin (30 nM) with morphine applied prior to and during the second capsaicin application (Figure 4-5 A). Morphine did not significantly inhibit capsaicin-mediated responses (Figure 4-5 B) in contrast to the strong inhibition observed in

PS-mediated responses (Figure 4-5 C).

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2+ Figure 4-5 Morphine does not inhibit capsaicin evoked [Ca ]i-responses

Cyclosporin (1µM) included in all solutions to reduce TRPV1 desensitization. A Traces showing DRG

2+ + [Ca ]i responses to three capsaicin (1 µM) challenges (indicated by black bars) followed by high K (50 mM KCl).

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Morphine (10 µM was present) before and during the second capsaicin challenge. Lower panels are pseudocolour images illustrating the change in F340/380 ratio in response to capsaicin and KCl. B Plots showing

2+ the change in [Ca ]i response amplitudes (second – first response) under control conditions (left, Caps/Caps)

2+ and when morphine was applied (middle, Caps/Mor + Caps). C 10 µM morphine reduced [Ca ]i responses evoked by 20 µM PS in same experiment. Amplitudes of responses evoked by first PS application (left) and second application of PS in presence of morphine (right). ***p<0.001

4.3.1.2 Inhibition of TRPM3 is independent of cAMP and does not rely on Gαi proteins

Activated Gαi subunits inhibit adenylate cyclase which decreases the production of cAMP and in turn reduces PKA activity. Several studies have characterised PKA-mediated regulation of ion channel function. Therefore, we determined whether opioid-mediated inhibition of

TRPM3 is caused by the reduction in cAMP levels. We examined this by testing the inhibitory effect of morphine application on PS-mediated responses in the presence of the membrane- permeable cAMP analogue, 8-bromo cAMP. 8-bromo cAMP had no effect on morphine- induced inhibition of PS-mediated responses (R = 11 ± 2 %; Figure 4-6 A and B) when compared to the inhibition in control experiments with only PS and morphine application (R

= 13 ± 2 %).

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Figure 4-6 Opioid-mediated inhibition of TRPM3 is independent of cAMP and Gαi subunits.

2+ A Traces showing the effect of 8-bromo cAMP (1 mM) on morphine (10 µM) induced inhibition of [Ca ]i– responses evoked by PS (20 µM) in DRG neurons. F(340/380) indicates fura-2 emission ratio. B Box and whisker and scatter plots displaying the average response amplitudes from experiments such as A., control, n = 96; morphine, n = 95; morphine +8 bromo cAMP, n = 118) ***p<0.001, compared to control. C Whole cell recordings illustrating morphine inhibition of PS-evoked outward membrane currents in CHO cells co-expressing TRPM3 and the µ opioid receptor (+60 mV) in the absence (left) and presence (right) of 100 µM NF023 in the pipette solution. D Box and whisker and scatter plots showing the percentage inhibition of 50 µM PS-evoked currents in the absence and presence of NF023. No significant difference noted (p=0.1).

Gα subunits can also directly act on ion channels, as has been demonstrated for Gαo-mediated inhibition of TRPM1 channels in retinal bipolar neurons (Koike et al., 2010; Xu et al., 2016). To

135 investigate whether MOR-mediated inhibition of TRPM3 is through Gαi subunits, we examined the effect of the selective Gαi inhibitor NF023 (Freissmuth et al., 1996) on morphine-induced inhibition of PS responses. NF023 has been reported to inhibit Gαi- mediated signalling when applied extracellularly to cells (Sarwar et al., 2015), however, this charged and relatively large compound is likely to exhibit limited membrane permeability.

Therefore, we examined its effect by supplying it intracellularly through the patch-pipette.

Inclusion of NF023 (100 µM) in the patch pipette in TRPM3/MOR expressing CHO cells had no effect on the inhibitory actions of morphine (Figure 4-6 C-E). These results demonstrate that opioid-induced inhibition of TRPM3 does not rely on a Gαi- and cAMP-mechanism.

4.3.1.3 Beta-gamma subunits of Gi proteins mediate inhibition of TRPM3

GPCR activation leads to the release of Gβγ subunits which can act as effector molecules and bind directly to ion channels modulating their function (Smrcka, 2008). We therefore investigated the effects of morphine, baclofen and PYY on PS evoked responses in the presence of the Gβγ inhibitor gallein in isolated DRG neurons. Treatment with gallein (20 µM)

2+ strongly prevented morphine-induced inhibition of PS evoked [Ca ]i-responses (Figure 4-7).

The response amplitude evoked by the second PS challenge was 109 ± 5 % in the combined presence of morphine and gallein compared to 10 ± 1 % in the presence of morphine alone.

Further experiments with three PS applications demonstrated a group of neurons with reduced responses (<30 % of the first response) to PS when morphine (10 µM) was applied but regained sensitivity (>50 % of first response) when gallein (20 µM) was also present

(Figure 4-7 C and D).

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Figure 4-7 βγ subunits mediate Gi/o inhibition of TRPM3.

A and B Effect of the Gβγ inhibitor gallein (20 µM) on morphine (10 µM) induced inhibition of PS-evoked [Ca2+]i- responses. A Plots show the relative response amplitudes evoked by a second PS challenge (20 µM) in control conditions, in the presence of morphine and gallein, control n = 52; morphine n = 314; morphine + gallein n = 153. B Representative traces demonstrating that gallein reverses morphine inhibition of PS (20 µM). Cells were perfused with 10 µM morphine 2 min before and during the second and third PS challenge and perfused with 20 µM gallein 7 min before and during the third PS challenge. The results shown are for morphine sensitive cells (PS response amplitude in presence of morphine < 30% of the first response amplitude). ***p<0.001

137 compared to control, †††p<0.001 compared to morphine, Kruskal Wallis. C Scatter and line plot showing the relationship between maximum response amplitudes (Δ Fura-2 ratio) for the first, second and third PS responses in control experiments without gallein and D in experiments with gallein (20 µM) present during the third PS application. E and F Plots displaying the relative response amplitudes evoked by the second (20 µM) PS challenge for control experiments and experiments where neurons were perfused with E 100 µM baclofen or 10 µM gallein and 100 µM baclofen (control, n = 88 baclofen, n = 236; baclofen and gallein, n = 102). ***p<0.001 compared to control, †††p<0.001 compared to baclofen, Kruskal Wallis. F Effect of 100 nM PYY or 10 µM gallein and 100 nM

2+ PYY on the relative amplitude of PS evoked [Ca ]i-responses (control, n = 65; PYY, n = 288; PYY and gallein, n = 183). ***p<0.001 compared to control, ††p=0.01 compared to PYY. Kruskal-Wallis. F(340/380) indicates fura-2 emission ratio.

We thereafter investigated whether gallein also prevents baclofen-induced inhibition of PS-

2+ evoked [Ca ]i-responses. Application of both gallein (10 µM) and baclofen (100 µM) during the second PS application produced a maximum response amplitude that was significantly higher than when baclofen was present alone (gallein and baclofen 38 ± 4 %, baclofen: 12 ± 2

2+ %; Figure 4-7 E). Similarly, the inhibitory effect of PYY (100 nM) on PS-evoked [Ca ]i-responses was attenuated in the presence of gallein (20 µM) where the maximum response amplitude in the presence of both gallein and PYY was significantly higher than the maximum response amplitude in the presence of PYY alone (PYY + gallein: 36 ± 3 %, PYY: 21 ± 2%; Figure 4-7 F).

Interestingly, the scatter plots suggest that the inhibitory effect of gallein on baclofen- and

PYY-mediated inhibition only occurred in a subset of neurons, unlike the strong effect observed in morphine inhibition in almost all cells. One reason for this difference may be due to NPY and GABAB receptors being in closer proximity to TRPM3 channels. Nevertheless, these results indicate that TRPM3 activity can be regulated by a Gβγ-dependent mechanism following activation of MOR, GABAB and NPY receptors.

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To determine whether TRPM3 can be inhibited directly by Gβγ subunits in the absence of

GPCR stimulation and Gαi/o activation, we examined the effect of intracellularly dialysing

TRPM3 expressing HEK293 cells with Gβγ subunits (50 nM) electrophysiologically. Application of PS (50 µM) for 10 s every minute evoked large outward currents in TRPM3 HEK293 cells, without any detectable desensitization (Figure 4-8 A and B). In contrast, intracellular dialysis with Gβγ-subunits (50 nM) produced a significant and progressive inhibition of the PS-evoked current amplitude (Figure 4-8 A and B). To confirm the inhibitory effects of Gβγ subunits on

TRPM3 we applied Gβγ subunits to the intracellular face of inside-out membrane patches from TRPM3 HEK293 cells stimulated with another, specific TRPM3 agonist, CIM0216.

Macroscopic CIM0216-evoked currents in these patches were greatly inhibited by 50 nM Gβγ subunits (Figure 4-8 C and E) but not by administration of Gβγ subunits that had been heat denatured by boiling (Figure 4-8 D and E). These results are consistent with direct inhibition of TRPM3 by Gβγ subunits.

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Figure 4-8 Effect of Gβγ subunits on PS-evoked currents.

A Outward currents (+40 mV) evoked by 10 s PS (50 µM) applications every minute in TRPM3-expressing HEK293 cells. Cells dialysed intracellularly with Gβγ subunits (50 nM, red trace) showed a progressive decline in the evoked current amplitude compared to control cells. B Column graph displaying normalised current (% of maximum current) for PS-evoked outward currents in TRPM3-expressing HEK293 cells. Columns represent mean ± SEM; **p<0.01, ***p<0.001, unpaired t-test (control, n = 3–5 cells; Gβγ, n = 3–5 cells). C Gβγ subunits

(50 nM) applied to the intracellular face of an inside out membrane patch from TRPM3-expressing HEK293 cell inhibit 10 µM CIM0216 evoked outward current (+60 mV). D Heat inactivated (100°C, 10 min) Gβγ subunits (50 nM) do not inhibit CIM0216 evoked currents. E Plots of percentage inhibition of CIM0216 evoked currents in inside out membrane patches for Gβγ subunits (n = 5) and boiled Gβγ subunits (n = 6).** p=0.002.

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4.3.1.4 Gi/o-coupled GPCRs modulate TRPM3-mediated nociceptive responses

We determined the effects of TRPM3 modulation in vivo by examining the nociceptive responses evoked by TRPM3 agonists in wild-type mice. Previous studies in TRPM3 knockout mice have shown that the behavioural responses to intraplantar injection of PS and CIM0216, are dependent on TRPM3 (Held et al., 2015a). The behavioural responses to either agonist alone were often mild. We therefore administered a combination of 5 nmole PS and 0.5 nmole CIM-0216 as these compounds have been shown to act synergistically in vitro (Held et al., 2015a). Intraplantar administration of the combined agonists evoked robust paw licking/flinching behaviour that was measured over a 2-min period. Prior intraplantar administration of morphine (130 nmole) essentially abolished the behavioural response evoked by PS/CIM0216 (Figure 4-9 A), and intraplantar injections of either baclofen

(240nmole) or PYY (235pmole) also significantly reduced the response (Figure 4-9 B).

Behavioural responses could be inhibited by morphine acting on, for example, voltage gated potassium channels in the periphery. We therefore examined the effects of intraplantar morphine on the paw licking/flinching responses evoked by intraplantar injection of capsaicin.

Morphine had no inhibitory effect on capsaicin-elicited behaviours (Figure 4-9 E) in marked contrast to its effect on PS/CIM0216 responses.

Naloxone and BIIE0246 act as inverse agonists at MOR and NPY Y2 receptors (Elbrønd-Bek et al., 2015; Wang et al., 2001) and consequently inhibit constitutive GPCR activity as well as inhibiting agonist evoked responses. As these ligands augmented TRPM3 activity in DRG neurons in vitro, we examined if they would increase the nociceptive effects of TRPM3 agonists. For these studies we used intraplantar injection of PS alone, which evoked relatively mild behavioural responses. Prior intraperitoneal administration of either 2.5 mg/kg naloxone

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(Figure 4-9 C) or 3 mg/kg BIIE0246 (Figure 4-9 D) resulted in significantly increased nociceptive responses to PS.

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Figure 4-9 Nociceptive responses to TRPM3 agonists are modulated by Gi/o GPCR ligands.

A and B Inhibitory effects of prior (5 min) intraplantar administrations of A 130 nmole morphine (n = 6 for each group) and B 240 nmole baclofen (n = 6) or 235 pmole PYY (n = 8) on the duration of licking/flinching evoked by intraplantar injection of 5 nmole PS plus 0.5 nmole CIM-0216 (n = 10 for PS + CIM0216). C and D Effects of prior

(30 min) intraperitoneal administration of either (c) 2.5 mg/kg naloxone (n = 9 for PS, n = 8 for PS + naloxone) or

(d) 3 mg/kg BIIE 0246 (n = 11 for PS, n = 8 for PS + BIIE 0246) on licking/flinching behaviour evoked by intraplantar administration of 5nmole PS. *p<0.05, **p<0.01, ***p<0.001; ANOVA followed by Tukey’s HSD test. E No effect of intraplantar morphine (130 nmole) on nociceptive responses evoked by intraplantar administration of capsaicin (33 nmole). Vehicle, 0.9% NaCl.

4.3.1.5 Adenosine inhibits TRPM3 in sensory neurons and in vivo through Gi/o-coupled A1 receptor activation

The Gi/o-coupled adenosine A1 receptor is expressed in DRG neurons and the dorsal spinal cord where its activation has been shown to promote antinociception in acute and chronic pain models (Gao et al., 2014; Gong et al., 2010; Schaddelee et al., 2005) We therefore examined whether adenosine A1 receptor activation can also inhibit TRPM3 channels in DRG neurons. We applied PS continuously in DRG neurons, which produced sustained increases in

2+ [Ca ]i (Figure 4-10 A). In control experiments, the amplitude of PS-evoked responses after 9 min of application was 82.2 ± 1.2% of the maximal response amplitude, consistent with a small degree of desensitization (Figure 4-10 A). Application of adenosine (50 µM) strongly and reversibly inhibited TRPM3-mediated responses (56.6 ± 4.4 %; Figure 4-10 B and C). This inhibition was fully reversible when DRG neurons were incubate with PTX overnight (200 ng/ml, 18 hrs; 90.6 ± 2.6 %; Figure 4-10 C). These results indicate that adenosine inhibits

TRPM3 through a Gi/o-coupled GPCR. Adenosine A1 and A3 receptors both couple to Gαi/o subunits, we therefore investigated which was responsible for adenosine-mediated inhibition

143 of TRPM3. Application of the A1 agonist, ENBA (1 µM), but not the A3 agonist, HEMADO (1

2+ µM), strongly inhibited PS-induced [Ca ]i-responses in DRG neurons (Figure 4-10 D-F).

Therefore, adenosine inhibits TRPM3 activity through activating the adenosine A1 receptor.

To further confirm adenosine-mediated inhibition of TRPM3, we used voltage-clamp recordings where adenosine (50 µM) strongly and reversibly inhibited PS-induced (50 µM) currents in isolated DRG neurons (by 78.8 ± 22.2 %, n = 4; Figure 4-10 G). As activation of adenosine A1 receptor inhibited TRPM3 channels in isolated DRG neurons, we next examined whether activation of the receptor can inhibit pain evoked by TRPM3 agonists in vivo.

Intraplantar administration of a combination of 5 nmole PS and 0.5 nmole CIM0216 evoked a robust paw licking/flinching behaviour in wild-type mice (Figure 4-10 H). Intraplantar administration of 75 nmole of adenosine 5 min before PS/CIM0216 administration completely inhibited pain-related paw licking and flicking behavioural responses evoked by the TRPM3 agonists (vehicle: 20.3 ± 3.6 s and adenosine: 0.43 ± 0.43s; Figure 4-10 H). When conducting the nociception behavioural experiment using morphine and capsaicin co-administration

(Figure4-9), we did not measure the duration of pain responses, but instead counted the number of nocifensive responses. This was due to the substantial number of behaviours and their brevity which would provide less accurate measurements than recording the number of pain responses. We therefore lowered the capsaicin concentration from 33 nmole to 5 nmole which led to less severe pain responses in mice that allowed us to measure the duration of pain responses more accurately. Nevertheless, Dembla et al., also demonstrated that administration of DAMGO significantly decreased the duration of pain responses following PS but not capsaicin administration (Dembla et al., 2017). Adenosine administration had no effect on capsaicin-evoked (5 nmole) behaviours (vehicle: 73.5 ± 5.1 s and adenosine: 54.5 ±

7 s; Figure 4-10 I). Therefore, adenosine A1 receptors are another example of Gi/o-coupled

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GPCRs capable of inhibiting TRPM3 channels in sensory neurons and in vivo providing a mechanism of action for how adenosine A1 receptor activation produces antinociception.

Figure 4-10 Adenosine A1 receptors regulate TRPM3 activity

2+ A [Ca ]i responses in isolated mouse DRG neurons challenged by PS (20 µM) followed by KCl (50 mM)

2+ application. B [Ca ]i responses in isolated mouse DRG neurons treated with PS (20 µM) and adenosine (50 µM).

