The Pennsylvania State University

The Graduate School

Intercollege Program of Neuroscience

STRUCTURE-FUNCTION RELATIONSHIPS AND BIOCHEMICAL

MANIPULATION OF 6-TM SUPERFAMILY ION CHANNELS

A Dissertation in

Neuroscience

by

Aditya Pisupati

 2018 Aditya Pisupati

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

December 2018

The dissertation of Aditya Pisupati was reviewed and approved* by the following:

Timothy Jegla Associate Professor of Biology Dissertation Advisor Chair of Committee

Melissa Rolls Professor of Biochemistry and Molecular Biology

Bernhard Luscher Professor of Biology, Biochemistry and Molecular Biology, Psychiatry

Santhosh Girirajan Associate Professor of Biochemistry and Molecular Biology

William Hancock Professor of Bioengineering

Kevin Alloway Professor of Neural and Behavioral Sciences Co-Director of Graduate Program in Neuroscience

*Signatures are on file in the Graduate School

ABSTRACT

The six-transmembrane (6-TM) superfamily of ion channels is a diverse group of tetramer-forming ion channels. Members of this superfamily are regulated by a many different physical and chemical stimuli, including (but not limited to): organic compounds, temperature, mechanical stress, and transmembrane potential. 6-TM channels are critical for regulating neuronal excitability, cardiac pacemaking, sensing of thermal and mechanical stimuli, and maintaining water balance in plants among many other functions. In this thesis, I will first present an overview of 6-TM containing channels. This will be followed by three different projects involving channels from three families within the 6-TM superfamily: Shaker, Transient Receptor

Potential Vanilloid (TRPV), and Cyclic Nucleotide Binding Domain (CNBD) channels. The project involving Shaker family channels identifies the stoichiometry of an obligatory heteromeric subunit and determines that this stoichiometry is regulated by a mechanism involving the activation gate. In the TRPV project, C. elegans and D. melanogaster TRPV channels are found to be activated by the vitamin B3 metabolite nicotinamide. In the investigations of CNBD containing channels, it is determined that heme appears to be an evolutionarily conserved

regulator of CNBD-containing channels in both plant and animal lineages. Finally, I will discuss

where these works fit into our current understanding of ion channels and physiology and will

speculate on the outcome of new questions posed by the results of these three projects.

iv

TABLE OF CONTENTS

List of Figures ...... vii

List of Tables ...... x

Acknowledgements ...... xi

Chapter 1 Background on 6-TM Ion Channels ...... 1

Ion channel gating ...... 1 Biological significance of 6-TM ion channels ...... 5 Shaker family channels ...... 7 TRPV family channels ...... 12 CNBD-containing family channels ...... 14 Preview of Thesis ...... 22 References ...... 24

Chapter 2 Experimental techniques for probing ion channel structure-function relationships ...... 32

Xenopus laevis oocytes as a model for studying ion channels ...... 32 Preparation of Xenopus oocytes for cRNA injections ...... 34 Two-Electrode Voltage Clamp (TEVC) ...... 35 How to perform Two-Electrode Voltage Clamp ...... 37 Analysis of TEVC Data ...... 38 Total Internal Reflectance Fluorescence (TIRF) Microscopy ...... 40 Performing TIRF Microscopy Experiments ...... 40 Analysis of TIRF data ...... 42 References ...... 44

Chapter 3 S6 gate divergence in regulatory subunits restricts the stoichiometry of heteromeric Shaker family channels ...... 45

Abstract ...... 46 Introduction ...... 47 Methods ...... 51 Molecular Cloning...... 51 cRNA synthesis and Xenopus oocyte preparation ...... 53 Electrophysiology...... 54 TIRF microscopy...... 55 Sequence Alignments ...... 56 Molecular Modeling ...... 56 Results ...... 57 The highly-conserved S6 activation gate sequence degenerates in regulatory subunits ...... 57 TIRF microscopy reveals multiple Kv2.1:Kv6.4 heteromer stoichiometries ...... 61

v The S6 gate of Kv6.4 limits formation of 2:2R heteromers ...... 64 The Kv6.4 gate blocks function of 2:2R heteromers ...... 67 Cd2+ block of cysteine mutants confirms 2:2R functionality of Kv6.4-PIPIIV- Kv2.1CT ...... 74 Discussion ...... 77 References ...... 87

Chapter 4 Activation of invertebrate TRPV heterotetramers by the vitamin B3 metabolite nicotinamide ...... 91

Introduction ...... 92 Methods ...... 93 Molecular cloning and cRNA synthesis ...... 93 Xenopus oocyte recordings ...... 93 TIRF Photobleaching ...... 93 Sequence Alignment ...... 94 Results ...... 94 Nicotinamide is an OSM9/OCR4 channel agonist ...... 94 Nicotinamide-evoked activity is conserved in the Drosophila TRPV homolog formed by Inactive and Nanchung ...... 95 OSM9 and OCR4 form a 2:2 heterotetramer ...... 96 Discussion ...... 98 Nicotinamide as a potential endogenous ligand for invertebrate TRPV channels ... 99 Phylogenetic analysis of TRPV evolution...... 100 The structural basis of interdependent heteromerization...... 102 References ...... 105

Chapter 5 Heme modulation of CNBD-containing Ion Channels ...... 107

Introduction ...... 107 Methods ...... 112 Molecular Cloning...... 112 cRNA synthesis ...... 113 Electrophysiology...... 113 Sequence Alignments ...... 114 Results ...... 114 The heme effect in human Elk1 channels is distinct from mode-shift ...... 114 The left-shift observed in human Elk1 channels is dependent on iron-containing heme moieties and is reversed in the presence of divalent cations ...... 115 Hemin leads to stabilization of the VSD-out state of Plant CNBD-containing channels ...... 117 The conserved S5 histidine is necessary, but not sufficient, for the heme effect in EAG family ...... 118 Discussion ...... 120 References ...... 124

Chapter 6 Summary, Biological Applications of Findings, and Future Directions ...... 126

Summary of Experiments ...... 126

vi The development and testing of a novel model for obligatory heteromeric stoichiometry formation ...... 126 Identification of a novel invertebrate TRPV ligand and exploration of invertebrate TRPV assembly ...... 127 Characterization of a conserved heme-channel interaction within the CNBD- containing channel family ...... 127 Biological Significance and Potential Applications of Findings...... 128 Restricted heteromeric stoichiometry prevents regulatory subunit mixing ...... 128 Invertebrate TRPV channels as an in vivo nicotinamide biosensor ...... 128 The interaction between heme and CNBD-containing channels ...... 129 Future Directions ...... 131 Testing the gate regulation model in other 6-TM channels ...... 131 Investigating the origins of nicotinamide sensitivity in TRPV channels ...... 132 Determining the mechanism and potential roles of the heme-CNBD interaction .... 133 References ...... 139

vii LIST OF FIGURES

Figure 1-1. The role of ion channels in the generation of an action potential...... 2

Figure 1-2. Biophysical states occupied by a gated ion channel...... 4

Figure 1-3. Schematic of a 6-TM ion channel subunit...... 5

Figure 1-4. Phylogenetic relationship of channels utilized in this Thesis...... 7