C Scatter plot representing the relative amplitude in A (n = 54), B (n = 53) and adenosine + PTX (200 ng/ml; n =

2+ 56) treated DRG neurons. D and E [Ca ]i responses in isolated mouse DRG neurons treated with PS (20 µM) and

ENBA (1 µM) or HEMADO (1 µM). F Scatter plot representing the relative amplitude in control (n = 87), D (n =

173) and E (n = 113). G Example trace of adenosine (50 µM)-mediated inhibition of PS (50 µM)-evoked currents in whole-cell voltage clamp recordings (+60 mV) in isolated DRG neurons. H Scatter plot representing the duration of pain responses in mice administered with intraplantar hind paw injections of PS/CIM0216 and

145 vehicle (n = 7) or adenosine (n = 7) ***p<0.001 two-tailed t-test. I Scatter plot representing the duration of pain responses in mice administered with intraplantar hind paw injections of capscaicin and vehicle (n = 6) or adenosine (n = 6).

4.3.2 Gq- and Gs-coupled GPCR regulation of TRPM3

As demonstrated above, TRPM3 is inhibited following activation of Gi/o-coupled GPCRs

(Badheka et al., 2017; Dembla et al., 2017; Quallo et al., 2017) and can also be inhibited by activation of heterologously expressed Gq-coupled GPCRs in recombinant systems (Badheka et al., 2017). This inhibition is independent of Gαi/o and Gαq subunits and is due to the direct interaction of Gβγ subunits with TRPM3 (Badheka et al., 2017; Dembla et al., 2017). Earlier studies of some voltage gated calcium (VGCCs) and G protein-coupled inwardly-rectifying potassium (GIRK) channels have shown that these can be promiscuously modulated by Gβγ released from different Gα subunits (see (Dascal, 1997; Dolphin, 2003; Yamada et al., 1998) for review). We therefore examined whether activation of Gs-coupled GPCRs can similarly modulate TRPM3 through liberated Gβγ subunits. Additionally, we assessed whether Gq- coupled GPCR activation modulates TRPM3 activity in native sensory neurons and in vivo.

4.3.2.1 Non Gi/o mediated inhibition of TRPM3

Since Gi/o-coupled GPCRs inhibit TRPM3 by an interaction with Gβγ (Badheka et al., 2017;

Dembla et al., 2017; Quallo et al., 2017), we examined whether Gβγ subunits liberated by other Gα proteins can exert a similar influence on TRPM3. To assess the effects of the irreversible Gα activator GTPγS and inhibitor GDPβS we studied HEK293 cells stably expressing TRPM3 by voltage-clamp. In the absence of either GTP analogue, currents evoked

146 by repeated application of PS (100 µM) showed a time-dependent, but modest decline in amplitude (Figure 4-11 A and D). Inclusion of GTPγS (300 µM) in the pipette solution significantly accelerated the desensitisation of TRPM3 after about 3 min (3rd PS application)

(Figure 4-11 B and D). In contrast, dialysing the cell with GDPβS (500 µM) lead to a progressive and significant increase in the PS-evoked current amplitude, which reached a maximum after about 4 min (4th PS application; Figure 4-11 C and D). To test whether the effects of GTPγS were partly mediated by non-Gi/o signalling, we incubated the cells with pertussis toxin (PTX,

200 ng/ml, for 24 hrs), which specifically locks the αi/o subunits into an inactive, GDP-bound state and inhibits Gαi/o subunits coupling to their cognate GPCRs. GTPγS was still able to inhibit PS-evoked currents in PTX-treated cells, albeit more slowly than in the absence of PTX

th (10 PS application) (Figure 4-11 E-G), which is consistent with an action independent of Gi/o signalling.

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Figure 4-11 GPCR-mediated regulation of TRPM3 activity in HEK293 cells.

A-C Example traces of multiple applications of PS (100 µM) in whole-cell voltage clamp recordings (+60 mV) of

TRPM3 HEK293 cells with control, GTPγS (300 µM) or GDPβS (500 µM) intracellular solutions. D Bar chart of the average responses ± SEM of A (n = 7), B (n = 6) and C (n = 8). E and F Example traces of multiple applications of

PS (100 µM) in whole-cell voltage clamp recordings (+60mV) of TRPM3 HEK293 cells treated with PTX (200ng/mL) with control or GTPγS (300 µM) intracellular solutions. G Bar chart of the average responses ± SEM of E (n = 7-

8) and F (n = 9-10). *p<0.05 and **p<0.01 compared to control, repeated measures two-way ANOVA with

Dunnett’s multiple comparisons test D and multiple t-test G.

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4.3.2.2 Gs-mediated inhibition of TRPM3

Since our experiments with GTPγS demonstrate that G proteins other than Gi/o can inhibit

TRPM3 currents, we examined the effects of agonists of GPCRs coupled to Gαs on TRPM3 activity. The adenosine A2B receptor is expressed at high levels in HEK293 cells (Cooper et al.,

1997) and couples to Gαs, thereby substantially increasing intracellular [cAMP] upon stimulation. To confirm that the receptor is expressed in our cell line, we used HEK293 cells stably expressing the pGLOSensor (Binkowski et al., 2009) to monitor the effect of A2B agonists on cAMP levels. Application of the endogenous full agonist adenosine and the selective partial A2B agonist BAY 60-6583 (Hinz et al., 2014) produced robust intracellular cAMP responses (Figure 4-12 A). The response profile and pEC50 values of adenosine (4.75 ±

0.05 SEM, n = 4) and BAY 60-6583 (6.04 ± 0.18 SEM, n = 4) agree with previous studies

(Goulding et al., 2018). To confirm that both agonists produced cAMP by activating the A2B receptor, we constructed inhibitory concentration response curves using the selective A2B antagonist PSB603 and sub-maximally effective (EC80) concentrations of adenosine (50 µM) and BAY 60-6583 (5 µM). PSB603 inhibited the responses with a similar pIC50 for both adenosine (7.18 ± 0.15 SEM, n = 4) and BAY 60-6583 (6.71 ± 0.3 SEM, n = 4) and almost completely inhibited intracellular cAMP responses at the highest concentrations tested

(Figure 4-12 B). To assess whether A2B can also couple to Gαq, we applied adenosine and used

2+ 2+ fura-2 to measure [Ca ]i (Figure 4-12 C). As a positive control for Gq-mediated [Ca ]i- increases, we used the selective PAR2 agonist, 2-Furyl LIGRLO-NH2 (2F-LIGRLO), as the receptor has been shown to couple to Gαq and produce calcium responses in HEK293 cells

(Kawabata et al., 1999). Stimulation of A2B evoked large intracellular cAMP production, but

2+ no detectable increase in [Ca ]i (Figure 4-12 A and C), and we therefore used activation of the A2B receptor to explore the influence of Gs-mediated activity on TRPM3.

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2+ The TRPM3 agonist PS (20 µM) evoked [Ca ]i-increases in TRPM3-expressing HEK293 cells, which were only subject to a minor degree of desensitisation during the 9 min application period (77.6 ± 0.81% of maximal amplitude, Figure 4-12 D and G). A2B receptor stimulation with adenosine (100 µM) or BAY 60-6583 (20 µM) significantly and reversibly reduced the amplitudes of the TRPM3-mediated responses to 34.2 ± 1.1% and 64.1 ± 0.5% (Figure 4-12 E-

G). The smaller degree of inhibition produced by BAY 60-6583 is consistent with its properties as a partial agonist at A2B receptors (Hinz et al., 2014) (Figure 4-12 A and G). Next we used patch-clamp recordings to confirm that the observed A2B mediated inhibition of PS-evoked

2+ [Ca ]i-responses was associated with a corresponding inhibition of TRPM3 currents.

Adenosine (100 µM) and BAY 60-6583 (20 µM) significantly inhibited PS-evoked (100 μM) currents to a similar extent (by 41.6 ± 6.4% and 43.1 ± 5.8%) (Figure 4-12 H, I and J). Since the current inhibition achieved by A2B stimulation was variable, we repeated the experiments using TRPM3 HEK293 cells overexpressing the A2B receptor. Overexpression of A2B generated more uniform and robust inhibition of PS-evoked currents (by 87.5 ± 3.8% for adenosine and 80.6 ± 6.7% for BAY 60-6583; Figure 4-12 K, L and M). Together these observations demonstrate that Gs-coupled GPCR activation can inhibit TRPM3.

150

151

Figure 4-12 Gs-coupled adenosine 2B receptor activation inhibits TRPM3-mediated responses in HEK293 cells.

A Concentration response curves of intracellular cAMP production in response to adenosine and BAY 60-6583 application mean ± SEM in pGLOSensor HEK293 cells. B Inhibitory concentration response curves using the selective A2B antagonist, PSB603, on intracellular cAMP production in response to adenosine (50 µM) and BAY

2+ 60-6583 (5 µM) application mean ± SEM. C Concentration response curve of [Ca ]i responses in HEK293 cells

2+ following application of 2F-LIGRLO and adenosine mean ± SEM (n = 4, both). D [Ca ]i responses in TRPM3

2+ HEK293 cells with PS application (20 µM). E and F [Ca ]i responses in TRPM3 HEK293 cells treated with PS (20

µM) and adenosine (100 µM) or BAY 60-6583 (20 µM). G Scatter plot representing the mean, median and SEM of the relative amplitude from D (n = 731), E (n = 588) and F (n = 1145). The minimum response amplitude during co-application of PS and A2B agonists was compared to the maximum amplitude during PS application. H and I

Example traces of adenosine (100 µM) and BAY 60-6583 (20 µM) mediated inhibition of PS (100 µM)-evoked currents in whole-cell voltage clamp recordings (+60 mV) in TRPM3 HEK293 cells. J Scatter plot representing

TRPM3-evoked current inhibition in H (n = 12) and I (n = 9). K and L Example traces of adenosine (100 µM) and

BAY 60-6583 (20 µM) mediated inhibition of PS (100 µM)-evoked currents in whole-cell voltage clamp recordings

(+60 mV) in HEK293 cells transfected with TRPM3 and A2B. M Scatter plot representing TRPM3-evoked current inhibition in J and K (n = 4, both). ***p<0.001 compared to control, ###p<0.001 compared to adenosine treatment, Kruskal-Wallis with Dunn’s multiple comparisons test.

4.3.2.3 Gs-mediated inhibition of TRPM3 in sensory neurons

2+ We next measured [Ca ]i-responses in isolated mouse DRG neurons to confirm that the Gs- mediated inhibition of TRPM3 operates in native cells. Applications of the inflammatory mediator PGE2 (1 μM), which has been widely used to sensitise nociceptive afferents (Meves,

2+ 2006), strongly inhibited PS-induced [Ca ]i-responses in a subpopulation of DRG neurons when compared to control (>50% inhibition in 97 out of 187 cells; Figure 4-13 A and B).

Treatment with PGE2 significantly reduced the amplitude of PS responses to 51.6 ± 2.6%

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(Figure 4-13 C). The effect of PGE2 was unaffected by the overnight incubation of neurons with

PTX (200 ng/ml, 18 hrs) (Figure 4-13 C). Conversely, PTX (200 ng/ml, 18 hrs) treatment fully prevented morphine-induced Gi/o-inhibition of PS-evoked responses in DRG neurons (Figure

4-13 D-F) (Badheka et al., 2017; Dembla et al., 2017; Quallo et al., 2017). These findings demonstrate that PGE2 inhibits PS responses independently of Gi/o.

PGE2 acts on two cognate Gs-coupled E prostanoid receptors, EP2 and EP4, which are both found in DRG neurons (Cruz Duarte et al., 2012). To determine the relative contribution of these receptors to the PGE2-mediated inhibition of TRPM3, we used the selective EP2 agonist butaprost and the selective EP4 agonist CAY10598. Butaprost (1 μM), significantly inhibited

2+ PS-induced [Ca ]i-responses in neurons (>50% inhibition in 90 out of 294 cells; Figure 4-13

G), reducing the amplitude of PS-evoked responses to 71.9 ± 1.78% compared to control experiments (87.2 ± 1.38 %; Figure 4-13 H). Butaprost (1 μM) was still able to inhibit PS- evoked responses in neurons treated with PTX (200 ng/ml, 18 hrs; Figure 4-13 H). In contrast to the EP2 receptor agonist butaprost, the selective EP4 receptor agonist CAY10598 (30 nM) was without effect on PS responses in DRG neurons compared to untreated controls (Figure

4-13 I), consistent with a limited co-expression of TRPM3 and EP4 in DRG neurons.

PS is an effective TRPM3 agonist but has previously been shown to activate NMDA

(Adamusová et al., 2013) and inhibit GABAA receptors (Akk et al., 2001). Here we found that

64.2% (52 out of 81) of DRG neurons studied by voltage-clamp responded to stimulation with

PS, which agrees well with previously reported results (Vriens et al., 2011). To confirm that

PS-evoked currents in DRG neurons were mediated by TRPM3, the selective TRPM3 antagonist ononetin (10µM) (I Straub et al., 2013) was applied during PS application. Ononetin inhibited PS evoked currents completely (by 98 ± 2%) and reversibly in all treated cells (Figure

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4-13 J). We therefore conclude that TRPM3 mediates PS-evoked current responses in DRG neurons.

We then used voltage-clamp recordings to test whether PGE2 and butaprost inhibited PS- evoked TRPM3 currents in isolated DRG neurons. Application of PGE2 (1 µM) or butaprost (1

µM) reversibly inhibited PS-evoked TRPM3 currents to a similar extent (by 65.7 ± 10.6% and

50.5 ± 8.9%) (Figure 4-13 K-M). These results demonstrate that activation of Gs-coupled EP2 receptor inhibits TRPM3 in DRG neurons.

154

155

Figure 4-13 Prostaglandin EP2 receptors inhibit TRPM3-mediated responses in sensory neurons

2+ A [Ca ]i responses in isolated mouse DRG neurons challenged by PS (20 µM) followed by KCl (50 mM)

2+ application. B [Ca ]i responses in isolated mouse DRG neurons treated with PS (20 µM) and PGE2 (1 µM). C

Scatter plot representing the relative amplitude in A (n = 207), B (n = 175) and PGE2 + PTX (200 ng/ml; n = 151)

2+ treated DRG neurons. D [Ca ]i responses in isolated DRG neurons treated with PS (20 μM) and morphine (10

2+ μM). E [Ca ]i responses in PTX-treated (200 ng/ml) isolated DRG neurons challenged with PS (20 μM) and morphine (10 μM). F Scatter plot representing the relative amplitudes of control (94.3 ± 1.6% SEM, n = 164), D

2+ (48.4 ± 2.7% SEM, n = 109) and E (86 ± 2.3% SEM, n = 135) neurons. G [Ca ]i responses in isolated mouse DRG neurons treated with PS (20 µM) and butaprost (1 µM). H Scatter plot representing the relative amplitude in control (n = 212), butaprost (n = 250) and butaprost + PTX (n = 140) treated DRG neurons. I Scatter plot representing the relative amplitude in control (n = 409), PGE2 (n = 185), butaprost (n = 294) and CAY10595 (30 nM; n = 207) treated DRG neurons. J Ononetin (10 μM) mediated inhibition of PS (50 μM)-evoked currents in whole-cell voltage clamp recordings (+60 mV) in isolated DRG neurons (n = 6). K and L Example traces of PGE2 and butaprost (1 µM, both)-mediated inhibition of PS (50 µM)-evoked currents in whole-cell voltage clamp recordings (+60 mV) in isolated DRG neurons. M Scatter plot representing TRPM3-evoked current inhibition in

K (n = 6) and L (n = 7). *** p<0.001 when compared to control, ###p<0.001 when compared to morphine-treated cells Kruskal-Wallis with Dun’s multiple comparisons test.

4.3.2.4 PAR2 activation inhibits TRPM3 through promiscuous Gi/o- and Gq-mediated signalling

Stimulation of Gq-coupled GPCRs have previously been shown to inhibit TRPM3-evoked currents, but not through a direct interaction between Gαq and TRPM3 (Badheka et al., 2017;

Dembla et al., 2017). Earlier studies have also demonstrated that PIP2 regulates TRPM3 activity, and since the principle canonical transduction mechanism of Gq-coupled receptors start with the activation of PLC and subsequent PIP2 hydrolysis we set out to identify the

156 mechanisms responsible for Gq-mediated inhibition of TRPM3 in more detail. PAR2 couples to both Gαq and Gαi/o (Traynelis and Trejo, 2007) and is well-expressed in HEK293 cells

2+ (Kawabata et al., 1999). Activation of Gq-linked pathways leads in an increase in [Ca ]I, but

2+ Gi/o-mediated signalling can also produce elevate [Ca ]i by Gβγ-mediated activation of PLC

(see Hermans, 2003 for review). To determine whether Gi/o-mediated signalling contributes

2+ to PAR2-evoked [Ca ]i responses in HEK293 cells, we therefore compared responses evoked by the selective PAR2 agonist 2F-LIGRLO with and without PTX pre-treatment. PTX-treatment

2+ (200 ng/ml, 24 hrs) reduced the maximal [Ca ]i-response amplitude evoked by 2F-LIGRLO to

85.8 ± 6% SEM (n = 4) of that in untreated cells but did not influence the apparent potency of

2F-LIGRLO (untreated cells pEC50 = 6.5 ± 0.12 and PTX-treated pEC50 = 6.43 ± 0.09, n = 4 both)

2+ (Figure 4-14 A). Our results indicate that the increase in [Ca ]i produced by activating PAR2 with 2F-LIGRLO is primarily, but not exclusively caused by Gq-mediated signalling.

2+ Initially, we examined the influence of PAR2 over TRPM3 activity using [Ca ]i-imaging. Since

2+ PAR2 activation can evoke [Ca ]i -responses, we applied a low concentration of 2F-LIGRLO

2+ (100 nM) that did not produce a [Ca ]i response in control experiments (Figure 4-14 B). 2F-

2+ LIGRLO (100 nM) significantly reduced the amplitude of TRPM3-mediated [Ca ]i responses to

40.9 ± 0.4%. In contrast, responses in control cells slightly desensitised over the 9-minute PS application period to 68.7 ± 0.4% of the maximal response (Figure 4-14 C and D).