Figure 1-5. General gating model of Shaker family channels...... 9

Figure 1-6. The N-terminal T1 domain determines homomeric and heteromeric channel assembly of Shaker family channels...... 11

Figure 1-7. CNBD-containing channels share similar tertiary structure to Shaker family but are gated distinctly...... 16

Figure 1-8. Summary of modulators of Voltage-gated CNBD-containing channels...... 17

Figure 1-9. Schematic of the conserved residues within the transmembrane core of CNBD-containing ion channels...... 17

Figure 2-1. Injection of Xenopus oocyte with ion channel encoding cRNA...... 33

Figure 2-2. Schematic of Two-Electrode Voltage Clamp (TEVC) using a Xenopus oocyte...... 36

Figure 2-3. Example analysis of Two Electrode Voltage Clamp Data...... 39

Figure 2-4. Using Total Internal Reflectance Fluorescence Microscopy (TIRFM) to identify ion channel stoichiometry...... 41

Figure 2-5. Example photobleaching data acquired by Total Internal Reflectance Fluorescence Microscopy...... 42

Figure 3-1. Tetrameric Shaker family K+ channels have two major intersubunit interfaces. .. 48

Figure 3-2. The S6 activation gate sequence differs from the Shaker family consensus in evolutionarily-independent regulatory subunit groups...... 59

Figure 3-3. Comparison of gating properties of Kv2.1 homomers and Kv2.1:Kv6.4 heteromers...... 60

Figure 3-4. Determination of Kv2.1:Kv6.4 heteromer stoichiometry by TIRF photobleaching assay...... 63

Figure 3-5. The Kv6.4 activation gate but not T1 limits the formation of Kv2.1:Kv6.4 2:2R heteromers...... 66

viii Figure 3-6. Biophysical properties of heteromeric currents from oocytes co-expressing Kv2.1:Kv6.4-PIPIIV-Kv2.1CT...... 68

Figure 3-7. Predicted effect of Kv2.1:Kv6.4 expression ratio on the number of functional channels for models in which (A) only 3:1R heteromers conduct or (B) both 3:1R and 2:2R heteromers conduct...... 70

Figure 3-8. Kv2.1:Kv6.4 expression ratio titrations predict that 2:2R heteromers have negligible conductance for Kv6.4 WT and conducting for Kv6.4PIPIIV-Kv2.1CT...... 72

Figure 3-9. Cd2+ block mediated by adjacent Kv2.1 I383C subunits predicts that 2:2R heteromers contribute to whole cell currents for Kv2.1:Kv6.4-PIPIIV-Kv2.1CT but not for Kv2.1:Kv6.4 WT...... 75

Figure 3-10. Structural homology models of Kv2.1 homomers and Kv2.1:6.4 heteromers suggest the Kv6.4 activation gate might perturb pore stability and function...... 79

Figure 3-11. Model for the evolution of Shaker family regulatory subunits adopting a 3:1R functional stoichiometry...... 83

Figure 4-1. Nicotinamide activates the heteromeric TRPV channel formed by the co- expression of OCR4 and OSM9 protein subunits in Xenopus laevis oocytes...... 95

Figure 4-2. Fly TRP channel subunits Inactive (IAV) and Nanchung (NAN) make a heteromeric ion channel that is activated by nicotinamide...... 96

Figure 4-3. TIRF photobleaching demonstrates channels that traffic to the membrane when OSM9 and OCR4 are coexpressed, but not individually...... 97

Figure 4-4. OSM9/OCR4 channels predominantly form 2:2 heteromers in Xenopus...... 98

Figure 4-5. TRPV subunits from the cnidarian Nematostella vectensis cluster with invertebrate TRPV clade...... 101

Figure 4-6. Disruption of the extracellular gate could be the mechanism of obligatory heteromerization among invertebrate TRPV channels...... 103

Figure 5-1. The highly conserved HxxxC motif in CNBD-containing ion channels does not have any currently known functions...... 108

Figure 5-2. Heme types are determined by differences in functional groups attached to the central porphyrin ring...... 109

Figure 5-3. Heme is a potent inhibitor of Nematostella vectensis HCN1 channels and an activator of human Elk1 channels...... 111

Figure 5-4. Heme and mode-shift are additive processes on Elk1...... 115

Figure 5-5. Metal substituted heme moieties and divalent cations prevent the heme induced left-shift in HsElk1...... 117

ix Figure 5-6. Outward voltage sensor stabilization by heme is conserved in Plant CNBD- containing channels...... 118

Figure 5-7. The heme effect can be reduced by mutating key residues in the pore domain of HsElk1 channels and the voltage sensor of GORK channels...... 119

Figure 5-8. Heme-stabilization of the VSD-out conformation in depolarization and hyperpolarization gated channels...... 122

Figure 6-1. Predicted effects of heme exposure on plant guard cells...... 130

Figure 6-2. Voltage Clamp Fluorometry could be used to identify the mechanism of channel modulation by heme...... 134

Figure 6-3. Nitric Oxide has heterogeneous effects on heme-shifted CNBD-containing channels...... 136

Figure 6-4. Elk1 D261A appears to be less responsive to hemin...... 137

x LIST OF TABLES

Table 3-1. Boltzmann Fit Paramaters of Kv2.1 and Kv2.1:Kv6.4 heteromers...... 86

xi ACKNOWLEDGEMENTS

There are many people that should be acknowledged for having a hand in the creation of this Dissertation; perhaps too many to list in the little space I have available. However, it would be a great disservice to not mention the following people, without whom none of this would be possible. First and foremost, I would like to thank my parents for giving me their blessings as I pursued my passions. I would also like to thank my brother; through your experiences and discussions, I’ve been able to envision a world more expansive than what exists in my own bubble. With regards to career development, I would first like to thank my advisor, Tim Jegla, for giving me the opportunity to develop into the scientist that I am today. You challenged me to scrutinize every detail to make the best arguments I possibly can, while simultaneously focusing on the bigger picture to look at how my work fits into the broader academic community. I would also like to acknowledge my Committee Members for their thoughts on how to frame this dissertation and for their advice on how to develop into a successful scientist. I would like to thank the following people who have contributed to my technical development as a scientist and academic: Xiaofan Li for tolerating my many questions as I learned valuable skills in the lab;

Keith Mickolajczyk and Awani Upadhyay for their ideas and eagerness to collaborate on projects;

Jessica Eason and Yunqing Zhou for allowing me to pass on my knowledge to a new generation in the lab; and to Greg Busey for inspiring in me a new perspective with which to solve problems.

Finally, I would like to thank the Penn State Medical Scientist Training Program for granting me the privilege to pursue dual degree training at this storied institution. I will be forever grateful for the mentorship I have received and the friendships I have made along the way.

This research was performed in part using funds from NSF (Award number 1621027) and

NIH (Award number R21NS093477). The findings and conclusions do not necessarily reflect the views of the National Science Foundation or the National Institutes of Health.

Chapter 1

Background on 6-TM Ion Channels

In this chapter, I will give an overview of the terminology and concepts involved in ion channel research, with a particular emphasis on the 6-TM channels that I will later discuss. Specifically, I will address: the concept of ion channel gating, and the biological significance 6-TM channels—specifically from Shaker, TRPV and

CNBD families—along with how my projects aim to solve open problems in the field. Finally, I will give a preview of the experiments on these various channels in my Thesis.

Ion channel gating

The nonpolar nature of the hydrocarbon tail groups within a lipid bilayer allows hydrophobic molecules to readily cross between the cytosol and the extracellular environment. However, this same physical property of lipid membranes renders the bilayer impermeable to polar molecules and ions. This poses a conundrum, as the movement of ions between the two compartments is vital for cell physiology and metabolism. To facilitate ion transport between the intracellular and extracellular compartments, organisms from all domains of life have a repertoire of membrane spanning proteins, called “ion channels”. By definition, ion channels only allow for the passive movement of ions; the driving force for ionic movement is the electrochemical gradient of the ions that the channels can permeate. While the movement of the ions through a channel is passive, the opening and closing of the channel typically requires some form energy, whether it is in the form of chemical energy (e.g. ligand binding), thermal energy, mechanical energy or electrical energy. The act of opening or closing of a channel is referred to as the gating of that channel, with the specific stimuli that can gate the channel being intrinsic to the channel.

2 Channels gated by electrical energy, which are the major focus of this Thesis, are also termed “voltage-

gated ion channels”. These channels are necessary for many different physiological processes. A simple example

to demonstrate the importance of voltage-gated ion channels is the classic action potential (Figure 1-1). An action

potential is a rapid depolarization of the cell membrane exhibited by excitable cells such as neurons. Neuronal

action potentials are used to transmit signals to other neurons or to effector cells such as muscle cells. The

initiation, propagation and termination of an action potential requires the activity of several different types of

voltage-gated ion channels; furthermore, the shape of the waveform and frequency of firing are also determined

by voltage-gated ion channel activity (Bean, 2007).

Figure 1-1. The role of ion channels in the generation of an action potential. The schematic on the right shows an action potential waveform with the distinct components labeled A-E. Each of the components of action potential requires contributions from several different types of gated ion channels. A) Excitatory post-synaptic currents (EPSCs) originating in the dendrites propagate through the soma to the proximal part of the axon, which is termed the Axon Initial Segment (AIS), and lead to membrane depolarization. These EPSCs typically originate from the opening of ligand-gated ion channels belonging to the excitatory ionotropic glutamatergic receptor family (iGluR). B) Once the membrane at the AIS is depolarized to the threshold voltage (VThreshold), Voltage-gated Sodium channels (NaV) open to allow rapid membrane depolarization. This rapid depolarization can propagate down the axon through the opening of more NaV channels at points further along the axon. Propagation of the action potential into the pre-synaptic terminals leads to synaptic vesicle fusion and neurotransmitter release. C) Inactivation of Voltage-gated Sodium channels causes the action potential waveform to peak. D) Opening of Voltage-gated Potassium channels (Kv) leads to the repolarization phase of the action potential. E) Membrane potential returns to the resting membrane potential (VRest) due to the activity of Kv channels. Currents from Kv channels can often lead to a membrane hyperpolarization beyond VRest that dissipates upon closure of these channels.

As shown in Figure 1-1 channels can generate rapid responses to changes in the cellular environment.

From this, it is easy to see how aberrations in channel function may lead to disease. Searching the word “channel”

3 in the Online Mendelian Inheritance of Man (OMIM) database results in 153 known diseases that are the result of ion channel mutations. Indeed, diseases caused by mutations in channel activity, also known as channelopathies, were some of the earliest diseases to be mapped within the human genome; these diseases include Cystic Fibrosis,

Hyperkalemic Periodic Paralysis, and Malignant Hyperthermia (Cutting et al., 1990; Fontaine et al., 1990;

MacLennan et al., 1990). Channelopathies can lead to disease by affecting the ability of a channel to assemble, traffic, and of course, gate (Ashcroft, 2006). Determining how mutations can influence an ion channel’s function would therefore lead to an understanding of how certain pathologies can arise, which would then be useful for developing treatment strategies for these pathologies. Prior to determining how specific mutations in channels lead to diseases, it is first important to understand how channels behave physiologically.

In general, gated channels can occupy one of three states at any given time: activated, deactivated, and inactivated (Figure 1-2). Activated channels are channels that have received the appropriate stimulus required to open the gate. It follows then, that deactivated channels are channels for which the activation stimulus is simply reversed. Inactivation of a channel, however, refers to a state from which the channel cannot activate, regardless of the magnitude of the stimulus that the channel receives. This is due to the closure of an independent inactivation gate. Once a channel is inactive, the inactivation gate must be re-opened before it can be activated again. Rates of activation and inactivation of ion channels can vary quite significantly. This is because the strength of coupling between the activation stimulus and the different gates is highly heterogeneous. An organism can therefore utilize several gated channels from the same family for different physiological purposes.

4

Figure 1-2. Biophysical states occupied by a gated ion channel. The activation gate is shown in red, and the inactivation gate is shown in blue. Opening and closing of the activation gate, while the inactivation gate is open, leads to transitioning between the deactivated state and activated state. Once the inactivation gate is closed, the inactivation must be removed and the channel must enter into the deactivated state prior to re-activation; channels cannot activate from an inactivated state. The number of activation and inactivation gates can vary based on the channel being studied. The rates of transitions between these states can vary based on the strength of the stimulus applied.

The classic example of an ion channel that displays activated, deactivated and inactivated states is the

Voltage-gated Sodium channel. As shown in Figure 1-1, this channel is crucial for the rising phase of the action

potential in many neurons. The behavior of the Voltage-gated Sodium channel was first described by Hodgkin

and Huxley in their famed experiments on the squid giant axon (Hodgkin and Huxley, 1952a, b). In this

experiment, the membrane potential of the squid giant axon was controlled by a Two-Electrode Voltage Clamp

(see Chapter 2) with one electrode placed in the extracellular bath, and the other electrode placed inside the axon.

By manipulating the concentrations of ions in the extracellular bath and the intracellular milieu, Hodgkin and

Huxley were able to demonstrate that the depolarization phase of the action potential is the result of a rapidly activating sodium conductance that subsequently inactivates. Subsequent deactivation of the sodium conductance occurs with membrane repolarizations. Remarkably, this characterization of the voltage-gated sodium channel was made almost 40 years prior to the cloning of the first Voltage-gated Sodium channel (Noda et al., 1986).

5 Biological significance of 6-TM ion channels

Before discussing how my Thesis works to solve problems in the field of six-transmembrane domain (6-

TM) ion channels, it is important to first understand what is currently known about this large, diverse superfamily.

All members of the 6-TM superfamily are made up of an intracellular N-terminal domain, followed by six

transmembrane domains—the last two of which always contribute to the pore domain—followed by an

intracellular C-terminal domain (Figure 1-3). Evolutionarily, 6-TM channels likely arose when channels from the

2-TM family combined with 4-TM voltage sensitive proteins. The evidence for this hypothesis stems from the existence of 2-TM channels in the inward-rectifying channel family (Ho et al., 1993; Kubo et al., 1993; Whorton and MacKinnon, 2011), and 4-TM voltage sensitive proteins that can themselves function as channels and enzymes (Murata et al., 2005; Ramsey et al., 2006; Sasaki et al., 2006). The common ancestor of all 6-TM channels was therefore voltage sensitive, and possibly voltage-gated. As will be shown below, currently known channels in the 6-TM family are known to be gated by many stimuli and have variable levels of voltage sensitivity.