Next, we examined the effects of PAR2 activation on TRPM3 electrophysiologically.

Application of 2F-LIGRLO (10 µM) inhibited TRPM3-evoked currents substantially by 80.6 ±

6.6% (Figure 4-14 E). To determine the relative contributions of Gi/o- or Gq-mediated signalling to this inhibition, we examined the effect of PTX pre-treatment (200 ng/mL). Applications of

2F-LIGRLO (10 µM) still inhibited TRPM3 after PTX treatment, but significantly less effectively

157

2+ (29.7 ± 3.6%; Figure 4-14 F and G). These results were also observed in [Ca ]i imaging experiments (Figure 4-14 G and H). Therefore, PAR2 activation inhibits TRPM3 via both Gq- and Gi/o-mediated signalling.

Since PAR2 can signal via both Gq and Gi/o and we know that Gi/o-mediated inhibition of TRPM3 is due to Gβγ subunits, we studied Gq-mediated signalling further. Gαq activation has previously been shown to inhibit TRPM3 via the hydrolysis of PIP2 (Badheka et al., 2015; Tóth et al., 2015). To determine whether this was the case, we examined if TRPM3 inhibition could be prevented by supplying PIP2 by including diC8 PtdIns(4,5)P2 (100 µM) in the pipette solution. PAR2-mediated inhibition of TRPM3 was unaffected by dialysis with diC8

PtdIns(4,5)P2 (77.2 ± 4.9%), indicating that PtdIns(4,5)P2 hydrolysis is not necessary for Gαq inhibition of TRPM3 (Figure 4-14 J and L). To examine the importance of PtdIns(4,5)P2 hydrolysis on PAR2-mediated inhibition of TRPM3 further, TRPM3 HEK293 cells were transiently transfected with PM-FRB-CFP and RFP-FKBP12-5ptpase. Rapamycin application induces heterodimerisation of these two constructs, followed by translocation of the resultant active phosphoinositide 5-phosphatase to the plasma membrane which leads to the hydrolysis of membrane-bound PtdIns(4,5)P2 (Varnai et al., 2006). Application of rapamycin

(500 nM) strongly reduced the amplitude of TRPM3-evoked currents (47.1 ± 7.7% SEM, n = 7;

Figure 4-14 K) in good agreement with previous reports (Badheka et al., 2015; Tóth et al.,

2015). Subsequent application of 2F-LIGRLO lead to the near complete elimination of TRPM3- evoked current (96.5 ± 2%). The inhibition produced by co-application of rapamycin and 2F-

LIGRLO was significantly greater than in the absence of rapamycin (Figure 4-14 L). The significantly enhanced inhibition observed by concomitant PtdIns(4,5)P2 hydrolysis and PAR2 suggests that the PtdIns(4,5)P2 hydrolysis produced by PAR2 stimulation is not sufficient to inhibit TRPM3 completely. These results indicate that PAR2-mediated inhibition of TRPM3

158 involves both Gq and Gi/o signalling, but that it is unlikely that PtdIns(4,5)P2 hydrolysis contributes significantly.

159

Figure 4-14 PAR2 receptor activation inhibits TRPM3-mediated responses through Gi/o- and

Gq/11-mediated signalling

2+ A Concentration response curve of [Ca ]i responses in HEK293 cells following application of 2F-LIGRLO ± SEM in

2+ control and PTX-treated cells. B [Ca ]i responses in HEK293 cells with application of different 2F-LIGRLO

2+ concentrations. C [Ca ]i responses in TRPM3 HEK293 cells treated with PS (20 µM) and 2F-LIGRLO (100 nM). D

Scatter plot representing the relative amplitude from C (n = 1370) and control cells (n = 959). *** p<0.001 Mann-

Whitney u test E and F Example traces of 2F-LIGRLO (10 µM)-mediated inhibition of PS (100 µM)-evoked currents in whole-cell voltage clamp recordings (+60 mV) in control and PTX-treated (200 ng/mL) TRPM3 HEK293 cells. G

2+ Scatter plot representing TRPM3-evoked current inhibition in E (n = 10) and F (n = 9) H [Ca ]i responses in PTX- treated (200 ng/ml) TRPM3 HEK293 cells challenged with PS (20 µM) and 2F-LIGRLO (100 nM). I Scatter plot representing the relative amplitudes of control (71.9 ± 0.4%, n = 578), 2F-LIGRLO (58.1 ± 0.6% SEM, n = 649)- and 2F-LIGRLO + PTX (65.5 ± 0.5% SEM, n = 712)-treated TRPM3 HEK293 cells. ***p<0.001 when compared to control, ###p<0.001 when compared to 2F-LIGRLO-treated cells, Kruskal-Wallis with Dun’s multiple comparisons test. J Example trace of 2F-LIGRLO (10 µM)-mediated inhibition of PS (100 µM)-evoked currents in TRPM3

HEK293 cells supplemented with diC8 PtdIns(4,5)P2 (100 µM) in the intracellular solution. K Example trace of 2F-

LIGRLO (10 µM)- and rapamycin (500 nM)-mediated inhibition of TRPM3 HEK293 cells transfected with PM-FRB-

CFP and RFP-FKBP12-5ptpase enzyme. L Scatter plot representing TRPM3-evoked current inhibition in E (n = 9),

J (n = 8) and K (n = 7). *p<0.05 when compared to control, one-way ANOVA followed by Dun’s multiple comparisons test I.

4.3.2.5 Activation of endogenous Gq-coupled purinergic receptors Y inhibit TRPM3 activity in HEK293 cells

Since activation of endogenous PAR2 receptors inhibits TRPM3 channels through promiscuous coupling to Gαi/o and Gαq, we investigated whether a solely Gq-coupled receptor can also inhibit TRPM3. Several reports suggest that various purinergic Y (P2Y) receptors that couple to Gαq are endogenously expressed in HEK293 cells. To determine whether any Gq-

160 coupled P2Y receptors are present in our cell line, we produced a concentration-response

2+ curve with the purinergic receptor agonist ATP. Application of ATP produced robust [Ca ]i- responses with an pEC50 value of 5.36 ± 0.05 (n = 6, Figure 4-15 A). Furthermore, pre-

2+ treatment with PTX (200 ng/ml, 24 hrs) had no effect on ATP-induced [Ca ]i-responses. We

2+ next examined the influence of ATP application on TRPM3 activity using [Ca ]i-imaging. As

2+ ATP application evokes [Ca ]i-responses we applied a low concentration of ATP (1 µM) that

2+ did not produce a [Ca ]i-response in control experiments (Figure 4- 15 B). ATP significantly

2+ reduced the amplitude of TRPM3-mediated [Ca ]i responses to 74 ± 0.32 %. In contrast, responses in control cells which desensitised slightly over the 9 -minute PS application period to 80 ± 0.48 % of the maximal response (Figure 4- 15 C and E). PTX treatment (200 ng/ml, 24 hrs) increased ATP-mediated inhibition of PS-evoked responses to 64.7 ± 0.26 % (Figure 4-15

D and E). ATP-mediated inhibition of TRPM3 was also observed in patch-clamp where application of ATP (100 µM) inhibited TRPM3-mediated currents by 37 ± 6.6 %. Dialysis of

TRPM3 HEK293 cells with bovine brain-derived PIP2 (10 µM) had no effect on observed ATP- induced inhibition of TRPM3-mediated currents (by 38.3 ± 10.6 %, Figure 4-15 F-H).

Altogether, these results demonstrate that ATP inhibits TRPM3 in HEK293 cells may predominantly be through Gq-mediated signalling in a largely PIP2-independent manner.

161

Figure 4-15 Endogenous P2Y receptor activation inhibits TRPM3-mediated responses through Gq/11-mediated signalling

2+ A Concentration response curve of [Ca ]i responses in HEK293 cells following application of ATP ± SEM in control

2+ and PTX-treated cells. B [Ca ]i responses in HEK293 cells with application of different ATP concentrations. C

2+ 2+ [Ca ]i responses in TRPM3 HEK293 cells treated with PS (20 µM) and ATP (1 µM). D Ca ]I responses in PTX- treated (200 ng/ml) TRPM3 HEK293 cells challenged with PS (20 µM) and ATP (1 µM). E Scatter plot representing the relative amplitudes of control (n = 515), ATP (n = 1221)- and ATP + PTX (n = 1187)-treated TRPM3 HEK293 cells. ***p<0.001 when compared to control, ###p<0.001 when compared to ATP-treated cells, Kruskal-Wallis with Dun’s multiple comparisons test. F and G Example traces of ATP (100 µM)-mediated inhibition of PS (100

µM)-evoked currents in whole-cell voltage clamp recordings (+60 mV) in control and bovine-brain derived PIP2

(10 µM)-treated TRPM3 HEK293 cells. H Scatter plot representing TRPM3-evoked current inhibition in F (n = 6) and G (n = 6).

162

We next attempted to determine which P2Y receptors may be responsible for ATP-induced inhibition of TRPM3. The Gq-coupled purinergic receptors Y1 is endogenously expressed in

HEK293 cells (Ecke et al., 2008; Moore et al., 2001). We therefore aimed to produce concentration response curves by using the selective P2Y1 agonists MRS2365, however,

2+ application of the agonist did not produce an increase in [Ca ]i-responses. Furthermore, inhibition of TRPM3 by ATP was relatively weak and variable, therefore, we decided to utilise overexpressed muscarinic M1 receptors.

4.3.2.6 Gq-mediated inhibition of TRPM3 channels by muscarinic M1 receptors

Activation of the Gq-coupled muscarinic acetylcholine M1 and BK2 receptors leads to the inhibition of TRPM3-evoked currents in recombinant cells (Badheka et al., 2017). Consistent with these results we found that activation of muscarinic M1 receptors inhibited TRPM3-

2+ mediated [Ca ]i-responses in HEK293 cells (Figure 4-16 A and B). M1 receptor activation can

2+ increase [Ca ]i and we therefore applied a low concentration of carbachol (0.1 µM) that did

2+ not evoke a [Ca ]i response in control experiments (Figure 4-16 C).

163

Figure 4-16 Muscarinic M1 receptor induced inhibition of TRPM3

2+ A [Ca ]i responses in TRPM3/muscarinic M1 HEK293 cells treated with PS (20 μM) and carbachol (0.1 μM) followed by application of a high concentration of carbachol (1 μM) to confirm M1 transfection. B Scatter plot representing the mean, median and SEM of the relative amplitude from control cells (60 ± 0.9% SEM, n = 431)

2+ and A (51.7 ± 1.0% SEM, n = 329), (Mann-Whitney U test), ***p<0.001. C [Ca ]i responses in TRPM3/muscarinic

M1 HEK293 cells with application of different carbachol concentrations.

164

4.3.2.7 Gq-mediated inhibition of TRPM3 in sensory neurons

We next examined whether Gq-coupled GPCRs can inhibit the activity of TRPM3 in its native environment by stimulating isolated DRG neurons with bradykinin, a pronociceptive mediator often used to sensitise nociceptors (Pethő and Reeh, 2012). Application of a maximally effective concentration of bradykinin (100 nM) strongly and rapidly inhibited PS-induced

2+ [Ca ]i-responses in a subpopulation of DRG neurons (>50% inhibition in 86 out of 293 cells)

(Figure 4-17 A). Bradykinin treatment significantly reduced the amplitude of PS responses to

65.6 ± 1.4% when compared to control experiments (78 ± 1.8%; Figure 4-17 B). The effect of bradykinin was not altered by PTX pre-treatment (200 ng/ml, 18 hrs; Figure 4-17 C).

Furthermore, pre-treatment with the selective BK2 receptor antagonist HOE 140 (50 nM) completely prevented the effect of bradykinin (control: 69.1  1.6%, bradykinin: 58.5  1.7%, bradykinin + HOE 140: 68.7  1.6%; Figure 4-17 D). In contrast, the BK1 receptor antagonist

SSR240612 (50 nM) was without effect on bradykinin-mediated inhibition of TRPM3 (control:

82.1  1.7%, bradykinin: 67  2.6%, bradykinin + SSR240612: 68.4  1.8%; Figure 4-17 E). It is important to note that GPCR co-expression with TRPM3 in DRG neurons is likely to be partial and that the means presented detect a substantial effect in a subset of neurons but that many neurons will be less affected or unaffected. Together, these results show that bradykinin inhibits TRPM3 in DRG neurons by activating BK2 receptors.

We also examined the effects of bradykinin on DRG neurons electrophysiologically.

Application of bradykinin (100 nM) to DRG neurons in the voltage-clamp configuration reversibly inhibited PS-evoked TRPM3 currents (by 53.7 ± 10.8% SEM, n = 5; Figure 4-17 F).

Therefore, activation of Gq-coupled GPCRs inhibits TRPM3 in DRG neurons.

165

Figure 4-17 Bradykinin BK2 receptor activation inhibits TRPM3-mediated responses in sensory neurons.

2+ A [Ca ]i responses in isolated mouse DRG neurons treated with PS (20 µM) and bradykinin (100 nM). B Scatter plot representing the relative amplitudes in control (n = 293) and bradykinin (n = 293)-treated DRG neurons

(Mann-Whitney U test) ***p<0.001. C Scatter plot representing the relative amplitudes in control (n = 190), bradykinin (n = 157)- and bradykinin + PTX (200 ng/ml; n = 165)-treated DRG neurons. D Scatter plot representing the relative amplitudes in control (n = 157), bradykinin (n = 197)- and bradykinin + HOE 140 (50 nM; n = 182)- treated DRG neurons. E Scatter plot representing the relative amplitudes in control (n = 118), bradykinin (n =

106) and bradykinin + SSR240612 (50 nM; n = 144)-treated DRG neurons. ***p<0.001 when compared to control,

###p<0.001 when compared to bradykinin-treated cells, and Kruskal-Wallis with Dun’s multiple comparisons test

C-E F Example trace of bradykinin (100 nM) mediated inhibition of PS (50 µM)-evoked currents in whole-cell voltage clamp recordings (+60 mV) in isolated DRG neurons.

166

4.3.2.8 Gs- and Gq-mediated inhibition of TRPM3 is independent of PKA and PKC activity

Canonical Gs- and Gq-coupled receptor signalling leads to the downstream activation of PKA and PKC. This has been previously shown to increase nociceptor excitability by sensitizing voltage-gated calcium channels (VGCCs) such as the Cav3.2 (Chemin et al., 2007; Kim et al.,

2006) and the noxious heat transduction channel TRPV1 (Bevan et al., 2014). We used the selective PKA inhibitor KT5720 to determine whether PKA contributed to the EP2-mediated

2+ inhibition of TRPM3. The butaprost-induced inhibition of PS-evoked [Ca ]i responses in DRG neurons was unaffected by KT5720 (1 µM) (control: 80.7  1.8%, butaprost: 70.8  2.6%, butaprost + KT5720: 66.7  2 %; Figure 4-18 A and B). Conversely, we used the adenylyl cyclase activator forskolin to raise intracellular cAMP and activate PKA. Forskolin (10 µM) was applied six minutes before and during PS and butaprost application. Forskolin treatment did not alter

2+ PS-induced [Ca ]i-responses in neurons and had no effect on butaprost-mediated inhibition of PS responses (control: 83.74  1.40%, butaprost: 63.82  2.15%, forskolin: 86.2  2.2%, butaprost + forskolin: 64.2  2.2%; Figure 4-18 C and D).

We examined whether PKC may be responsible for the inhibition of TRPM3 by BK2 receptor activation by applying the selective PKC inhibitor BIM VIII (Wilkinson et al., 1993) (1 µM) before and during bradykinin application. BIM VIII treatment did not prevent bradykinin-

2+ mediated inhibition of PS-induced [Ca ]i-responses in neurons but in fact increased it

(control: 78.1  1.2%, bradykinin: 59.1  1.5%, bradykinin + BIM VIII: 44.08  1.4%; Figure 4-

18 E and F). This effect may be related to an off-target effect of BIM VIII or possibly influenced by PKC inhibition preventing it from acting on other targets. Together, these observations strongly suggest that Gs- and Gq-coupled GPCRs inhibit TRPM3 independently of cAMP, PKA and PKC.

167

Figure 4-18 TRPM3 inhibition by butaprost and bradykinin is independent of PKA and PKC.

2+ A [Ca ]i responses in isolated mouse DRG neurons treated with PS (20 µM), butaprost (1 µM) and KT5720 (1

µM). B Scatter plot representing the relative amplitudes in control (n = 112), butaprost (n = 151)- and butaprost

2+ + KT5720 (n = 151)-treated cells. C [Ca ]i responses in isolated mouse DRG neurons treated with PS (20 µM), butaprost (1 µM) and forskolin (10 µM). D Scatter plot representing the relative amplitudes in control (n = 185),

2+ butaprost (n = 183)-, forskolin (n = 148)- and butaprost + forskolin (n = 155)-treated cells. E [Ca ]i responses in isolated mouse DRG neurons treated with PS (20 µM), bradykinin (100 nM) and BIM VIII (1 µM). F Scatter plot representing the relative amplitudes in control (n = 236), bradykinin (n = 257)- and bradykinin + BIM VIII (n =

248)-treated cells. **p<0.01 and ***p<0.001 when compared to control, ###p<0.001 when compared to bradykinin-treated cells, Kruskal-Wallis with Dun’s multiple comparisons test.