Figure 1-3. Schematic of a 6-TM ion channel subunit. All subunits in the 6-TM family consist of variable intracellular N- and C- termini. The six transmembrane domains are enumerated as S1-S6. The pore domain (PD) is made up of the S5 and S6 domains with the large linker between these two domains; the PDs of all four subunits lines the permeation pathway for the ions to traverse the membrane. Because of its importance in ion permeation, the linker region between S5 and S6 is often termed the pore loop, or P-loop. The S1-S4 domains collectively make up the voltage sensor domain (VSD). A series of highly conserved positively charged residues in the S4 domain is thought to be the major detector of changes in membrane potential. Although not all 6-TM channels are voltage-gated, voltage sensing was likely present in the common ancestor of 6-TM channels, and voltage sensitivity is conserved in many 6-TM channels.

6

Functional 6-TM channels are cation selective channels that are typically assembled with four individual

subunits combining in a family dependent way; thus, most 6-TM channels are considered tetrameric. The only

exceptions to tetrameric assembly in the 6-TM superfamily are: the Voltage-gated Sodium channel (NaV) family

and the Voltage-gated Calcium channel (CaV) family, which are monomers (Payandeh et al., 2011; Wu et al.,

2015); and the Two-Pore Channel (TPC) family, which are dimers (Ishibashi et al., 2000). These various channels

are referred to as “pseudo”-tetramers due to the fact that they consist of tandem linkages of two repeats of 6-TM

subunits for the dimeric TPCs and four repeats of 6-TM subunits for the NaV and CaV channels. The ancestral 6-

TM channel was therefore likely a true tetramer.

6-TM channels are structurally and functionally distinct from other ion channel superfamilies. Examples

of other non-6-TM superfamilies include the pentameric Cys-loop receptor superfamily and the trimeric

Degenerin/Epithelial Sodium Channel (Deg/ENaC) superfamily (Ashcroft, 2006). Whereas 6-TM subunits are

characterized by the signature six-transmembrane domain sequence, Cys-loop receptor subunits have four transmembrane helices and Deg/ENaC subunits contain only two transmembrane helices (Benos and Stanton,

1999; Sine and Engel, 2006). In addition to these structural differences, 6-TM channels are functionally very distinct from Cys-loop receptors and Deg/ENaCs. As mentioned above, voltage sensitivity is a hallmark of 6-TM channels. In contrast, Cys-loop receptors are primarily ligand-gated ion channels; two of the most well-known classes of Cys-loop receptors are the Nicotinic Acetylcholine (nAch) and GABAA receptors (Sine and Engel,

2006). Deg/ENaCs meanwhile are predominantly gated by mechanical or chemical stimuli (Benos and Stanton,

1999).

The channels that will be discussed in this Thesis belong to the following families within the 6-TM

superfamily: Shaker family, Transient Receptor Potential Vanilloid (TRPV) family, and cyclic nucleotide binding

domain (CNBD) containing family (see Figure 1-4). Shaker family consists of four major subfamilies:

Kv1/Shaker, Kv2/Shab, Kv3/Shaw, and Kv4/Shal. As will be discussed below, there also exist various expansions

of regulatory subfamilies (KvR). The TRPV family is actually a subclass of the broader TRP channel family.

However, for the purposes of this Thesis, discussion will be limited to TRPV structure-function and differences

7 between the invertebrate lineage and the vertebrate lineage. The CNBD-containing family is composed of the

Ether-A-Go-Go (EAG) family, Hyperpolarization-activated Cylic Nucleotide-gated (HCN) family, and the

Plant CNBD-containing channels. The Cyclic Nucleotide Gated (CNG) channel family also belongs to the

CNBD-containing clade, however these channels are not used in this Thesis and will not be discussed. Though

Shaker, TRPV and CNBD-containing channels all belong to the 6-TM superfamily, these families diverged going

back to prokaryotes (Jegla et al., 2009). As will be highlighted below, these channels are gated and regulated

through diverse mechanisms, but all are important for neuronal signaling.

Figure 1-4. Phylogenetic relationship of channels utilized in this Thesis. Channels that are analyzed in this Thesis belong to the subfamilies circled in blue. Shaker, TRPV and CNBD families all have varying levels of voltage sensitivity but are all tetrameric. Shaker family channels all open with depolarization of the membrane and are potassium-selective. TRPV channels non-specifically conduct cations; they may have some voltage-sensitivity but are primarily polymodal sensory channels. They can be gated by: ligand binding, osmomechanical changes, and temperature changes depending on the lineage of the channel being considered. CNBD containing channels have highly heterogenous responses to voltage. EAG family channels, like Shaker family, are all opened by depolarization and are potassium-selective. The Plant CNBD channels GORK and SKOR share this phenotype. By contrast, HCN family channels, as well as KAT and AKT, open when the membrane is hyperpolarized. HCN channels are non-specific cation channels, whereas KAT and AKT are potassium-selective. CNG channels are exclusively opened by cyclic nucleotide binding.

Shaker family channels

Shaker channels are a large group of channels in metazoans that are activated by depolarization of the membrane; thus, these channels are all voltage-gated channels. These channels are known to be potassium selective due to the presence of a near universally conserved selectivity filter in the pore-loop (Long et al., 2005).

8 Within the Shaker family, there are four subfamilies that are common to animals that can form homomeric

channels: Shaker (Kv1), Shab (Kv2), Shaw (Kv3), and Shal (Kv4) (Butler et al., 1989; Butler et al., 1990). The

name Shaker stems from the fact that mutations in the founding member of the gene family, Shaker, lead to a

distinct leg-shaking phenotype in Drosophila (Salkoff, 1983; Kamb et al., 1987; Papazian et al., 1987).

Although the precise structural basis of all of the transitions that Shaker family channels undergo in order

to activate, deactivate and inactivate are still being elucidated, the biophysical characterization of these channels

has been a matter of intense study. A general gating model of the types of transitions that Shaker channels can

undergo is shown in Figure 1-5. Activation of Shaker channels requires extracellular, or outward, movement of

the voltage sensor domains (VSDs) of each subunit, which is then coupled to the pore. Outward movement of the

VSDs occurs in at least two steps; during the first step of the VSD-out transition, voltage sensors move independently. The second step is cooperative movement of the VSDs (Zagotta et al., 1994; Mannuzzu and

Isacoff, 2000). Opening of the gate occurs through an as yet unclear mechanism, but is thought to involve interactions between the S4-S5 linker and the distal S6 gate (Pathak et al., 2005). Inactivation of Shaker channels can occur in two ways: the well-characterized N-type inactivation, and the less well understood C-type inactivation (Hoshi et al., 1990; Zagotta et al., 1990; Hoshi et al., 1991; Hoshi and Armstrong, 2013). As shown in

Figure 1-5, N-type inactivation, which is also known as “ball-and-chain” inactivation, occurs when a portion of the cytoplasmic N-terminus occludes the pore. C-type inactivation is not as well characterized, however recently published structures suggest that this occurs due to collapse of the extracellular portion of the permeation pathway

(Pau et al., 2017).