168

4.3.2.9 Gβγ subunits mediate inhibition of TRPM3 in heterologous and endogenous systems

Since the major canonical signalling events activated by Gαs and Gαq did not affect TRPM3 inhibition, we next examined the potential involvement of Gβγ protein. TRPM3 HEK293 cells were transiently transfected with the Gβγ sink βARK1-ct (C-terminus of the β-adrenergic receptor kinase-1) (Koch et al., 1994) and the effects of A2B activation on TRPM3 examined

2+ using [Ca ]i measurements. βARK1-ct transfection abolished the inhibitory effect of adenosine (100 µM) and BAY 60-6583 (20 µM) application on TRPM3-mediated responses when compared to untransfected cells (control: 83.7  1.40%, adenosine: 34.2 ± 1.1%, adenosine + βARK1-ct: 75.3 ± 0.8%, BAY 60-6583: 64.1 ± 0.5% and BAY 60-6583 + βARK1-ct:

80.5 ± 0.6%; Figure 4-19 A, B D and E). Similarly, TRPM3 inhibition by muscarinic M1, PAR2 and P2Y receptor activation was completely prevented by transfection with βARK1-ct (control

M1: 59.9  0.86 %, carbachol: 51.7  0.95 % and carbachol + βARK1-ct: 60.3  0.84 %; control

PAR2: 68.7 ± 0.4 %, 2F-LIGRLO: 40.9 ± 0.4 % and 2F-LIGRLO + βARK1-ct: 63.4 ± 0.4 %; control

P2Y: 83.3 ± 0.4 %, ATP: 74.9 ± 0.3 % and ATP + βARK1-ct: 84.4 ± 0.4 %; Figure 4-19 C, F, G and

H).

169

Figure 4-19 Gs- and Gq-coupled GPCR inhibition of TRPM3 is mediated by Gβγ protein.

2+ A-C [Ca ]i responses in control (upper panel) and βARK1-ct transfected (lower panel) TRPM3 HEK293 cells treated with PS (20 µM) and adenosine (100 µM, n = 852), BAY 60-6583 (20 µM, n = 553) or carbachol (0.1 µM, n = 386) . D-F Scatter plots representing the relative amplitudes of A-C with βARK1-ct untransfected control and treated TRPM3 HEK293 cells. G and H Scatter plots representing the relative amplitudes of control and βARK1- ct transfected cells treated with PS (20 µM) and 2F-LIGRLO (100 nM, control: 71.9 ± 0.4 %, n = 959; 2F-LIGRLO:

170

58.1 ± 0.6 %, n = 1370 and βARK1-ct: 65.5 ± 0.5 %, n = 1391) or PS and ATP (1 µM; control: 82.6 ± 0.4 %, n = 714;

ATP: 74.6 ± 0.3, n = 1392 and βARK1-ct: 84.3 ± 0.4 %, n = 622). ***p<0.001 when compared to control and

###p<0.001 when compared to treated cells, Kruskal-Wallis with Dun’s multiple comparisons test.

We also used GRK2i, a polypeptide that corresponds to the Gβγ binding region of G-protein- coupled receptor kinase 2 (GRK2, also known as βARK1, β-adrenoreceptor kinase 1), to further confirm the involvement of Gβγ in adenosine’s effect. GRK2i (10 µM) dialysis in TRPM3

HEK293 cell patch-clamp experiments for >5 min significantly reduced adenosine-mediated inhibition of TRPM3-evoked currents (control: 67.4 ± 6.4% and GRK2i: 36.6 ± 6.3%; Figure 4-

20 A and B). We next examined the involvement of Gβγ in EP2- and BK2-mediated inhibition of TRPM3 by dialysing GRK2i in isolated DRG neurons. GRK2i (10 µM) dialysis significantly prevented butaprost- and bradykinin-mediated inhibition of TRPM3-evoked currents (control butaprost: 50.5 ± 8.9%, GRK2i butaprost: 17.6 ±1.9%, control bradykinin: 53.7 ± 10.8% and

GRK2i bradykinin: 26.2 ± 3.2%; Figure 4-20 C-F). These results demonstrate that Gβγ subunits are responsible for TRPM3 in heterologous and native cell systems inhibition produced by activation of GPCRs coupled to Gαs and Gαq.

171

Figure 4-20 Gs- and Gq-coupled GPCR inhibition of TRPM3 in sensory neurons is reliant on

Gβγ protein.

A, C and E Example traces of adenosine (100 µM), butaprost (1 µM) and bradykinin (100 nM) mediated inhibition of PS (100 µM for HEK cells and 50 µM for DRG neurons)-evoked currents in whole-cell voltage clamp recordings

(+60 mV) in HEK293 cells (for adenosine) and isolated DRG neurons (for butaprost and bradykinin) in control

(upper panel) and GRK2i (10 µM, lower panel) intracellular solutions. B, D and F Scatter plots representing

TRPM3-evoked current inhibition in A (control n = 10, GRK2i n = 7, two-tailed t-test), C (control n = 7, GRK2i n =

5, two-tailed t-test) and E (control: n = 5, GRK2i: n = 6, two-tailed t-test) *p<0.05 and **p<0.01.

172

4.3.2.10 Gs- and Gq-coupled GPCRs inhibit TRPM3 mediated nociception

As activation of EP2 and BK2 receptors inhibits TRPM3 in isolated DRG neurons, we next attempted to determine whether proinflammatory mediators activating EP2 and BK2 receptors can inhibit pain evoked by TRPM3 agonists in vivo. Previous studies in Trpm3-/- mice showed that the behavioural nocifensive responses to intraplantar injections of PS and a second TRPM3 agonist CIM0216 are dependent on TRPM3 (Held et al., 2015a). Intraplantar administration of a combination of 5 nmole PS and 0.5 nmole CIM0216 evoked a robust paw licking/flinching behaviour in wild-type mice. Intraplantar administration of 0.3 nmole of either PGE2 (10 min before PS+ CIM0216; vehicle: 19.4 ± 3s and PGE2: 10.4 ± 0.9s), butaprost

(10 min before PS+ CIM0216; vehicle: 23.8 ± 3.8s and butaprost: 10.3 ± 4.2s) or bradykinin (5 min before PS+ CIM0216; vehicle: 13.8 ± 4.7s and bradykinin: 2.3 ± 1.4s) before PS/CIM0216 administration, significantly reduced the pain-related paw licking and flicking behavioural responses evoked by the TRPM3 agonists (Figure 4-21 A-C). The doses of PGE2, butaprost and bradykinin were chosen not to evoke pain-related behavioural responses directly.

To test whether this effect is specific for TRPM3-dependent nociception, we examined whether activation of EP2 and BK2 receptors could also inhibit TRPV1-mediated behavioural responses, since many TRPM3 expressing neurons also express TRPV1 (Vriens et al., 2011).

However, neither PGE2 (vehicle: 68.5 ± 3.7s and PGE2: 62.7 ± 3.7s), butaprost (vehicle: 77 ±

7.3s and butaprost: 72 ± 7.9s), nor bradykinin (vehicle: 57 ± 4.2s and bradykinin: 65 ± 10.7s) influenced the capsaicin-evoked behaviours (Figure 4-21 D-F), in marked contrast to their inhibitory effect on responses elicited by PS/CIM0216.

TRPM3 is required for the development and maintenance of inflammatory heat hyperalgesia, which consequently has been found to be absent from Trpm3-/- mice (Vriens et al., 2011). We

173 compared the heat sensitivity of Trpm3+/+ and Trpm3-/- mice 3 days after intraplantar injections of Freund’s complete adjuvant (FCA, 15µl). FCA reduced the paw withdrawal latency in wildtype mice in the hot-plate test (50°C) but was without effect in Trpm3-/- mice, in good agreement with earlier observations (Vriens et al., 2011), indicating that TRPM3 is of critical importance for inflammatory heat hypersensitivity (Figure 4-21 G). Administration of the selective TRPM3 inhibitor ononetin (10mg/kg, i.p.), completely reversed established FCA- induced heat hypersensitivity (Figure 4-21 H), demonstrating that the loss of hypersensitivity in Trpm3-/- mice was unlikely to be caused by developmental or compensatory mechanisms and suggests that TRPM3 may be a tractable target for inflammatory pain. FCA has long been used as a model for inflammatory hypersensitivity and pain, but the precise mechanisms by which it produces pain and hypersensitivity are not known. To determine the influence of local, intraplantar injections of PGE2 and bradykinin on established FCA-induced heat hyperalgesia, we administered the same doses that inhibited the behavioural response to

TRPM3 agonists (Figure 4-21 A and C), but that did not evoke a behavioural response on their own (5 or 10 min before hot-plate test). Perhaps counterintuitively, intraplantar administration of either PGE2 or bradykinin fully reversed heat hyperalgesia in FCA-treated mice (Figure 4-21 I). These observations are consistent with our observations that EP2 and

BK2 receptor activation inhibit TRPM3 in vitro and mimic the effects of pharmacological blockade or genetic inactivation of TRPM3 on heat hypersensitivity.

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Figure 4-21 TRPM3 dependent antinociceptive and analgesic effects of Gs- and Gq-coupled

GPCR activation prevents mouse nociceptive behaviour in response to TRPM3 agonists and reverses heat hyperalgesia in FCA-treated mice.

A-C Scatter plots representing the duration of pain responses in mice administered with intraplantar hind paw injections of PGE2 (n = 10, two-tailed t-test), butaprost (Mann-Whitney U test, p<0.05, n = 8) and bradykinin

(Mann-Whitney U test, p<0.01, n = 10) (all at a dose of 0.3 nmole) or with vehicle prior to PS (5 nmole)/CIM0216

(0.5 nmole) administration, *p<0.05, **p<0.01. D-F Scatter plots representing the duration of pain responses in mice administered with intraplantar hind paw injections of PGE2, butaprost and bradykinin (0.3 nmole, n = 6, all) or with vehicle prior to capsaicin (5 nmole) administration. G Bar chart comparing heat withdrawal latencies of mice from 50 °C hot plate pre- and post-72 hours of FCA (15 µL) injection in Trpm3+/+ (two-tailed t-test) and

Trpm3-/- mice (two-tailed t-test) (n = 8, both). H Bar chart comparing heat withdrawal latencies of mice from 50

°C hot plate pre-, post-72 hours of FCA (15 µL) injection and post-vehicle or ononetin (10 mg/kg) application (n

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= 6, both). I Bar chart comparing heat withdrawal latencies of mice from 50 °C hot plate pre-, post-72 hours of

FCA (15 µL) injection and post-treatment with vehicle, PGE2 (0.3 nmole) or bradykinin (0.3 nmole) (n = 6, each).

*p<0.05, **p<0.01 when compared to control and #p<0.05, ##p<0.01 when compared to 72 hours post FCA, two- way ANOVA followed by Tukey’s multiple comparisons test.

4.3.2.11 Identifying binding site of Gβγ protein on TRPM3 channels

Work published in a doctoral thesis from the Oberwinkler lab suggested that a short amino acid sequence on TRPM3 channels (PKALKLLGME) that is encoded by exon 17 may be the site that Gβγ proteins bind to and regulate TRPM3 channels (Mohr, 2014). This is because

TRPM3 variants α4 and α5, which lack exon 17, were not inhibited by Gβγ (Mohr, 2014). We therefore had this peptide synthesised (TRPM3 peptide) to test whether it alleviates TRPM3 inhibition following GPCR activation by sequestering Gβγ protein. Inclusion of TRPM3 peptide (10 µM) in the pipette solution had no effect on adenosine-mediated inhibition of

TRPM3 in HEK293 whole-cell patch clamp experiments (inhibition in control: 29.4 ± 7.5 % and TRPM3 peptide-treated cells: 40.6 ± 11.7 %; Figure 4-22 A and B). Therefore, this amino acid sequence, in its free form, does not bind Gβγ protein.

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Figure 4-22 TRPM3 peptide sequence does not sequester Gβγ protein

A Example traces of adenosine (100 µM)-mediated inhibition of PS (100 µM)-evoked currents in whole-cell voltage clamp recordings (+60 mV) in HEK293 cells with control (upper panel) and TRPM3 peptide (10 µM, lower panel) intracellular solutions. B Scatter plots representing TRPM3-evoked current inhibition in A (control n = 5 and TRPM3 peptide n = 5).

4.4 Discussion

4.4.1 Gi/o-mediated inhibition of TRPM3

TRPM3 is expressed in DRG neurons where it is proposed to play a role in the transduction of thermal (heat) stimuli in normal conditions and notably in the development of heat hypersensitivity in inflammatory conditions (Vriens et al., 2011). Studies of TRPM3 have been facilitated by the discovery of PS and the synthetic compound CIM0216 as TRPM3 agonists

(Held et al., 2015a; Wagner et al., 2008) that can be used to probe the functions and characteristics of TRPM3, and we have utilised these compounds to investigate the regulation of TRPM3 channels by GPCR ligands. Like other TRP channels, TRPM3 activity is regulated by intracellular PIP2 and other phosphoinositides (Badheka et al., 2015; Tóth et al., 2015) but

177 otherwise little is known about the mechanisms that regulate the activity of this channel.

Other sensory neuron TRP channels, notably TRPV1 and TRPA1, are regulated downstream of

GPCR signalling by mechanisms that involve either Gq activation, PIP2 hydrolysis and protein kinase C or Gs activation leading to protein kinase A mediated phosphorylation. These and other pathways are important regulatory mechanisms for TRP channels particularly in inflammatory conditions (see Veldhuis et al., 2015 for review). Activation of the Gi coupled µ opioid receptor is a potent analgesic and antinociceptive, with peripheral effects contributing to the predominantly central actions and opioid agonists such as morphine can inhibit TRPV1 activity by reducing phosphorylation levels; however, this is evident only when TRPV1 is sensitized via the cAMP-PKA pathway (Endres-Becker et al., 2007; Vetter et al., 2008, 2006).

This latter observation is consistent with our observation that morphine has little or no

2+ inhibitory effect on capsaicin evoked [Ca ]i responses when desensitization is blocked by cyclosporin.

Our results demonstrate for the first time that agonists acting at several Gi coupled GPCRs (µ opioid, GABA-B, NPY and A1 receptors) exert an inhibitory effect on TRPM3 activation in DRG neurons. This inhibitory effect was PTX-sensitive demonstrating a Gi/o protein involvement.

The inhibition was not, however, mediated by the canonical Gα subunit/adenylate cyclase pathway as the inhibitory effect was not abrogated by either application of the Gα subunit inhibitor NF023 or by providing a membrane permeable cAMP analogue that is well known to activate PKA. Instead, our results indicate that the TRPM3 inhibition is mediated by Gβγ subunits. The Gαi inhibitor NF023, which inhibits Gαi subunit interactions with effector molecules, was without effect when dialysed into cells at 100 µM, much higher than its reported IC50 value (~300–400 nM) (Freissmuth et al., 1996). However, Gα subunits bind to effector molecules with picomolar to nanomolar affinities and the effectiveness of NF023 will

178 depend on its relative binding affinity for Gαi compared to that of the effector molecules. We therefore cannot be certain that NF023 effectively inhibits Gαi signalling even at the concentration used. In contrast, we have clear evidence for a role for Gβγ subunits.

The inhibitory effects of morphine on TRPM3 were prevented by gallein, which is thought to be a specific inhibitor of Gβγ signalling (Lin and Smrcka, 2011), although we cannot exclude a non-Gβγ ‘off target’ effect. Critically, we found that PS evoked currents were inhibited by direct application of purified Gβγ subunits to either whole cells or excised inside-out membrane patches, without concomitant activation of Gαi or G-protein-coupled receptors.

The effect of the Gβγ subunits was lost after heat denaturation indicating that proteins in the sample were responsible for the inhibitory activity. Our findings are therefore consistent with activation of a µ opioid receptor and an action of Gβγ subunits to inhibit TRPM3. A direct interaction between the released Gβγ subunits and TRPM3 is likely as found for some other ion channels (Elbrønd-Bek et al., 2015; Veldhuis et al., 2015; Wang et al., 2001). Gβγ subunits can, however, activate other molecules such as phospholipase C which may remain in excised patches (Rebecchi and Pentyala, 2000), and such an action would hydrolyse PI(4,5)P2 and reduce TRPM3 activity (Badheka et al., 2015; Tóth et al., 2015). Inhibition of TRPM3 activity could therefore be due to a Gβγ/PLC-mediated loss of PIP2. PIP2 levels are not maintained in isolated membrane patches and decline rapidly especially in the absence of Mg-ATP, which is required for phosphoinositide kinase mediated generation of PIP2 (Zakharian et al., 2011). This loss of PI(4,5)P2, accounts for the greatly reduced TRPM3 channel activity previously reported in excised membrane patches (Badheka et al., 2015; Tóth et al., 2015). In our excised inside- out patch-clamp experiments, recordings were made minutes after excision into Mg-ATP free solution, which will deplete PIP2, and no further run down of channel activity was noted during the recordings. The finding that Gβγ subunits exerted a robust inhibitory effect on

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TRPM3 currents in membrane patches in these conditions suggests that the Gβγ inhibition does not involve PLC mediated hydrolysis of PIP2. Such a conclusion is further supported by the observations in DRG Ca2+-imaging experiments where, in contrast to TRPM3, TRPV1 responses were not affected by morphine. As the activities of both channels are modulated by PIP2 hydrolysis (Badheka et al., 2015; Cao et al., 2013), the differential, strong inhibition of

TRPM3 cannot be explained by PLC activation and a reduction in PIP2.