9

Figure 1-5. General gating model of Shaker family channels. Activation of channels requires outward movement of the S4 domains (shown in white), each of which can move independently. S4 domains can only move outward through depolarization of the membrane. Pore opening is a concerted effort between all four subunits and only occurs once all four S4 domains are in the appropriate outward position. Once a channel is active, it can deactivate through repolarization of the membrane, or it can enter into one of two inactivated states. N-type inactivation occurs when a portion of the cytoplasmic N-terminus occludes the pore. C-type inactivation occurs through more sophisticated mechanisms and may involve collapse of the permeation pathway.

The physiological role of Shaker channels relies on the biophysical properties of each specific channel, which in turn is dependent on the lineage of the specific channel in question. Generally, currents generated by

Shaker channels can be classified as “fast-inactivating” or “delayed-rectifying”. The term “delayed-rectifier” originated from the fact that these types of currents activate upon depolarization somewhat more slowly compared

Voltage-gated Sodium channels (Hille, 1992); delayed-rectifying channels do not significantly inactivate at physiologically relevant timescales. Fast-inactivating type channels, as the name suggests, rapidly inactivate

10 following upon strong depolarization. Currents from fast-inactivating channels are sometimes termed IA, after the

genus Anisodoris in which these types of currents were first characterized (Connor and Stevens, 1971); these

types of channels typically regulate subthreshold excitability.

Drosophila Shaker channels display a fast-inactivating current; it was the loss of this inactivating

potassium current, specifically at the neuromuscular junction, that underlies the leg-shaking phenotype that gives

rise to the name of the Shaker family. (Salkoff and Wyman, 1981; Salkoff, 1983; Wu et al., 1983). Shab channels

are non-inactivating and appear to play a role in determining action potential frequency and synaptic transmission

(Ueda and Wu, 2006; Peng and Wu, 2007). Shaw channels are non-inactivating and appear to be important to regulate resting membrane potential (Tsunoda and Salkoff, 1995). Drosophila Shal channels are also fast-

inactivating, much like Shaker, however they tend to open at more hyperpolarized potentials (Ping et al., 2011).

Though the currents generated by the mammalian Shaker lineage are typically similar to their Drosophila

homologs, there is prominent divergence within certain subfamilies. Mammalian Kv2 channels encode a delayed-

rectifying current much like their Drosophila homolog; these channels are expressed in virtually all neurons and

play a key role in regulating various aspects of neuronal excitability (Malin and Nerbonne, 2002; Misonou et al.,

2005). Mammalian Kv4 channels, likewise, display fast-inactivation similar to Drosophila Shal channels, and also appear to play a role in regulating action potential threshold (Sheng et al., 1992; Serodio et al., 1996). Mammalian

Kv1 channels, however, predominantly display delayed-rectifier type currents with little inactivation (Christie et al., 1989). Although it should be noted that two mammalian Kv1 channels, Kv1.4 and Kv1.7, display fast- inactivation similar to Drosophila (Sheng et al., 1992; Kalman et al., 1998), so fast-inactivating Shaker currents are still somewhat conserved in mammals. Mammalian Kv3 channels display both delayed-rectifier and fast- inactivating type currents and appear to play a role in determining neuronal firing rate (Massengill et al., 1997;

Waters et al., 2006).

Classically, Shaker family channels have been studied as homomers, however it is known that subunits can combine in a subfamily specific way to generate heteromers that have novel biophysical properties

(Covarrubias et al., 1991). This subfamily specificity of channel assembly is due to a well-characterized N-

11 terminal region that is aptly named the Tetramerization domain, or T1 (Shen and Pfaffinger, 1995; Xu et al.,

1995; Kreusch et al., 1998). The mechanism by which the T1 domain regulates subfamily specificity is through

direct contact of adjacent subunits (Kreusch et al., 1998; Robinson and Deutsch, 2005); in this way, “like”

subunits associate with other “like” subunits (Figure 1-6). It should be noted that the T1 mechanism of channel

assembly is actually unique to Shaker family; as will be discussed below, the cytoplasmic C-terminus plays a

more prominent role in assembly of other 6-TM channels. Though the specific combinations of subunits, and their

respective stoichiometries, that can form in vivo is not clear, heteromeric channels are likely more physiologically

relevant than the homomeric channels that are typically studied (Coleman et al., 1999; Plane et al., 2005).

Figure 1-6. The N-terminal T1 domain determines homomeric and heteromeric channel assembly of Shaker family channels. A) T1 domains of subunits from the same subfamily can combine in a lock-and-key type mechanism. In this way, the subfamily represented with the blue T1 can combine with other blue T1-containing subunits and red T1-containing subunits can combine with other red T1-containing subunits B) Subunits from different subfamilies exclude each other due to incompatibility at the interface. Therefore, blue T1-containing subunits cannot combine with red T1-containing subunits to form a heteromer.

Interestingly, there are several independent expansions of so-called “regulatory” subunits present within

the Shaker family. These regulatory subunits cannot form homomeric channels, but rather combine with

homomer-forming subunits in order to form obligatory heteromers (Post et al., 1996; Jegla and Salkoff, 1997;

Kramer et al., 1998; Ottschytsch et al., 2002; Li et al., 2015b). This regulatory phenotype is the result of a self- incompatible T1 domain, which can be rescued when homomer-forming subunits with a compatible T1 are expressed (Kramer et al., 1998; Ottschytsch et al., 2002). Heteromers formed by coexpression of regulatory

12 subunits with homomer-forming subunits have biophysical characteristics that are distinct from the corresponding

homomeric channels (Kerschensteiner and Stocker, 1999; Stocker et al., 1999; Ottschytsch et al., 2005).

The major expansion of regulatory subunits in mammals is made up of the Kv5, Kv6, Kv8 and Kv9

subfamilies, and these are all known to combine with Kv2 subunits. These regulatory subunits are expressed in a

cell-type specific manner and display distinct biophysical properties that influence their physiological function

(Bocksteins, 2016). Mammalian obligatory heteromers for which the stoichiometry is known consistently display

only one regulatory subunit per functional tetramer (Kerschensteiner et al., 2005; Bocksteins et al., 2017). This

poses a conundrum, as the T1 domain alone would not be able to prevent a second regulatory subunit from being

inserted into a functional tetramer (Figure 1-6); this suggests that other portions of the channel are also important

for determining channel stoichiometry.

The project presented in Chapter 3 of this Thesis was done to test the hypothesis that the other major

contributor to subunit stoichiometry in mammalian regulatory subunits is the activation gate. This was predicted

based on the fact that the gate region is the only consistently divergent portion of the various regulatory lineages.

The stoichiometry of the mouse Kv2.1/Kv6.4 channel was chosen for study due to its importance in fast-spiking neurons, particularly those found in the spinal cord (Muller et al., 2014). Moreover, the Kv2.1/Kv6.4 heterotetrameric channels display a closed-state inactivation phenotype not found in the Kv2.1 homotetramer

(Bocksteins et al., 2012), which allows for easier discrimination of homomeric and heteromeric currents. The results from this project give rise to a model by which the activation gate of regulatory subunits from any evolutionary expansion could change over time to determine channel stoichiometry.