Direct effects of G protein subunits on ion channel functions have been well studied for P/Q- and N-type voltage gated calcium channels (Dolphin, 2003) and GIRK channels (Dascal, 1997;

Yamada et al., 1998). In contrast, there is relatively little information available about direct interactions between G protein subunits and TRP channels. TRPM1 channels are inhibited by activation of Go–linked GPCRs and the available evidence is consistent with regulation by interactions direct between TRPM1 and both Gαο and Gβγ subunits (Devi et al., 2013; Koike et al., 2010; Shen et al., 2012; Xu et al., 2016). In other studies, TRPM8 was shown to be inhibited by an interaction with Gαq (Li and Zhang, 2013; Zhang et al., 2012) while TRPA1 activation following stimulation of MrgprA3 receptors in DRG neurons was inhibited by gallein and phosducin, consistent with a Gβγ-mediated mechanism (Wilson et al., 2011). TRP channel regulation by direct interactions with G protein subunits is therefore emerging as an important and novel mechanistic concept.

DRG neurons showed differential sensitivity to activation of different GPCRs. The TRPM3 responses in many neurons were inhibited by more than one GPCR agonist consistent with expression of multiple GPCRs in these neurons. Presumably the inhibition pattern reflects the distribution and expression levels of the different GPCRs. However, not all Gi/o GPCR agonists inhibited TRPM3. The group III mGluR agonist, L-AP4, and agonists at δ- and κ-opioid receptors

180 did not significantly inhibit TRPM3 responses although they are expressed in DRG neurons

(Carlton and Hargett, 2007; Govea et al., 2012; Honsek et al., 2015; Scherrer et al., 2009; Wang et al., 2010). In part this could be explained by lack of TRPM3-GPCR co-expression in individual

DRG neurons. For example, δ-opioid receptors are expressed by about 15% of, mainly larger diameter, DRG neurons (Bardoni et al., 2014; Scherrer et al., 2009). There is also likely to be an overlap in expression of Gi/o –coupled metabotropic glutamate GPCRs (Carlton and

Hargett, 2007; Govea et al., 2012) with the small-medium diameter DRG neurons that express

TRPM3 (Vriens et al., 2011). A significant inhibition of TRPM3 responses would therefore be expected in some DRG neurons, as seen for CB1 receptor activation (Agarwal et al., 2007;

Veress et al., 2013). The effect of CB1 receptor activation in DRG neurons was restricted to a small number of neurons, although our experiments with heterologously expressed TRPM3 and CB1 indicated that CB1 receptor activity can regulate TRPM3. It is possible that specific macromolecular assemblies are required for the efficient interaction between types of GPCRs and TRPM3 and that δ- and κ-opioid and mGluR receptors do not contribute to these complexes, and that CB1 receptors are variably coupled in DRG neurons. The pronounced inhibition of TRPM3 by µ-opioid receptor activation is interesting as this receptor sub-type is expressed in mice in heat sensitive DRG neurons (Honsek et al., 2015; Scherrer et al., 2009) and may be functionally relevant for opioid control of heat sensation.

In addition to demonstrating GPCR agonist induced inhibition of TRPM3, our in vitro studies have revealed that inverse agonists that act at µ-opioid and NPY receptors (naloxone and

BIIE0246) can potentiate TRPM3 mediated responses. These findings are consistent with the concept that constitutive activity of µ-opioid and NPY receptors provides a level of tonic inhibition of TRPM3. A potentiating effect of these ligands was also noted in vivo, where they potentiated the behavioural effects of local intraplantar injection of PS. This result could

181 reflect an inhibition of constitutive GPCR activity, as suggested by the in vitro findings, or inhibition of endogenous GPCR agonists produced in the tissues. Intraplantar administration of morphine, baclofen, PYY or adenosine inhibited the strong nociceptive behavioural responses evoked by combined local application of PS and CIM0216. Such a peripheral antinociceptive action could be due to an action that inhibits action potential transmission, perhaps by activation of voltage gated potassium channels. However, morphine and adenosine did not inhibit TRPV1-mediated, capsaicin-evoked pain responses so a general inhibitory action of morphine on the transmission of nociceptive signals can be ruled out. The inhibition of TRPM3-mediated nociceptive responses by the GPCR agonists are thus specific to TRPM3-evoked pain, consistent with the observation of direct Gβγ-mediated inhibition of the channel. Our results demonstrate that GPCR modulation of TRPM3 occurs in vivo and that the effects are localized to the regions of sensory nerve terminals rather than systemic effects operating at the level of the spinal cord or higher centres in the nociception pathway.

Our results demonstrate that TRPM3 in sensory neurons is subject to Gi/o GPCR regulation via a Gβγ subunit action. Activation of µ-opioid and GABA-B receptors are important inhibitory mechanisms that operate both peripherally and centrally, including actions on the central terminals of sensory nerves in the spinal dorsal horn. Given the co-expression of TRPM3 and these GPCRs in nociceptive sensory neurons and the emerging role of TRPM3 in nociception, our findings highlight the importance of determining the role of TRPM3 in pathophysiological pain conditions.

4.4.2 Gs- and Gq-mediated inhibition of TRPM3

Our results demonstrate for the first time that TRPM3 is inhibited by activation of GPCRs coupled to any of the three major classes of Gα in heterologous and native cellular

182 environments. Based on our findings with GDPβS-treated cells, we show that TRPM3 is tonically inhibited by G-proteins in HEK293 cells, in good agreement with the tonic inhibition of TRPM3 previously observed in DRG neurons and in vivo (Quallo et al., 2017). Here we show that stimulation of Gs-coupled GPCRs exerts an inhibitory effect on TRPM3 in HEK293 cells and DRG neurons. This inhibition is PTX-insensitive, demonstrating that Gi/o is not responsible.

Furthermore, the inhibition is not mediated by the canonical signalling pathways engaged by

Gs activation, since it is unaffected by PKA inhibition or by stimulation of cAMP production. In contrast, Gs-coupled GPCR inhibition of TRPM3 is fully prevented by expression of βARK1-ct in HEK293 cells, demonstrating that Gβγ protein is responsible for this inhibition.

Previous investigations demonstrated that Gαq does not co-immunoprecipitate with TRPM3

(Dembla et al., 2017) and that inhibition of TRPM3 by heterologous expression of Gq-coupled muscarinic M1 and BK2 receptors is unaffected by PtdIns(4,5)P2 supplementation but is reduced by βARK1-ct (Badheka et al., 2017). Here, we confirmed these findings by showing

2+ that TRPM3-mediated [Ca ]i-responses in HEK293 cells is inhibited by muscarinic M1 receptor activation and that this inhibition is prevented by expression of βARK1-ct and we have further shown that BK2 receptor activation inhibits TRPM3 in vivo and in isolated DRG neurons. This inhibition is mediated by Gβγ and is independent of PKC. Along with our observations that βARK1-ct and GRK2i reversed TRPM3 inhibition by A2B and EP2 activation, these results clearly demonstrate that Gβγ mediates Gi/o-, Gs- and Gq-induced inhibition of

TRPM3.

Gβγ proteins directly modulate N and P/Q type VGCCs and GIRK channels independently of

Gα and irrespective of whether they are liberated from Gαi/o, Gαs or Gαq (See Dascal, 1997;

Dolphin, 2003; Yamada et al., 1998 for review). Our results with GTPγS in control and PTX-

183 treated TRPM3 HEK293 cells show that although PS-evoked currents were gradually inhibited after PTX treatment, the inhibition developed more quickly in the absence of PTX. A more effective inhibition of TRPM3 by Gi/o-coupled GPCRs is supported by the stronger inhibition observed following activation of µ-opioid, NPY and GABAB receptors (Quallo et al., 2017) than with Gs- or Gq-coupled receptors. This finding agrees with the dominant modulation of VGCCs and GIRK channels by activating Gi/o-coupled GPCRs (Dolphin, 2003). Promiscuous GPCR activation of GIRK channels by Gβγ has been observed, but primarily in heterologous overexpression systems. In sinoatrial node pacemaker cells, GIRK is preferentially activated by the Gi/o-coupled M2 muscarinic receptor rather than by the Gs-coupled β2-adrenergic receptor (Digby et al., 2008; Hein et al., 2006; Touhara and MacKinnon, 2018). Reports that

Gβγ proteins dissociate more readily from Gαo than from Gαs (Digby et al., 2008) lend further support to the notion that Gi/o-coupled Gβγ is more effective at inhibiting TRPM3.

The proinflammatory mediators PGE2 and bradykinin, are both thought to produce pain and hypersensitivity at least in part by sensitization of sensory neuron TRP channels such as

TRPA1 and TRPV1, downstream of EP2 and BK2 receptors (reviewed by (Bautista et al.,

2013) and (Veldhuis et al., 2015)). Here, low doses of PGE2 and bradykinin that did not evoke a behavioural response on their own, significantly reduced the pain-related behavioural responses evoked by topical injections of a combination of the TRPM3 agonists

PS and CIM0216, in a manner similar to that observed with application of agonists of the

Gi/o-coupled µ opioid receptor (Badheka et al., 2017; Dembla et al., 2017; Quallo et al.,

2017). This effect appears to be specific for TRPM3, since pain-related behaviours produced by the TRPV1 agonist capsaicin were not inhibited by application of PGE2, butaprost and bradykinin.

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Dose-response curves with intraplantar bradykinin injections in mice demonstrate a significant increase in overt nociception following the administration of 3 nmole of bradykinin and above when compared to PBS and vehicle injections (Ferreira et al., 2004).

This effect was attributed to the activation of BK2 receptors as it was mimicked by the administration of the BK2-selective agonist Tyr8-BK and reversed by the selective antagonist

HOE 140. Administration of BK1 selective agonists and antagonists had no effect on bradykinin-induced overt nociception. This study also demonstrated that administration of

0.3 and 1 nmole of bradykinin did not produce significantly different overt nociception when compared to control and PBS injections. Another study examining the role of bradykinin sensitisation of TRPV4 channels and induction of mechanical hyperalgesia also used various concentrations of bradykinin (Costa et al., 2018). In this study, 0.3 nmole of bradykinin was only capable of sensitising mechanical hyperalgesia 120 minutes post administration. Based on these two studies, we therefore relied on utilising 0.3 nmole of bradykinin, which in our hands also did not evoke a behavioural response on its own. We believe that weak activation of these receptors is enough to inhibit TRPM3 activity and that stronger activation with the use of higher doses is required to produce a nociceptive effect through sensitisation of other TRP channels like TRPV1. Altogether, activation of EP2 and BK2 receptors is unlikely to inhibit pain produced by TRPM3 agonists by interfering with action potential generation or membrane excitability, since many TRPM3 expressing neurons also express TRPV1 (Vriens et al., 2011).

We evaluated the effects of PGE2 and bradykinin on heat hypersensitivity produced by intraplantar FCA. The roles of PGE2 and bradykinin in adjuvant-induced hypersensitivity are not clear and the behavioural effects produced by FCA are, for example, unchanged in mice lacking both BK1 and BK2 bradykinin receptors (Cayla et al., 2012). Intra-articular injections

185 of FCA in the rat produced mechanical hyperalgesia, which was unaffected by inhibition of

BK2 receptors three days after FCA-induction, whereas co-administration of a BK2 antagonist together with FCA prevented the development of hypersensitivity, suggesting a role for BK2 in the development, rather than maintenance of hypersensitivity (Perkins et al., 1993). An evaluation of the efficacy of analgesic drugs in rats treated with intraplantar FCA demonstrated that the non-selective NSAIDs indomethacin and diclofenac, at doses that would completely prevent cyclo-oxygenase mediated PGE2 formation, produced no, or only a minor reduction of heat hypersensitivity (Nagakura et al., 2003). Here, we found that FCA failed to produce heat hypersensitivity in Trpm3-/- mice, and that a selective TRPM3 antagonist, ononetin, produced a complete reversal of the behavioural sensitization produced by FCA in wildtype mice. Surprisingly, local intraplantar injections of either of the proinflammatory mediators PGE2 or bradykinin also produced a complete reversal of the established FCA-induced heat hypersensitivity in wild-type mice. Taken together, our behavioural analysis thus confirms that TRPM3 is critically important for inflammatory heat hyperalgesia (see (Vriens et al., 2011)), and strongly indicates that activation of GPCRs may produce analgesia by inhibiting TRPM3. Somewhat surprisingly, this was the case following local administration of PGE2 and bradykinin. These latter findings are consistent with the absence of high concentrations of PGE2 and bradykinin in the inflamed paw at this post-FCA time point.

Our results demonstrate that TRPM3 is promiscuously inhibited by Gβγ after activation of receptors coupled to any of the major classes of G-proteins and thus may act as a pan-GPCR effector molecule. We show that Gs- and Gq-coupled GPCRs inhibit TRPM3 in cell lines and in isolated sensory neurons in vitro and that they can produce antinociception and analgesia by inhibiting TRPM3 in vivo.

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4.5 Conclusion

TRPM3 is promiscuously inhibited by GPCRs that couple to Gαi/o, Gαq and Gαs through direct

Gβγ protein binding (summarised in Figure 4-23). This inhibition occurs in heterologous expression systems, in native cells and in vivo. Furthermore, TRPM3 may be a tractable target for inflammatory pain, since genetic inactivation, pharmacological inhibition as well as GPCR mediated inhibition of TRPM3 completely prevented or reversed heat hypersensitivity.

Figure 4-23 Schematic representation of promiscuous GPCR-mediated inhibition of TRPM3 ion channels

Active TRPM3 ion channels conduct cations through the channel pore when GPCRs remain inactive (left of the dotted line). Activated GPCRs, whether constitutively or by agonists, leads to the dissociation of Gβγ proteins from Gαi/o, Gαq and Gαs subunits (right of the dotted line). Subsequent binding of Gβγ to TRPM3 inhibits the channel and prevents ionic conductance.

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Chapter 5. GPCR regulation of TRPM2, TRPM7 and TRPM8

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5.1 Introduction

GPCRs have been shown to directly modulate TRPM1 (Gαo, Gβγ) (Koike et al., 2010; Xu et al.,

2016), TRPM3 (Gβγ) (Alkhatib et al., 2019; Quallo et al., 2017) and TRPM8 (Gαq) (Zhang et al.,

2012) through G-proteins. Several studies observed GPCR-mediated modulation of other

TRPM family channels including TRPM2 and TRPM7 through indirect signalling such as PIP2 hydrolysis and protein kinase-mediated phosphorylation. Since G-proteins directly regulate

TRPM1, 3 and 8 channels, I have explored whether other members of the TRPM family are modulated by GPCRs.

5.1.1 TRPM2 activation and modulation by GPCRs

5.1.1.1 TRPM2 activation

The C-terminus of TRPM2 channels contains an adenosine diphosphate ribose (ADPR) pyrophosphatase domain, which is termed Nudix-like domain or NUDT9 homology domain

(NUDT9-H) (Kühn and Lückhoff, 2004). ADPR binds to the NUDT9-H domain where it gates channel opening in the presence of Ca2+ (Csanády and Törocsik, 2009). TRPM2 channels are termed ‘chanzyme’ because they display enzymatic activity by cleaving ADPR and converting it into adenine monophosphate and ribose-5-phosphate (Perraud et al., 2003, 2001).

For a long period, TRPM2 was thought to be activated by other nucleotides including cyclic

ADPR (cADPR) (Kolisek et al., 2005), nicotinamide-adenine dinucleotide (NAD) (Naziroğlu and

Lückhoff, 2008), nicotinic acid-adenine dinucleotide (NAAD) and NAAD-phosphate (NAADP)

(Beck et al., 2006; Lange et al., 2008; Tóth and Csanády, 2010). However, it is unknown whether these nucleotides play a direct role in TRPM2 gating. This is exemplified in the observation that NAD activates TRPM2 with low affinity and that this activation may be due

189 to ADPR contamination (Kolisek et al., 2005). It is also difficult to interpret TRPM2 activation by these intracellular nucleotides since cells expresses several enzymes that can generate and degrade nucleotides thereby making it difficult to control nucleotide concentrations and to identify the active principle with certainty. TRPM2 activation requires intracellular Ca2+ as a coactivator where four Ca2+ ions bind to intracellular crevices near the pore of the channel

(Csanády and Törocsik, 2009).

Investigators of one study purified cADPR and AMP enzymatically and applied them to inside- out excised membrane patches containing TRPM2 channels, and failed to detect activation of the channel (Tóth et al., 2014). Furthermore, activation of TRPM2 by cADPR was shown to be due to contamination by ADPR (Tóth and Csanády, 2010). A later study using similar methods removed contaminant ADPR from NAD and NAAD and removed ADPR phosphate (ADPRP) from NAADP and applied these compounds to excised inside-out membrane patches. NAD,

NAAD and NAADP were all shown to be ineffective at activating TRPM2 and did not compete with ADPR for a binding site within TRPM2 channels whereas ADPRP was demonstrated to be a direct TRPM2 agonist (Tóth et al., 2015). Furthermore, NAADP did not potentiate TRPM2 channel opening by ADPR as previously suggested (Beck et al., 2006). Additionally, TRPM2 channel gating was found to be unrelated to its intrinsic enzyme activity (Tóth et al., 2014).

More recently, the NUDT9-H domain in TRPM2 has been shown to exert no ADPRase enzymatic activity when solubilised chimeras of it were expressed in E.coli cells. Replacement of the NUDT9H in TRPM2 with the soluble chimera retained ADPR-dependent channel gating thereby confirming the functionality of the chimeric domains (Iordanov et al., 2016).