TRPV family channels

Transient Receptor Potential Vanilloid (TRPV) channels are 6-TM channels that serve to primarily transduce physical changes in the environment, although they also can be opened through chemical binding

(Caterina et al., 1999; Montell, 2005). Vertebrate TRPV channels are celebrated for their role in heat sensation,

13 however vertebrate TRPV4 channels appear to be unique in that they also appear to be gated by osmotic or

mechanical changes (Caterina et al., 1999; Liedtke and Friedman, 2003; Vriens et al., 2004). The biophysical

underpinnings of thermosensitive and mechanosensitive gating among these channels are still unclear.

Structurally, TRPV subunits contain anywhere between three and five ankyrin repeats in the N-terminus (Montell,

2005). TRPV channels also have a have a signature sequence in the C-terminus referred to as the “TRP box”, which appears to be important for gating in general (Montell, 2005; Cohen and Moiseenkova-Bell, 2014).

Tetrameric assembly of vertebrate TRPV channels is known to occur through a coiled-coiled domain distal the

TRP box (Tsuruda et al., 2006; Zhang et al., 2011).

Mechanosensitive gating among vertebrate TRPV4 channels appears to require interactions with cytoskeletal proteins, such as actin, and may also require PIP2 interactions with the N-terminal tail (Ramadass et

al., 2007; Becker et al., 2009; Garcia-Elias et al., 2013). The nature of the temperature sensor in vertebrate TRPV

channels is still a matter of controversy. Some studies suggest that portions of the N-terminus are responsible for temperature sensitivity (Yao et al., 2011; Liu and Qin, 2017), while others point to residues in the C-terminus

(Vlachova et al., 2003; Joseph et al., 2013). Still other groups have demonstrated that the pore domain is critical for temperature-dependent gating (Grandl et al., 2010; Zhang et al., 2018). The only aspect of TRPV temperature sensitive gating that is clear is that this feature is intrinsic to the channel itself (Cao et al., 2013).

Unlike their vertebrate homologs, invertebrate TRPV channels do not appear to have any role in temperature sensitivity; these channels appear to be exclusively activated by osmotic or mechanical stress

(Colbert et al., 1997; Tobin et al., 2002; Gong et al., 2004; Zhang et al., 2013). Additionally, invertebrate TRPV subunits appear to be interdependent, which is to say that functional TRPV channels require the combinatorial expression of at least two different types of TRPV subunits (Tobin et al., 2002; Gong et al., 2004). Invertebrate

TRPV channels appear to be sensitive to noxious chemicals such as benzaldehyde and certain insecticides

(Colbert et al., 1997; Nesterov et al., 2015), the latter of which appears to activate TRPV-expressing neurons in vivo.

14 Though there are differences in the specific sensory functions and in the assembly of vertebrate and

invertebrate TRPV channels, both types of TRPV channels generally activate at noxious thresholds for their

respective stimuli. In this regard, a common characteristic of both vertebrate and invertebrate TRPV channels is

that both types of channels are required for nociception (Caterina et al., 2000; Montell, 2003; Tobin and

Bargmann, 2004). Investigating invertebrate TRPV channels would therefore help clarify the mechanisms by

which nociceptive signaling occur. Invertebrate TRPV channels, due to their lack of temperature sensitivity, can

also serve as a valuable outgroup in order to study the evolution of temperature sensation. Despite identification

of possible channel agonists, as well as known mechanosensory and osmosensory roles, invertebrate TRPV

channels have never been previously shown to generate currents in a heterologous expression system. The lack of

a heterologously expressed invertebrate TRPV current limits the utility of these channels since the role of specific

residues and domains with regards to channel gating cannot be addressed as easily.

In Chapter 4 of this Thesis, I describe a project where, with the help of a serendipitous discovery by my

colleague Awani Upadhyay, invertebrate TRPV currents were heterologously expressed for the first time.

Specifically, invertebrate TRPV channels were shown to be activated by the Vitamin B3 metabolite nicotinamide.

It was also shown through TIRF microscopy that invertebrate TRPV channels only traffic to the membrane when two different subunits are coexpressed and appear to take a predominantly 2:2 stoichiometry. These results are significant because this was the first time invertebrate TRPV channels were expressed in a heterologous system.

CNBD-containing family channels

The Cyclic Nucleotide Binding Domain (CNBD) -containing channels are characterized by a C-terminal cyclic nucleotide binding domain. Evolutionarily, these channels were likely present in early prokaryotes. The evidence for this comes from channels identified in spirochetes that are gated by cyclic nucleotide binding, that share significant homology to the eukaryotic CNBD-containing channels, particularly in the eponymous CNBD

(Brams et al., 2014; Kesters et al., 2015; James et al., 2017). Among eukaryotes, there are several groups of

15 CNBD-containing channels, that have many diverse physiological roles. The voltage-gated members of eukaryotic CNBD-containing channels, which are the channels that I focused on in this Thesis, fall into three major groups: the Ether-a-go-go (EAG) family of channels, the Hyperpolarization-activated Cyclic Nucleotide gated (HCN) family of channels, and the Plant CNBD-containing channels. The Cylic Nucleotide Gated (CNG) channels, which are found in both plants and animals, display some levels of voltage sensitivity but require cyclic nucleotide binding in order to gate (Goulding et al., 1992; Leng et al., 1999; Li et al., 2017). Among bilaterians,

CNG channels are important for the transduction of olfactory and visual sensory information (Craven and

Zagotta, 2006).

The general model by which membrane potential influences the gating of these channels is similar to that found in Shaker channels (see Figure 1-4). However, the mechanism of VSD-PD coupling is known to be distinct from Shaker (Whicher and MacKinnon, 2016). Whereas the S4-S5 linker appears critical for gating in Shaker family, the linker in CNBD-containing channels does not appear to be as prominent (Whicher and MacKinnon,

2016). Indeed, coexpression of the isolated VSD and isolated PD of CNBD-containing channels in EAG family generates functional channels with near identical properties compared to the intact subunit (Lorinczi et al., 2015;

Tomczak et al., 2017). This is not to say that the S4-S5 linker does not play a role in gating, because the linker is still important for CNBD-containing channels to gate appropriately (Chen et al., 2001; Nieves-Cordones and

Gaillard, 2014; Tomczak et al., 2017). However, the mechanism by which S4-S5 linker appears to be distinct between Shaker and CNBD-containing channels. Interactions between the CNBD of a subunit and the

PD/VSD/C-linker of an adjacent subunit function alongside the S4-S5 linker in order to couple voltage sensor and pore in CNBD-containing channels (Figure 1-7). Another difference between CNBD-containing channels and

Shaker family channels is that tetrameric assembly of CNBD-containing channels occurs primarily through C- terminal domains (Daram et al., 1997; Ludwig et al., 1997; Jenke et al., 2003; Dreyer et al., 2004).