TRPM2 can also be activated by reactive oxygen and nitrogen species (ROS/RNS) such as H2O2

(Hara et al., 2002; Wehage et al., 2002). This activation is believed to be through the

190 generation of ADPR which can be caused by two pathways. In the first pathway, ROS/RNS damage DNA and activate poly-ADPR- polymerase (PARP) and poly-ADPR glycohydrolase

(PARG). PARP and PARG then convert NAD into polymers of ADPR that are then hydrolysed further into monomeric ADPR, which thereafter bind to and activate TRPM2 (Blenn et al.,

2011; Fonfria et al., 2004). The second pathway is due to mitochondrial NUDT9 ADPRase- mediated breakdown of NAD into ADPR. This was demonstrated by expressing NUDT9 targeted to mitochondria in TRPM2-transfected HEK293 cells. NUDT9 degraded mitochondrially generated ADPR and abrogated TRPM2 activation by H2O2 (Perraud et al.,

2005). However, some studies demonstrated that H2O2, but not ADPR, activated a splice variant of TRPM2 found in neutrophil granulocytes with a deletion in the C-terminus (amino acids 1292-1325) (Wehage et al., 2002) suggesting that H2O2-mediated activation of TRPM2 can be independent of ADPR production.

5.1.1.2 GPCR regulation of TRPM2

There are no studies to date that tested whether GPCR activation regulates TRPM2, however, there are some studies that examined the effects of GPCR signal mediators such as PIP2, PKA and PKC on TRPM2 activity. Membrane-bound PIP2 has been shown to be important for

TRPM2 activity (Tóth and Csanády, 2012). Masking or neutralization of PIP2 headgroups with polylysine abolished TRPM2 currents in TRPM2-transfected HEK293 cells. Application of diC8

PIP2 following polylysine removal restored TRPM2 channel activity. PIP2 depletion was shown to decrease TRPM2’s affinity for Ca2+ from 20 µM to 1 mM (Tóth and Csanády, 2012).

Therefore, TRPM2 channels may be regulated by GPCR-mediated activation of PLC and subsequent PIP2 hydrolysis.

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TRPM2 currents in TRPM2-transfected HEK293 cells were potentiated by application of forskolin (Togashi et al., 2006). This potentiation was inhibited by the PKA inhibitor H-89. In pancreatic β-cells, insulin release following forskolin application was reduced in siTRPM2- treated cells suggesting that endogenous TRPM2 may contribute to cAMP-mediated insulin release (Togashi et al., 2006). Furthermore, activation of the Gαs-coupled glucagon-like peptide 1 (GLP-1) receptor increased insulin release and this increase was inhibited by both

H-89 and siTRPM2 treatment (Togashi et al., 2006). Therefore, it was suggested that PKA phosphorylates TRPM2 thereby increasing its activity (Togashi et al., 2006).

Finally, TRPM2 has been suggested to be potentiated by PKC activation in DRG neurons

(Nazıroğlu, 2017). The first study that demonstrated this used patch-clamp electrophysiology to record currents elicited by very high concentration of H2O2 (10 mM) (Nazıroğlu, 2017). The study attributed these currents to be conducted by TRPM2 and utilised the non-specific TRP channel blockers 2-APB and ACA to further support their claim but without providing direct evidence (Nazıroğlu et al., 2011).

5.1.2 TRPM7 activation and modulation by GPCRs

5.1.2.1 TRPM7 activation

Native and heterologously expressed TRPM7 channels are constitutively active and are regulated by various endogenous and molecular modulators (Runnels et al., 2002). Two of the most significant endogenous regulators are Mg2+ and Mg-complexed nucleotides, which tonically inhibit TRPM7 and are the main reason for the low activity observed in resting cells

(Nadler et al., 2001). Dialysing TRPM7-transfected HEK293 cells with intracellular solutions containing physiological levels of Mg2+ (700-900 µM) but lacking MgATP produced large

TRPM7-mediated currents in whole-cell patch-clamp electrophysiology experiments (Nadler

192 et al., 2001). Increasing the MgATP concentration while keeping the Mg2+ concentration constant demonstrated a concentration-dependent inhibition of TRPM7-mediated currents, with a complete suppression at physiological concentrations of MgATP (3-4 mM) and where higher concentrations inhibited TRPM7 constitutive current (Nadler et al., 2001). This MgATP- mediated inhibition of TRPM7 was also observed in Jurkat T cells, RBL-2H3 and human retinoblastoma cells (Hanano et al., 2004). MgATP-mediated inhibition was also shown to be independent of ATP hydrolysis since the nonhydrolyzable MgATPγS also inhibited TRPM7 currents (Nadler et al., 2001). Additionally, other nucleotides such as GTP were able to inhibit

TRPM7 currents (Demeuse et al., 2006; Nadler et al., 2001).

Free nucleotides do not inhibit TRPM7 since perfusing cells with NaATP did not inhibit TRPM7- mediated currents but did actually enhance them (Demeuse et al., 2006; Nadler et al., 2001).

This effect was also observed in another study that interpreted the results as demonstrating a kinase-mediated activation mechanism (Runnels et al., 2001). However, Mg2+ inhibits

TRPM7 on its own (Nadler et al., 2001) and since ATP is the principal intracellular Mg2+ buffer,

2+ Na2ATP potentiates TRPM7 by chelating free Mg , thereby relieving channel block. Removal of both Mg2+ and Mg-complex nucleotides and chelating residual Mg2+ with the buffer N- hydroxyethylenediaminetriacetic acid (HEDTA) or Na2ATP maximally activated TRPM7

(Demeuse et al., 2006).

The first and only small molecule activator of TRPM7 currently known is the DOR anatagonist naltriben (Hofmann et al., 2014). Naltriben reversibly activates native and heterologously expressed TRPM7 channel with an EC50 value of 20 µM (Hofmann et al., 2014). Naltriben was also able to compete with Mg2+-mediated inhibition of TRPM7 and the TRPM7 antagonist

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NS8593 (Hofmann et al., 2014). Mutational studies demonstrated that naltriben likely binds to the TRP domain (Hofmann et al., 2014).

5.1.2.2 GPCR regulation of TRPM7

TRPM7 regulation by GPCR-mediated signalling has been controversial with various studies demonstrating opposing conclusions. One of the modulatory effects of GPCR activation on

TRPM7 activity is proposed to be hydrolysis of membrane PIP2 levels (Runnels et al., 2002).

Carbachol application to M1R-transfected HEK293 cells that endogenously express TRPM7 reversibly inhibited TRPM7-mediated currents (Runnels et al., 2002). This inhibition was found to correlate with PLC activation by measuring PIP2 hydrolysis and IP3 generation

(Runnels et al., 2002). This study also demonstrated that the C2 domain of PLC-β1-3 and PLC-

γ1 interacts with a 146-long amino acid C-terminal segment of the TRPM7 kinase termed the phospholipase C interacting kinase (PLIK). Furthermore, inhibition of TRPM7 following M1R stimulation did not involve any of the other Gαq-mediated pathways such as generation of IP3 and DAG, Ca2+ release from intracellular stores and PKC activity (Runnels et al., 2002). TRPM7 current recovery was also quicker when cells were perfused with PIP2 and was significantly slower when PIP2 generation was inhibited (Runnels et al., 2002). Activation of epidermal growth factor receptor which hydrolyses PIP2 through tyrosine kinase activation of PLC-γ also inhibited TRPM7 currents but to a lesser extent to that observed with M1R activation (Runnels et al., 2002).

Conversely, TRPM7 has also been shown to be activated following application of PLC activators (Clark et al., 2006; Langeslag et al., 2007). Application of bradykinin to TRPM7- transfected N1E-115 cells that endogenously express BK2 receptors caused a sustained Ca2+ influx that was not observed in untransfected cells. Activation of other Gq-coupled GPCRs such

194 as PARs by thrombin also caused the same effect. Additionally, altering cAMP levels by use of forskolin or agonists for Gs- and Gi-coupled GPCRs had no effect on TRPM7-mediated responses. These results indicate that TRPM7 activation is influenced specifically by PLC activation, but not by cAMP and Ca2+ signalling. However, the same study found that bradykinin inhibited TRPM7-mediated currents in TRPM7-transfected N1E-115 cells in the whole-cell patch-clamp configuration. Conversely, bradykinin application activated TRPM7 in the perforated patch-clamp configuration. The exact mechanism behind this difference is unknown, however, the study suggests that PLC regulation of TRPM7 channels in intact cells such as in Ca2+ measurement and perforated patch-clamp experiments are more likely to generate more physiologically relevant results, compared to whole-cell voltage-clamp where the intracellular solution is dialysed and essentially replaced by the pipette solution. Indeed, other studies have also reported PLC-mediated activation of TRPM7 (Kim et al., 2005).

Another controversial modulatory effect of GPCR activation on TRPM7 activity is through cAMP-mediated activation of PKA. The first study examining this effect stated that increased

PKA activity potentiated TRPM7 channels (Takezawa et al., 2004). In this study, carbachol application inhibited TRPM7-mediated currents in TRPM7-transfected HEK293 cells.

2+ Carbachol application increased [Ca ]I through PLC-mediated signalling in untransfected

HEK293 cells which indicates that TRPM7 was inhibited by Gq-mediated signalling (Takezawa et al., 2004). However, these Ca2+ responses were not in all cells tested and were much smaller than those elicited by application of thrombin, which activates endogenous PAR1 receptors

(Takezawa et al., 2004). When thrombin was applied to TRPM7-transfected cells, there was only a small effect on TRPM7-mediated currents thereby suggesting that carbachol’s inhibitory effect was not due to Gq-mediated signalling (Takezawa et al., 2004). Indeed, treatment of TRPM7-transfected cells with PTX reversed carbachol-mediated inhibition of

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TRPM7 suggesting that endogenous muscarinic receptors M2 and M4 that couple to Gαi/o were responsible for TRPM7 inhibition (Takezawa et al., 2004). The study did also observe

2+ that overexpression of TRPM7 prevented increases in [Ca ]i following carbachol and thrombin application. This effect was suggested to be due to TRPM7 affecting PLCβ-mediated

IP3 production by binding to and inhibiting PLC since experiments with pipette-perfused IP3 produced normal Ca2+ release in TRPM7 overexpressing cells (Takezawa et al., 2004). Finally, activation of endogenous Gs-coupled β-adrenergic receptor was shown to potentiate TRPM7- mediated currents (Takezawa et al., 2004). This potentiation was prevented by inhibiting PKA with H89 and KT5720. Additionally, superfusion of TRPM7-transfected HEK293 cells with cAMP also potentiated TRPM7 currents where this effect was reversed with H89 application

(Takezawa et al., 2004). Finally, cAMP and H89 were shown to have no effect on TRPM7 kinase activity in vitro (Takezawa et al., 2004). Altogether, these results suggest that TRPM7 is only regulated by Gi/o- and Gs-coupled GPCRs through their modulator effect on PKA activity.

Several other studies contradicted these findings. Langeslag et al., applied forskolin to

TRPM7-transfected N1E-115 cells and did not witness any increased currents in perforated

2+ patch-clamp experiments and did not observe Ca influx-responses (Langeslag et al., 2007).

Broertjes et al., demonstrated that cAMP-mediated signalling actually has an inhibitory effect on TRPM7 (Broertjes et al., 2019). Application of forskolin inhibited TRPM7-mediated Ca2+ influx in TRPM7-transfected N1E-115 cells (Broertjes et al., 2019). This effect was due to PKA activity as application of H89 reversed forskolin-induced inhibition of TRPM7-mediated Ca2+ influx (Broertjes et al., 2019). PKA mainly phosphorylates serine residues of target proteins, therefore, several TRPM7 with single-point mutations at serine residue sites were generated.

One of these, TRPM7-S1269A, was not inhibited following application of forskolin to TRPM7-

S1269A expressing N1E-115 cells suggesting that PKA inhibits TRPM7 activity by

196 phosphorylating serine residue 1269 (Broertjes et al., 2019). Another study published at the same time also came to the same conclusions (Tian et al., 2018).

The variability between these studies could be attributed to the use of different experimental techniques to measure TRPM7 activity. Takezawa et al., measured TRPM7 activity solely through the use of whole-cell patch clamp electrophysiology (Takezawa et al., 2004) whereas

2+ Langeslag et al., and Broertjes et al., used less invasive [Ca ]i-measurements (Broertjes et al.,

2019; Langeslag et al., 2007). However, the results obtained by Tian and colleagues that agreed with those of Broertjes and colleagues’ results were obtained using whole-cell patch clamp (Tian et al., 2018). Furthermore, none of the studies examining the effects of PLC and

PKC on TRPM7 considered any direct G-protein involvement.

5.1.3 TRPM8 activation and modulation by GPCRs

5.1.3.1 TRPM8 activation

TRPM8 channels are better characterised than TRPM2 and TRPM7 channels where a variety of chemical agonists exist for TRPM8. TRPM8 is gated by cold temperatures, chemical stimuli such as menthol and icilin and by voltage (Brauchi et al., 2004; McKemy et al., 2002; Peier et al., 2002a; Voets et al., 2004a). Activation of TRPM8 by cold and menthol may involve shifting voltage-dependent activation towards more negative potentials thereby increasing the probability of TRPM8 channel openings and increasing inward currents at physiological membrane potentials (Brauchi et al., 2004; Mälkiä et al., 2007; Voets et al., 2004a).

5.1.3.2 GPCR regulation of TRPM8

TRPM8 was found to be inhibited during inflammation through the actions of released inflammatory mediators such as histamine, PGE2 and bradykinin (Linte et al., 2007;

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Premkumar et al., 2005; Zhang et al., 2012). Premkumar and colleagues demonstrated that activation of bradykinin functionally downregulates TRPM8 in DRG neurons. This effect was attributed to PKC-mediated dephosphorylation of TRPM8 and was reversed by phosphatase inhibitors (Premkumar et al., 2005). These findings were further corroborated by Linte and colleagues who demonstrated that bradykinin, acting through PKC, and PGE2, acting through

PKA, both reduced responses to cooling and shifted temperature thresholds to cold levels in cold- and menthol-sensitive DRG neurons (Linte et al., 2007). TRPM8 inhibition by Gq-coupled

GPCRs was also attributed to PLC-mediated hydrolysis of membrane PIP2 (Liu and Qin, 2005;

Rohács et al., 2005). Both studies demonstrated that PIP2 hydrolysis inhibited TRPM8- mediated responses. Direct application of PIP2 onto excised inside-out membrane patches restored TRPM8 current rundown in TRPM8-transfected HEK293 cells and activation of the tyrosine receptor kinase platelet-derived growth factor β receptor inhibited TRPM8 activity through PLC-γ-mediated hydrolysis of PIP2 (Liu and Qin, 2005; Rohács et al., 2005). However, histamine H1 and BK2 receptor activation were later found to inhibit TRPM8 directly through

Gαq subunit binding (Zhang et al., 2012). This inhibition was abolished following deletion of

Gαq, and rescued by expression of a Gαq chimaera that lacked the ability to activate downstream signalling pathways (Zhang et al., 2012). Direct application of activated purified

Gαq proteins but not Gβγ proteins onto excised inside-out membrane patches of TRPM8- transfected HEK293 cells inhibited TRPM8-mediated currents (Zhang et al., 2012). This Gαq- mediated inhibition was only demonstrated in heterologous expression systems and not in native sensory neurons and did not distinguish between inhibition produced directly by Gαq or its downstream signalling events.

More recently, direct Gαq binding to TRPM8 following BK2 activation was found to be the sole mechanism for TRPM8 inhibition in sensory neurons (Zhang, 2019a). This effect did not

198 involve PLCβ-PIP2 signalling or Gα11 subunits (Zhang, 2019a). The number of TRPM8-mediated action potentials (spike events) in DRG neurons from Gαq KO mice in cell-attached patch- clamp experiments were significantly higher than those in wildtype mice thereby suggesting that Gαq tonically inhibits TRPM8 activity. Furthermore, bradykinin application inhibited

TRPM8-mediated spike events in wildtype DRG neurons but had no effect on neurons from

Gαq KO animals (Zhang, 2019a). This effect was not due to impaired PLCβ signalling as Gα11 was able to sensitise TRPV1 activity through PKC activation which was prevented by application of U73122 (Zhang, 2019a). These results also demonstrate that Gαq and not Gα11 inhibits TRPM8 channels, agreeing with previous studies (Li and Zhang, 2013). Zhang demonstrated that three basic arginine residues on TRPM8 (R364, R 368 and R470) were found to function as Gαq effector sites (Zhang, 2019a). When these residues were mutated to glutamine, causing a TRPM8 triple mutant (TRPM8-TM), TRPM8-TM exhibited much larger currents than wildtype TRPM8 in transfected-HEK293 cells and was not inhibited by Gαq

(Zhang, 2019a). Bradykinin application was shown to robustly inhibit TRPM8-currents in

TRPM8/BK2-transfected HEK293 cells (Zhang, 2019a). This inhibition was abolished with the expression of TRPM8-TM suggesting that BK2-mediated inhibition of TRPM8 is solely reliant on Gαq protein binding and not PIP2 hydrolysis (Zhang, 2019a). Furthermore, activation of H1 receptors was still able to inhibit 43% of inward TRPM8-TM-mediated current where this inhibition was completely abolished with U73122 application (Zhang, 2019a). Therefore, BK2- mediated inhibition of TRPM8 is solely due to Gαq protein binding whereas H1-mediated inhibition is reliant on both Gαq and PIP2 hydrolysis. Interestingly, BK2 coexpression with

TRPM8 or TRPM8-TM protected against TRPM8 inhibition following PIP2 hydrolysis whereas coexpression with H1 receptors did not (Zhang, 2019a). The reason behind this may be because BK2 receptors bind to TRPM8 whereas H1 receptors do not (Zhang, 2019a).

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Altogether, TRPM8 channels may be inhibited following Gq-coupled GPCR activation solely through Gαq subunits or together with PIP2 hydrolysis.

TRPM8 regulation by Gi/o- and Gs-coupled GPCRs has not been as extensively studied. Two studies demonstrate Gi/o-mediated inhibition of TRPM8 channels through PKA activity

(Bavencoffe et al., 2010; De Petrocellis et al., 2007). Stimulation of Gi/o-coupled α2A- adrenoreceptors in rat DRG neurons inhibited TRPM8 activity (Bavencoffe et al., 2010). In

TRPM8 transfected-HEK293 cells, application of forskolin had no effect on menthol-evoked

TRPM8 currents whereas treatment with the adenylyl cyclase inhibitor SQ22536 caused

TRPM8 currents to decrease by ~60% (Bavencoffe et al., 2010). Forskolin application did prevent inhibition by activation of α2A-adrenoreceptors and this effect was also witnessed when endogenous Gs-coupled β-adrenergic receptors were activated in HEK293 cells

(Bavencoffe et al., 2010). Furthermore, application of KT5720 and H89 reduced cold- and menthol-activated TRPM8 currents (Bavencoffe et al., 2010). These results suggest that α2A- adrenoreceptor activation inhibits TRPM8 by decreasing PKA activity. S9D and T17D single point mutations in TRPM8 protected against inhibition by α2A-adrenoreceptor activation indicating that these residues serve as phosphorylation sites for PKA (Bavencoffe et al., 2010).

Conversely, De Petrocellis et al., demonstrated that increased activity of PKA inhibits TRPM8

2+ (De Petrocellis et al., 2007). Application of forskolin or 8-bromo-cAMP in [Ca ]i measuring experiments produced a rightward shift in icilin and menthol dose-response curves in TRPM8- transfected HEK293 cells where this effect was prevented by cotreatment with the selective

PKA inhibitor Rp-cAMP-S (De Petrocellis et al., 2007).

The discrepancy between both studies may be due to the same reason behind the discrepancies seen with TRPM7. Bavencoffe et al., relied solely on whole-cell patch-clamp

200 electrophysiology while Petrocellis et al., only utilised Ca2+-measuring experiments

(Bavencoffe et al., 2010; De Petrocellis et al., 2007). Furthermore, no study has examined whether TRPM8 can interact directly with Gαi/o or Gαs subunits.

5.2 Aims of the present study

Since we have established the activity of endogenous HEK293 cell GPCRs that signal through all three Gα proteins in intact cells using Ca2+ imaging experiments, we set out to examine

Gi/o-, Gq/11- and Gs-coupled GPCR regulation of TRPM2, TRPM7 and TRPM8 channels.

These experimental aims were investigated using single cell and cell population Ca2+- measurements of TRPM2, TRPM7 or TRPM8 transfected HEK293 cells.

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5.3 Results

5.3.1 GPCR modulation of TRPM2

No synthetic compound has been found to activate TRPM2 channels in a reversible manner.

To investigate TRPM2 modulation by GPCRs in Ca2+-imaging experiments, we utilised two activators of TRPM2: H2O2, which activates the channel in an irreversible manner and heat.

5.3.1.1 GPCR modulation of H2O2-mediated activation of TRPM2

Application of high concentrations of H2O2 can oxidise cysteine residues in proteins other than

TRPM2 and I therefore compared H2O2 concentrations response curves in untransfected and

2+ TRPM2-transfected HEK293 cells. Application of H2O2 robustly increased [Ca ]i-responses in

TRPM2-transfected cells with an EC50 of 30.1 ± 1.7 µM at 25 °C (Figure 5-1 A). In contrast, H2O2

2+ only evoked [Ca ]i-responses at high concentrations of H2O2 (5 mM, Figure 5-1 A).

We next performed Ca2+-imaging experiments in TRPM2-transfected cells treated with a maximally effective H2O2 concentration (500 µM). Application of H2O2 (500 µM) produced

2+ strong [Ca ]i-responses that plateaued after roughly 10 minutes of H2O2 application and did not desensitise (relative amplitude of 101 ± 0.6 %; Figure 5-1 B and E). The shape of H2O2 activation of TRPM2 demonstrates positive feedback by Ca2+. PAR2 activation leads to signalling through both Gαi/o and Gαq/11, we therefore utilised this promiscuity to test the effect of Gi/o- and Gq/11-mediated signalling on TRPM2 activity. Application of a low concentration of 2F-LIGRLO (100 nM) that can activate both Gi/o and Gq/11 proteins had no effect on H2O2-mediated activation of TRPM2 when compared to control (103 ± 0.49 %; Figure

5-1 B and E). We next assessed the effect of activating endogenous Gs-coupled A2B receptors on TRPM2 activity. Application of a maximally activating concentration of adenosine (100 µM)

202 had no effect on H2O2-mediated activation of TRPM2 when compared to control (107.5 ± 0.6

%; Figure 5-1 D and E). These results suggest that TRPM2 is not regulated by Gαi/o, Gαq/11, Gαs and Gβγ subunits.

Figure 5-1 GPCR activation does not regulate TRPM2 channel activation by H2O2

2+ A Concentration response curve of [Ca ]i responses in TRPM2 transfected or untransfected HEK293 cells

2+ following application of H2O2 ± SEM. B [Ca ]i responses in TRPM2 HEK293 cells with H2O2 application (500 µM).

2+ C [Ca ]i responses in TRPM2 HEK293 cells with H2O2 application (500 µM) followed by application of 2F-LIGRLO

2+ (100 nM). D [Ca ]i responses in TRPM2 HEK293 cells with H2O2 application (500 µM) followed by application of adenosine (100 µM). E Scatter plot representing the relative amplitudes of control (n = 573), 2F-LIGRLO (n =

854)- and adenosine (n = 462)-treated TRPM2 HEK293 cells.

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5.3.1.2 GPCR modulation of heat-mediated activation of TRPM2

Since activation of TRPM2 by H2O2 appears irreversible and is produced by oxidation of methionine rather than cysteine residues (Kashio et al., 2012), it is possible that this covalent mode of activation prevented protein-protein interactions with G-proteins. In addition to being activated by H2O2 and ADPR, TRPM2 is a heat sensor that can be activated by warm or hot temperatures. We therefore used two subsequent heat ramps to activate the channel and tested the effect of GPCR agonists on the second heat response. In control experiments,

2+ heat ramps elicited strong [Ca ]i-responses in TRPM2-transfected HEK293 cells at 50 °C that appeared to be fully desensitised in the second heat application (relative amplitude of 40.7 ±

0.5 %, Figure 5-2 A). When done in untransfected HEK293 cells, the first heat ramp elicited

2+ significantly smaller [Ca ]i-responses than in TRPM2-transfected HEK293 cells and the response did not desensitise during the second heat ramp (98.4 ± 1.1 %; Figure 5-2 B). These

2+ results demonstrate that the heat ramp-induced [Ca ]i-responses in TRPM2-HEK293 cells are mainly mediated by TRPM2 channel activation but that they fully desensitise. Application of adenosine (100 µM) prior to and during the second heat response had no effect on the second heat ramp when compared to control (40.3 ± 0.4 %; Figure 5-2 C). To test whether PKA activation affects TRPM2 activity, we applied forskolin prior to and during the second heat

2+ ramp and observed no difference in [Ca ]i-responses when compared to control experiments

(38.5 ± 0.5 %; Figure 5-2 E). In contrast, application of 2F-LIGRLO (100 nM) elicited significantly

2+ larger [Ca ]i-responses during the second heat ramp when compared to control that were not affected by co-treatment of the PKC inhibitor BIM VIII (control: 40.6 ± 0.5 %; 2F-LIGRLO:

54.4 ± 0.6 % and 2F-LIGRLO + BIM VIII: 56.2 ± 0.6 %; Figure 5-2 F and G). Similarly, ATP

2+ application significantly increased [Ca ]i-responses during the second heat ramp when compared to control (control: 40.3 ± 0.4 %; ATP: 93.8 ± 3 %; Figure 5-2 H and I. However, this

204 effect was also observed in untransfected HEK293 cells where 2F-LIGRLO (100 nM) application

2+ elicited significantly larger [Ca ]i-responses during the second heat ramp when compared to control (control: 98.4 ± 1.1 %; 2F-LIGRLO: 156 ± 1.2%; Figure 5-2 J and K). This effect may possibly be due to increased kinetic rates for Gq/11-mediated signalling caused by increases in

2+ energy (heat). This in turn may increase the rate of activation of PLCβ leading to larger [Ca ]i- responses through downstream signalling via IP3 receptor activation and subsequent release of intracellular Ca2+ stores. Altogether, these results suggest that TRPM2 activity by heat cannot be rescued by GPCR activation but are inconclusive in determining whether GPCR activation inhibits TRPM2 activity.

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Figure 5-2 GPCR activation does not regulate TRPM2 channel activation by heat

2+ 2+ A [Ca ]i responses in TRPM2 HEK293 cells stimulated with two subsequent heat ramps (50 °C). B [Ca ]i

2+ responses in untransfected HEK293 cells stimulated with two heat ramps (50 °C). C [Ca ]i responses in TRPM2

HEK293 cells stimulated with two heat ramps (50 °C) and treated with adenosine (100 µM). D Scatter plot representing the relative amplitude in A (n = 573) and C (n = 462). E Scatter plot representing the relative amplitude in control (n = 564), adenosine- (n = 467) and forskloin-treated (10 µM, n = 467) TRPM2 HEK293 cells

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2+ (n = 462) F [Ca ]i responses in TRPM2 HEK293 cells stimulated with two heat ramps (50 °C) and treated with 2F-

LIGRLO (100 nM). G Scatter plot representing the relative amplitude in control (n = 494), 2F-LIGRLO- (n = 561) and 2F-LIGRLO + BIM VIII-treated (1 µM, n = 551) TRPM2 HEK293 cells. I Scatter plot representing the relative

2+ amplitude in control (n = 561) and ATP-treated (n = 585) TRPM2 HEK293 cells. J [Ca ]i responses in untransfected

HEK293 cells stimulated with two heat ramps (50 °C) and treated with 2F-LIGRLO (100 nM). K Scatter plot representing the relative amplitude in control (n = 506) and 2F-LIGRLO-treated (n = 654) untransfected HEK293 cells.

5.3.2 GPCR modulation of TRPM7

5.3.2.1 Gi/o- and Gq-coupled GPCR modulation of TRPM7

We utilised the DOR agonist naltriben which was recently shown to be an activator for TRPM7

2+ to study whether GPCRs regulate TRPM7 activity in [Ca ]i-imaging experiments. When we first applied naltriben (30 µM) to transiently transfected TRPM7 HEK293 cells, we did not

2+ 2+ observe any [Ca ]i-responses. To determine whether this was due to including Mg block of

TRPM7 by Mg2+ ions in the extracellular solution. We therefore applied Mg2+-free solution to

2+ TRPM7 and observed the development of strong [Ca ]i-responses (Figure 5-3 A). Afterwards,

2+ application of naltriben (30 µM) elicited even greater [Ca ]i-responses that were reversible following washout of the drug (Figure 5-3 A). Reapplication of extracellular solutions with

Mg2+ inhibited TRPM7 activity (Figure 5-3 A). Together, these results demonstrate that TRPM7 activity is blocked by extracellular Mg2+ ions and, although constitutively active in Mg2+-free environments, TRPM7 can be further activated by naltriben.

Naltriben application in Mg2+-free solutions was able to evoke robust, non-desensitising and

2+ reversible [Ca ]i-responses in these cells (relative amplitude 97.8 ± 0.7 %; Figure 5-3 B and

2+ D). Next, we applied 2F-LIGRLO (100 nM) and observed no effect on TRPM7-mediated [Ca ]i-

207 responses (96.6 ± 1.3 %; Figure 5-3 C and D). These results demonstrate that neither Gαi/o nor

Gαq/11 regulate the channel.

Figure 5-3 PAR2 activation does not regulate TRPM7 channel activity

2+ 2+ 2+ A [Ca ]i-responses in TRPM7 transfected HEK293 cells in the presence of Mg -containing and Mg -free

2+ solutions along with application of naltriben (30 µM) B [Ca ]i responses in TRPM7-transfected HEK293 cells with

2+ naltriben application (30 µM). C [Ca ]i responses in TRPM7-transfected HEK293 cells with naltriben and 2F-

LIGRLO (100 nM) application. D Scatter plot representing the relative amplitude from A (n = 436) and B (n = 209).

5.3.2.2 Gs-coupled modulation of TRPM7

In contrast to control experiments, application of adenosine (100 µM) significantly increased

2+ naltriben-evoked (30 µM) [Ca ]i-responses in a reversible manner (control: 97.8 ± 0.7 % and adenosine: 121.7 ± 1.2 %; Figure 5-4 A and B). To test whether this effect relied on TRPM7 being activated by naltriben, we used a sole adenosine application (100 µM) and observed a

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2+ reversible increase in [Ca ]i-responses (Figure 5-4 C). To ensure that this response was in only

TRPM7-transfected HEK293 cells, we subsequently applied naltriben (30 µM) and observed

2+ an increase in [Ca ]i (Figure 5-4 C). Adenosine was applied again during the naltriben-

2+ mediated response and it was able to reversibly increase [Ca ]i-responses (Figure 5-4 C). In

2+ contrast, adenosine (100 µM) had no effect in [Ca ]i-responses in cells that were not responsive to naltriben thereby demonstrating that adenosine’s effect is specific to TRPM7- transfected cells. These results indicate that adenosine is able to reversibly and repeatedly increase both constitutive TRPM7 activity and channel activation evoked by the agonist naltriben.

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Figure 5-4 A2B activation potentiates TRPM7 channel activity

2+ A [Ca ]i responses in TRPM7-transfected HEK293 cells with naltriben application (30 µM) and adenosine (100

µM) application. B Scatter plot representing the relative amplitude from control (n = 436) and A (n = 281). C

2+ [Ca ]i responses in TRPM7-transfected HEK293 cells with adenosine (100 µM) and naltriben (30 µM) + adenosine application (n = 72).

We next tested whether adenosine-mediated potentiation of TRPM7 was mediated by PKA.

Application of the selective PKA inhibitor H89 (10 µM) prior to and during adenosine application had no effect on adenosine-mediated potentiation of TRPM7 responses (control:

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109.9 ± 1 %; adenosine: 119.3 ± 0.72 % and adenosine + H89: 118.7 ± 1.6 %; Figure 5-5 A and

B). To test whether TRPM7 channels were being potentiated by Gβγ protein, we co- transfected TRPM7 and βARK1-ct in HEK293 cells. βARK1-ct expression had no effect on adenosine-mediated potentiation of TRPM7 channels (control: 97.8 ± 0.7 %; adenosine: 112.8

± 1 % and adenosine + βARK1-ct: 112.9 ± 0.9 %; Figure 5-5 C and D). These results indicate that TRPM7 potentiation following A2B activation is not due to Gβγ proteins or increased PKA activity.

Figure 5-5 A2B-mediated potentiation of TRPM7 does not rely on PKA and Gβγ protein

2+ A [Ca ]i responses in TRPM7-transfected HEK293 cells with naltriben application (30 µM), adenosine (100 µM) and H89 (10 µM) application. B Scatter plot representing the relative amplitude from control (n = 543).

2+ adenosine (n = 605) and adenosine + H89 (n = 683). C [Ca ]i responses in TRPM7- and βARK1-ct-transfected

HEK293 cells with naltriben (30 µM) and adenosine (100 µM) application. D Scatter plot representing the relative amplitude from control (n = 436). adenosine (n = 522) and adenosine + βARK1-ct (n = 660).

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5.3.3 GPCR modulation of TRPM8

Although TRPM8 has been shown to be directly inhibited by Gαq subunits, interactions with other Gα proteins were not examined or reported. Here, we utilised the TRPM8 agonist

2+ menthol to investigate the effect of GPCR activation on TRPM8-mediated [Ca ]i-responses.

5.3.3.1 TRPM8 is solely regulated by Gαq proteins

Transiently transfected TRPM8 channels in HEK293 cells were activated by two subsequent applications of menthol (50 µM) and did not desensitise (relative amplitude of 101.2 ± 0.7 %;

Figure 5-4 A and C). Application of adenosine (100 µM) prior to and during the second

2+ menthol application had no effect on [Ca ]i-response amplitudes (98.2 ± 0.7 %; Figure 5-4 B and C). In contrast, application of 2F-LIGRLO (100 nM) prior to and during the second menthol

2+ application lead to a significant decrease in menthol-evoked [Ca ]i-responses (82 ± 0.8 %;

Figure 5-4 D and F). As described earlier, PAR2 activation leads to promiscuous signalling through Gαi/o and Gαq/11 subunits. Therefore, to distinguish which G-protein was responsible for PAR2-mediated inhibition of TRPM8, we pre-treated TRPM8-transfected HEK293 cells with

PTX (200 ng/ml, 24 hours). PTX treatment had no effect on 2F-LIGRLO-mediated inhibition of

TRPM8 activity (84.2 ± 1.2 %; Figure 5-4 E and F). This data suggests that PAR2 is inhibiting

TRPM8 channel activity through Gq/11-mediated signalling. To test whether Gβγ protein was responsible for this inhibition, TRPM8-transfected HEK293 cells were also transfected with

βARK1-ct. Expression of βARK1-ct had no effect on PAR2-mediated inhibition of TRPM8 (72.1

± 0.6 %; Figure 5-4 G and H). Altogether, these results demonstrate that Gαi/o, Gαs and Gβγ do not regulate TRPM8 activity and confirm TRPM8 regulation by Gq/11-mediated signalling.