16

Figure 1-7. CNBD-containing channels share similar tertiary structure to Shaker family but are gated distinctly. A) An extracellular view of a CNBD-containing channel. The shortened linker between the VSD (blue) and PD (black) demonstrates a non-domain swapped quarternary architecture. B) The longer S4-S5 linker of Shaker family channels enables a domain swapped quarternary structure where the VSD (purple) of one subunit has the PD (gold) of the adjacent subunit in between itself and the central permeation pathway. C) Schematic of CNBD- containing channel; the front and back subunits are removed for display. CNBD-containing channels, as given by their name, are characterized a prominent C-terminal cyclic nucleotide binding domain (CNBD, shown in yellow). The CNBD is connected to the PD (black) with a helical domain termed the C-linker (shown in red). The CNBD interacts with the PD, C-linker and VSD (blue) of the adjacent subunit to facilitate voltage-dependent gating.

Activity of CNBD-containing channels can be influenced by many different biologically relevant molecules and stimuli. The effect of these various other molecules and stimuli on CNBD-containing channels appear to be heterogenous and require interaction with different portions of the subunit based on the type of

CNBD-containing channels one is interested in (Figure 1-8). As will be detailed within the subsections of each

type of CNBD-containing channel below, these interactions include: divalent cations, pH, PIP2, reactive oxygen species, and of course cyclic nucleotides. This diverse regulation of CNBD-containing channels allows these channels to be utilized in unique ways by eukaryotes.

17

Figure 1-8. Summary of modulators of voltage-gated CNBD-containing channels. Various small molecules and physiological stimuli interact with (A) EAG (B) HCN and (C) Plant CNBD-containing channels through the sites shown in the voltage sensor (blue), pore (black), C-linker (red) or CNBD (yellow). Certain modulators appear to have an effect on the various CNBD-containing channels but are not currently mapped to any known sites; an example of this is the interaction between PIP2 and HCN channels.

Despite these differences in regulation, it should be emphasized that a number of positions within the transmembrane core are universally or near universally conserved (Figure 1-9). For some of these highly conserved residues, there is functional conservation alongside the structural conservation. An example of this would be the string of basic residues present in S4, that serve to detect changes in membrane potential. In other cases, conservation of primary structure does not yield the same functional outcome across the various channel families. For example: the highly conserved G[F/Y]G motif of the selectivity filter within the pore loop allows

EAG channels to be strongly potassium selective, but does not confer significant potassium selectivity to HCN channels (Whicher and MacKinnon, 2016; Lee and MacKinnon, 2017). Some of these highly conserved residues do not have any known function, despite intense scrutiny.

Figure 1-9. Schematic of the conserved residues within the transmembrane core of CNBD-containing ion channels. Conserved residues are shown as red circles. The highly conserved HxxxC motif in the S5 domain is demarcated by the filled in circles.

18 In Chapter 5 of this Thesis, I will discuss a project that began with a search for the function of a near universally conserved HxxxC motif in the S5 transmembrane domain of CNBD-containing channels. It was hypothesized that this motif confers heme sensitivity to all CNBD-containing channels. The results suggest that heme does indeed interact with CNBD-containing channels in a manner demonstrating evolutionary conservation.

While there is some evidence that this heme effect occurs through the HxxxC motif, it remains to be seen whether or not this interaction is conserved across all lineages. I have focused my investigations of CNBD channels to just the eukaryotic voltage-gated channels in this Thesis and will therefore limit the discussion of CNBD-containing channel physiology to just the three groups voltage-gated CNBD-containing channels mentioned above.

EAG family channels

The EAG family of channels consists of three subfamilies of voltage-gated potassium selective channels.

These subfamilies are: the Ether-a-go-go (Eag) subfamily, the Eag-related gene (Erg) subfamily, and the Eag-like

(Elk) subfamily. Physiologically, EAG channels play many different roles that are dependent on the subfamily.

Channels from the Eag subfamily are thought to play an important role in regulating subthreshold excitability among neurons (Saganich et al., 1999; Saganich et al., 2001). Mutations in human Eag1 are known to cause both

Zimmerman-Laband Syndrome and Temple-Baraitser Syndrome. These are neurodevelopmental disorders, in which patients display overt craniofacial and skeletal defects in addition to intellectual disability and epilepsy

(Kortum et al., 2015; Simons et al., 2015). Interestingly, mice lacking Eag1 channels develop normally, and do not display any severe behavioral phenotype (Ufartes et al., 2013). Elk channels are also predominantly expressed in nervous tissue, and while there no currently known Elk mutations that lead to disease, Elk2 knockout mice display epilepsy (Trudeau et al., 1999; Zou et al., 2003; Zhang et al., 2010). Erg channels meanwhile, specifically human Erg1 channels, are celebrated for their role in cardiac repolarization (Sanguinetti et al., 1995; Trudeau et al., 1995). Indeed, mutations in human Erg1 channels are known to cause Long QT Syndrome Type 2, which can lead to sudden cardiac death (Curran et al., 1995). EAG family channels have recently been a focus as a

19 prominent cancer target, as some estimates suggest >80% of non-CNS tumors overexpress members of this family

and targeting specific EAG channels pharmacologically inhibits cancer proliferation (Hemmerlein et al., 2006;

Huang et al., 2015).

EAG family channels are known not to be sensitive to cyclic nucleotides; the CNBD of EAG channels is

therefore better characterized as the cyclic nucleotide binding homology domain (CNBHD). While the CNBHD

does not interact with cyclic nucleotides, the canonical cyclic nucleotide binding pocket within the CNBHD is

occupied by a portion of the N-terminus, termed the “intrinsic ligand” (Haitin et al., 2013; Whicher and

MacKinnon, 2016; Wang and MacKinnon, 2017). The bulk of the N-terminus bears homology to the so-called

Per-Arnt-Sim (PAS) domain; in the context of EAG family channels, the PAS domain is referred to as the eag domain. The interaction between the eag domain and the CNBHD is known to be important for channel gating

(Zhao et al., 2017; Dai et al., 2018). Indeed many disease-causing mutations in EAG channels have been mapped to the interface between these two domains (Haitin et al., 2013).

Despite lacking a response to cyclic nucleotides, EAG channels display exquisite sensitivity to a number of other biologically active small molecules. This includes sensitivity to divalent cations, such as calcium, magnesium and zinc; specifically, divalent cations prevent outward movement of the voltage sensor domain by binding to specific residues in S2 and S3 (Silverman et al., 2000). Interestingly, the divalent cation selectively is known to vary based on the subfamily within EAG, owing to differences in S4 (Zhang et al., 2009). EAG channels also appear to have sensitivity to the lipid signaling molecule PIP2. Though PIP2 is not explicitly

necessary for voltage sensor-pore coupling, as it is with the more distantly related KCNQ channel (Zaydman et

al., 2013), PIP2 can modulate gating in significant ways. PIP2 has been shown to inhibit channel opening in human

Eag1 channels as well as human Elk1 channels (Li et al., 2015a; Han et al., 2016). EAG family channels are also

known to be exquisitely sensitive to the pH of the extracellular environment; this pH sensitivity appears to require

the same residues necessary for divalent cation sensitivity (Silverman et al., 2000).