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Figure 5-6 TRPM8 activity is only regulated by Gq/11-mediated signalling

2+ A [Ca ]i responses in TRPM8 transfected HEK293 cells treated with two short applications of the TRPM8 agonist

2+ menthol (50 µM). B [Ca ]i responses in TRPM8 transfected HEK293 cells treated with menthol and adenosine

2+ (100 µM). C Scatter plot representing the relative amplitude from A (n = 1117) and B (n = 868). D [Ca ]i responses

2+ in TRPM8 transfected HEK293 cells treated with menthol and 2F-LIGRLO (100 nM). E [Ca ]i responses in PTX-

(200 ng/ml) pretreated TRPM8 transfected HEK293 cells stimulated with menthol and 2F-LIGRLO. F Scatter plot

2+ representing the relative amplitude from control (n = 1117), D (n = 964) and E (n = 652). G [Ca ]i responses in

TRPM8/βARK1-ct transfected HEK293 cells treated with menthol and 2F-LIGRLO. H Scatter plot representing the relative amplitude from control (n = 1117), 2F-LIGRLO (n = 964) and βARK1-ct-transfected TRPM8 HEK293 cells

(n = 1034).

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5.4 Discussion

5.4.1 GPCR modulation of TRPM2

5.4.1.1 GPCR activation has no effect on TRPM2 activation by oxidative stress

Our results confirm that TRPM2 channels are activated by H2O2 and that TRPM2 activation

2+ 2+ leads to increases in [Ca ]i-responses. H2O2 application also produced [Ca ]i-responses in native HEK293 cells, however, these were only observed at high concentrations of H2O2 (5 mM). These results throw into doubt the findings that PKC potentiated TRPM2 activity in DRG neurons following application of 10 mM H2O2 (Nazıroğlu, 2017). We show that activation of endogenous A2B and PAR2 receptors, which collectively signal through Gαi/o, Gαq/11 and Gαs,

2+ had no effect on TRPM2-mediated [Ca ]i-responses. These findings indicate that neither Gα subunits nor Gβγ proteins have a direct effect on TRPM2 gating.

5.4.1.2 GPCR activation has no effect on heat-induced activation of TRPM2

Since H2O2-mediated activation of TRPM2 is irreversible it may prevent any modulatory interaction with activated G-proteins and we therefore used heat as another activator to test whether TRPM2 activity can be modulated by GPCRs. In this study, we demonstrate that

TRPM2 is activated at high temperature thresholds (50 °C) and that this activation strongly desensitises which is evidenced by the weaker activation following a second heat ramp. In

2+ contrast, heat in native HEK293 cells only elicited small increases in [Ca ]i that were identical during both heat ramps. These results indicate that TRPM2 is strongly activated by heat and but that it strongly desensitises after the first stimulus. Activation of A2B receptors had no effect on the second heat-elicited response. Furthermore, treatment with forskolin did not resensitise TRPM2 responses in the second heat application. These results disagree with work

214 demonstrating that PKA activation by forskolin or downstream of Gs signalling sensitises

TRPM2 channels (Togashi et al., 2006).

The effect of PAR2 and endogenous P2Y receptor activation on TRPM2-mediated responses to heat ramps could not be accurately investigated as heat challenges increased Gq-mediated

Ca2+ responses. This effect was observed in both native and TRPM2-expressing HEK293 cells which further supports an increase in Gq-mediated signalling rather than a TRPM2-related effect.

5.4.2 GPCR modulation of TRPM7

Our results confirm TRPM7 block by extracellular Mg2+ ions as Mg2+-free conditions led to robust TRPM7-mediated responses both in the presence and absence of naltriben.

Interestingly, TRPM7 can be further activated by naltriben in Mg2+-free conditions suggesting that the channel is not maximally activated during constitutive activity. Activation of PAR2

2+ had no effect on TRPM7-mediated [Ca ]i-responses indicating that Gαi/o- and Gαq proteins do not regulate the channel.

Conversely, activation of A2B receptors lead to a clear and reversible potentiation of TRPM7- mediated responses. This potentiation was observed both during constitutive TRPM7 activity and naltriben-induced TRPM7 activation. Furthermore, inhibition of PKA had no effect on adenosine-mediated potentiation of TRPM7 responses.

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5.4.3 GPCR modulation of TRPM8

Our results demonstrate that TRPM8 activation by menthol is not affected by activation of endogenous A2B receptors. Based on [cAMP] measurement assays in native HEK293 cells

(Chapter 4), 100 µM adenosine application lead to maximal [cAMP] increases likely to activate

2+ PKA. Since adenosine application had no effect on TRPM8-mediated [Ca ]i-responses, we conclude that neither Gαs nor PKA activity regulate the channel. This disagrees with the published literature which shows conflicting data on PKA-mediated inhibition or activation of

TRPM8 activity (Bavencoffe et al., 2010; De Petrocellis et al., 2007).

2+ Conversely, PAR2 activation lead to a significant decrease in TRPM8-mediated [Ca ]i- responses. This inhibition was not affected by PTX treatment. Therefore, in contrast to the promiscuous inhibition of TRPM3 by PAR2 through Gi/o- and Gq-signalling, TRPM8 was only inhibited by PAR2 through PTX-insensitive Gq-mediated signalling. Co-transfecting TRPM8 cells with βARK1-ct had no effect on PAR2-mediated inhibition of TRPM8. Together, these results demonstrate that TRPM8 regulation by GPCRs is solely through Gαq proteins and their downstream signalling events.

5.5 Conclusion

The results of this chapter demonstrate that members of the TRPM subfamily are differentially regulated by GPCRs and that not all can be regulated by direct G-protein interactions. They also confirm that TRPM3, hitherto, is unique among the TRPM subfamily in that it can be directly regulated by GPCRs that signal through different Gα subunits.

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Chapter 6. General discussion, study limitations and future directions

The first aim of this thesis was to characterise TRPM3 activity so that we could determine how to accurately examine its role in somatosensation. Many ion channels including TRP channels are regulated by GPCRs and therefore the second aim of this thesis was to explore

GPCR modulation of TRPM3 channels in heterologous and native systems. This modulation was further explored in an inflammatory model of chronic pain to make these studies more physiologically relevant. Since we, along with others, initially discovered that TRPM3 channels are inhibited by GPCRs that couple to Gαi/o, and then demonstrated promiscuous inhibition by Gαq- and Gαs-coupled receptors (Alkhatib et al., 2019; Badheka et al., 2017;

Dembla et al., 2017; Quallo et al., 2017), the final aim of this thesis was to explore whether

GPCRs can modulate other members of the TRPM family such as TRPM2, TRPM7 and

TRPM8.

In the first results chapter (Chapter 3), TRPM3 activation by pregnenolone sulphate and

CIM0216 was characterised using Ca2+ fluorometric assays and patch-clamp electrophysiology. Both agonists produced sustained concentration-dependent, TRPM3-

2+ mediated increases in [Ca ]i. In patch-clamp experiments, PS-evoked TRPM3 currents were outwardly rectifying, did not plateau at high voltages of up to +200 mV and desensitised in a

Ca2+-independent manner. Conversely, CIM0216-evoked TRPM3 currents were biphasic, double-rectifying, plateaued at high voltages and desensitised in a Ca2+-dependent manner.

The desensitisation with both agonists in patch-clamp experiments was not observed in fluorometric assays. TRPM3 current desensitisation was not complete and was approximately 50% of the peak current.

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The mechanism behind TRPM3 current desensitisation was further explored by testing the

2+ effect of intracellular PIP2 supplementation or extracellular Ca exclusion. Supplementing cells with PIP2 had no effect on the desensitisation of PS-evoked currents. This would rule out PLCδ -mediated PIP2 hydrolysis being the reason. However, bovine brain-derived PIP2 was used for these experiments which is not readily solubilised in aqueous solutions (Tóth et al., 2015). diC8-PIP2, which is water soluble, should be used in future experiments to rule out the involvement of PLCδ in TRPM3 current desensitisation. Excluding extracellular Ca2+ had no effect on PS-evoked current tachyphylaxis but almost completely prevented tachyphylaxis of CIM0216-evoked currents. This suggests that CIM0216-mediated activation of TRPM3 is strongly affected by Ca2+ influx and the role of PLCδ and calmodulin should be further explored in the future.

One limitation in this study is that the recovery of TRPM3 tachyphylaxis was not examined.

This could have been done by assessing whether increasing the time between PS applications would allow for TRPM3 currents to recover from being desensitised. This approach may have helped determine whether PS-evoked TRPM3 current tachyphylaxis is due to an intrinsic, self-regulating mechanism in the channel or the involvement of another molecule. However, this approach would not work with CIM0216 as the compound does not fully wash off after application.

The effect of PS and CIM0216 co-application should also be examined more thoroughly in future studies. Application of a low concentration of CIM0216 that does not produce a

TRPM3-mediated current on its own strongly sensitised the channel to PS applications (Held et al., 2015a). We utilised this synergy in in vivo behavioural experiments but did not examine it further in patch clamp experiments. Future studies can investigate whether co-

218 application of these agonists is also susceptible to tachyphylaxis. The effect of osmolarity and heat on TRPM3 activation by PS and CIM0216 and subsequent channel desensitisation should also be examined in further detail.

In the second results chapter (Chapter 4), the modulation of TRPM3 channels by GPCRs was examined using fluorometric assays, single-cell imaging, patch-clamp experiments and in vivo behavioural experiments. We discovered that activation of Gi/o protein-coupled receptors in HEK293 cells and isolated DRG neurons inhibited TRPM3 activity through Gβγ proteins binding directly to TRPM3 channels. This inhibition was observed with the activation of multiple Gi/o-coupled receptors such as MOR, NPY, GABAB and adenosine A1 receptors. However, activation of other types of receptors such as CB1 receptors did not significantly inhibit TRPM3 activity, which possibly indicates that some GPCRs may be better co-expressed with TRPM3 than others. Gi/o-coupled receptor inhibition of TRPM3 was also observed in in vivo behavioural experiments with mice. This inhibition was specific to

TRPM3 activity as administration of morphine or adenosine did not affect the number of pain responses or their duration following capsaicin administration.

Since Gβγ protein is liberated from GPCRs that couple to Gαq and Gαs and other ion channels such as GIRKs and VGCCs exhibit promiscuous inhibition by GPCRs, we investigated whether TRPM3 channels can also be promiscuously inhibited by GPCRs. Activation of endogenous HEK293 Gq-coupled purinergic receptors and promiscuously Gi/o- and Gq- coupled PAR2 receptors, exogenous muscarinic M1 receptors and endogenous DRG BK2 receptors all inhibited TRPM3 through release of Gβγ protein. This effect does not rely on

PIP2 hydrolysis as supplementing cells with bovine-brain derived and diC8 PIP2 had no significant effect on purinergic receptor- and PAR2-mediated inhibition of TRPM3 in HEK293

219 cells. Furthermore, membrane PIP2 hydrolysis using rapamycin with the PM-FRB-CFP and

RFP-FKBP12-5ptpase constructs did not decrease PAR2-mediated inhibition of TRPM3 activity. However, since PAR2 was found to promiscuously inhibit TRPM3 through a Gi/o- and

Gq-coupled mechanism, as evidenced by using PTX, we cannot be fully certain of the level of contribution that PIP2 hydrolysis plays in Gq-protein coupled receptor inhibition of TRPM3.

Future studies should attempt to investigate the role of PIP2 hydrolysis using GPCRs that solely signal through Gαq. Nevertheless, Gq-protein coupled receptor inhibition of TRPM3 was fully prevented when βARK1-ct was co-expressed in HEK293 cells and significantly lowered with the inclusion of GRK2i in patch pipettes in DRG neurons. This indicates that liberated Gβγ proteins are the major signalling pathway involved in Gq-protein coupled inhibition of TRPM3. Similarly, activation of endogenous HEK293 Gs-coupled adenosine A2B and endogenous DRG prostaglandin EP2 receptors both inhibited TRPM3 activity. This inhibition was through liberated Gβγ proteins as expressing βARK1-ct or inclusion of GRK2i in the patch pipette prevented inhibition of TRPM3. Gq-coupled BK2- and EP2-mediated inhibition of TRPM3 was also observed in in vivo experiments. This inhibition was specific to

TRPM3 as activating either receptor had no effect on capsaicin-elicited responses.

TRPM3 KO studies demonstrated that the channel is required for the development of heat hyperalgesia associated with inflammation (Vriens et al., 2011). We therefore administered the TRPM3 antagonist ononetin, bradykinin and PGE2 in mice following FCA treatment to observe whether TRPM3 inhibition would reverse the developed heat hypersensitivity.

Indeed, administration of all three compounds fully reversed heat hyperalgesia associated with inflammation and produced similar results to a TRPM3 KO model. This further confirms that the concentrations of bradykinin and PGE2 can activate BK2 and EP2 receptors sufficiently enough to inhibit TRPM3 in a similar manner to selective TRPM3 antagonists.

220

One limitation to this study is that we did not explore whether activating GPCRs that couple to Gα12/13 are also able to inhibit TRPM3 activity. One receptor that should be investigated is the lysophosphatidic acid (LPA) receptor 5. LPA5 is expressed in DRG neurons of mice and signals through Gα12/13 and Gαq proteins (Lee et al., 2006). KO of LPA5 receptors in mice has been shown to protect against the development of neuropathy following partial sciatic nerve ligation (Lin et al., 2012). Future experiments should determine whether activating

LPA5 receptors affects TRPM3 activity through a Gα12/13-mediated mechanism and if so whether it is by liberated Gβγ proteins. If TRPM3 is also inhibited by Gβγ that is liberated from activated Gα12/13 subunits, then TRPM3 would potentially be an effector molecule for all GPCRs regardless of what Gα protein they signal through.

In the third results chapter (Chapter 5), the modulation of TRPM2, TRPM7 and TRPM8 channels by GPCRs was explored using single-cell Ca2+-imaging experiments in HEK293 cells.

In these experiments, we utilised PAR2’s promiscuous signalling through Gαi/o and Gαq and

A2B receptor signalling through Gαs. Currently there are no agonists that can reversibly activate TRPM2 that can be applied externally. We therefore utilised H2O2 which leads to irreversible activation of TRPM2 and heat ramps which reversibly activate the channel.

Application of PAR2 and A2B receptor agonists had no effect on H2O2 activation of TRPM2.

However, since H2O2 activation of TRPM2 is irreversible, this could prevent modulation of the channel by G-proteins and their signalling pathways. We then applied two subsequent heat ramps on TRPM2 where its activity appeared to be fully desensitised following the first ramp. Application of A2B agonists did not affect the Ca2+-response to the second ramp. This indicates that activation of GPCRs that couple to Gαs does not resensitise the channel. Since the response to the second heat ramp is fully desensitised, we cannot claim for certain whether GPCR signalling affects TRPM2 activity. Future experiments should attempt to

221 apply control solution and GPCR agonists prior to the first heat ramp and observe whether

GPCR agonists influence the initial TRPM2-mediated heat response. Furthermore, TRPM2- transfected cells can be patch-clamped and supplied internally with reversible TRPM2 activators. GPCR agonists can then be applied by the superfusion system and any effects on

TRPM2-mediated currents recorded.

TRPM7 was recently shown to be reversibly and externally activated by the DOR antagonist naltriben. Activation of PAR2 had no effect on TRPM7-mediated Ca2+-responses thus suggesting that Gαi/o- and Gαq-mediated signalling does not interfere with TRPM7 activity. In contrast, activation of A2B receptors led to a significant and reversible increase in TRPM7- mediated responses. This sensitisation was present in the absence of naltriben, was not reliant on PKA and was unaffected by co-expressing the Gβγ sink βARK1-ct. These results indicate that TRPM7 may be sensitised by Gαs directly binding to the channel. To confirm this, future experiments can utilise the inside-out patch-clamp technique where GTPγS- activated Gαs can be applied directly onto the intracellular side of TRPM7.

With TRPM8, only activation of PAR2 inhibited the channel in a Gαq-dependent manner as

PTX treatment did not affect this. This data agrees with previously published literature that

Gαq protein binding to TRPM8, and possibly Gαq-mediated PIP2 hydrolysis, inhibit TRPM8 activity (Zhang et al., 2012; Zhang, 2019b). Co-expression of βARK1-ct did not prevent this inhibition. These results suggest that Gαi/o, Gαs and Gβγ do not affect TRPM8 activity.

Future work should also examine whether TRPM1 and TRPM4-6 can also be affected by G- proteins. Since TRPM1 has been shown to be inhibited by both Gαo and Gβγ, it is at least likely that it will be promiscuously inhibited by Gβγ released from other Gα subunits, as is the case with TRPM3. Direct G-protein binding to TRPM6 has not been studied and could be

222 a possibility since its closely related channel, TRPM7, appears to be activated by Gαs proteins.

The TRPM family is involved in several physiological processes and it appears that numerous members are directly modulated by G-proteins. This provides an exciting avenue into understanding the diverse relationship between GPCRs and ion channels which should be further explored.

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