20 HCN family channels

HCN channels are hyperpolarization activated cyclic nucleotide gated channels that are non-specific cation permeable. Because these channels allow permeation of sodium and calcium in addition to potassium, channel opening typically results in depolarization of the membrane. Depolarization of the membrane following channel opening will then lead to subsequent closure of these channels typically alongside with action potential generation. Once the membrane repolarizes, HCN channels will begin to open once again and the cycle begins anew (Santoro and Tibbs, 1999). This quirky property of HCN channels, wherein opening of the channels leads to its own closure, led to the current generated by these channels being known as the “funny current” or If and is critical for pacemaking type neurons and cells throughout the body, including in the heart (Santoro et al., 1997;

Ludwig et al., 1998; Santoro et al., 2000).

Even though HCN channels open upon hyperpolarization of the membrane, the direction of movement of the Voltage Sensor Domain in response to changes in membrane potential is still conserved with other CNBD channels (Mannikko et al., 2002). This property of the VSD is important to keep in mind because it suggests that the coupling mechanism between the VSD and PD is somehow reversed in HCN channels. Recently published structures of the human HCN1 channel will be helpful in determining how this coupling is reversed, however one limitation is that these structures only depict the voltage sensor in the outward position (Lee and MacKinnon,

2017).

In contrast to EAG channels, HCN channels do interact with cyclic nucleotides, specifically cyclic AMP.

The interaction between HCN channels and cyclic AMP weakens the coupling of voltage sensor and pore domains, allowing for channel opening at more depolarized potentials (Wainger et al., 2001; Lee and MacKinnon,

2017). This allows for complex physiological responses, such as adrenergic regulation of cardiac pacemaking.

However, it should be noted that the sensitivity of HCN channels to cAMP is highly heterogenous; HCN4 channels show more greatly facilitated opening by cAMP binding than HCN1 channels (Ludwig et al., 1998;

Santoro et al., 1998; Santoro and Tibbs, 1999). HCN channels are also known to be regulated by intracellular magnesium (Lyashchenko and Tibbs, 2008; Vemana et al., 2008), but the effects of extracellular divalent cations

21 on HCN channels have not been as closely investigated. Like EAG family channels, PIP2 has been shown to affect

HCN channel gating as well (Zolles et al., 2006). While PIP2 has been shown to inhibit EAG channels, it appears

to facilitate HCN channel opening (Pian et al., 2006; Flynn and Zagotta, 2011; Baker et al., 2015). This would

suggest that PIP2 regulates HCN channels in a manner similar to EAG channels given the reversed VSD-PD

coupling, however it is not clear which residues are important for the HCN-PIP2 interaction (Baker et al., 2015).

Plant CNBD-containing channels

Like the metazoan EAG channels, Plant CNBD-containing channels are voltage-gated, potassium selective, and do not display significant sensitivity to cyclic nucleotides. Regulation of potassium levels by

Plant CNBD-containing channels is critical for plant growth, as well as responses to environmental stress

(Anschutz et al., 2014). There are generally two types of Plant CNBD-containing channels: the outward rectifiers, and the inward rectifiers. The outward rectifiers, such as GORK and SKOR, are gated by depolarization

(Gaymard et al., 1998; Ache et al., 2000). However, much like HCN channels, the inward rectifiers, such as KAT and AKT, are gated by hyperpolarization (Schachtman et al., 1992; Sentenac et al., 1992). Though plants do not contain excitable cells such as neurons or muscle cells, Plant CNBD-containing channels play a key role in the transport of water, ions and other key nutrients necessary for metabolism. GORK and the inward rectifiers in the

KAT and AKT subfamilies are specifically important in guard cells (Pilot et al., 2001; Szyroki et al., 2001; Hosy et al., 2003). Guard cells are cells that surround organelles called stoma that open and close in response to various environmental factors to facilitate transpiration. SKOR meanwhile, regulates the flow of potassium into the

Xylem (Gaymard et al., 1998).

Among the Plant CNBD-containing channel family, only KAT1 has been shown to be modulated by intracellular levels of cyclic nucleotides; specifically, this inwardly rectifying channel appears to be inhibited by cGMP (Hoshi, 1995). It is not clear from the data, whether this interaction requires the CNBD, and because only

KAT1 has been shown to be affected, cGMP binding may not be a common property among Plant CNBD-

22 containing channels. PIP2 appears to be necessary for KAT1 and SKOR gating (Liu et al., 2005). The data suggest that PIP2 prevents loss of current when channels are separated from the intracellular environment, but the precise

nature of the PIP2-channel interaction is still unclear. It is possible that PIP2 has an identical role among Plant

CNBD-containing channels that is shown in HCN and EAG channels, though more studies need to be done in order to discern the nature of the channel-PIP2 interaction. Divalent cations have been shown to influence the gating of KST1, an inward rectifier channel found in potato guard cells that is related to KAT1, however this interaction appears to require distinct portions of the voltage sensor as compared to EAG channels (Hoth and

Hedrich, 1999). Divalent cations do not appear to influence the gating of other Plant CNBD-containing channels.

SKOR has been shown to be sensitive to external pH (Lacombe et al., 2000), but it is unclear if this is conferred to the channel through the same mechanism as in EAG channels. SKOR and GORK are both modulated by reactive oxygen species or ROS (Demidchik et al., 2010; Garcia-Mata et al., 2010). ROS, particularly in the form of hydrogen peroxide, is known to play many diverse roles in plant metabolism (Apel and Hirt, 2004).

Preview of Thesis

Chapter 2 of this Thesis pertains to the techniques utilized in Chapters 3, 4 and 5; this Chapter gives some background information on how the techniques work and why these techniques were chosen over alternatives that can be used to study ion channels. In Chapter 3 of this Thesis, I will discuss a project that aims to determine mechanisms that regulate stoichiometry of a regulatory subunit associated with the Kv2 subfamily. The results are then used to develop an evolutionary model, which can potentially be used to predict the stoichiometry of other related regulatory subunits. In Chapter 4, I will present part of a project that identified a novel endogenous agonist of invertebrate TRPV channels, which is the vitamin B3 metabolite nicotinamide. I will discuss what inferences can be made due to the identification of the nicotinamide-channel relationship and speculate on the physiological significance of channel-nicotinamide relationship. In Chapter 5, I will discuss an unpublished project involving the CNBD channel family. In this project, heme was found to significantly alter CNBD channel gating; I aimed to

23 further characterize the heme-channel interaction. Finally, in Chapter 6, I will present a summary of the findings

from Chapters 3, 4 and 5. I will then discuss the impact of these findings on the field-at-large and will speculate on future directions based on these results.

24 References

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32 Chapter 2

Experimental techniques for probing ion channel structure-function relationships

In this chapter, I will discuss in detail the techniques used in Chapters 3, 4 and 5. In particular, I will give a brief overview of: the use of Xenopus laevis oocytes for heterologous expression of wild-type and mutant/chimeric channels, Two-Electrode Voltage Clamp (TEVC) and Total Internal Reflectance Fluorescence

(TIRF) Microscopy. I will give a background on these techniques, discuss their limitations, and then explain why these techniques are appropriate to answer the questions I am asking.

Xenopus laevis oocytes as a model for studying ion channels

Xenopus oocytes have been used to investigate membrane bound ion channels and neurotransmitter

receptors for over 30 years (Gundersen et al., 1983). There are several characteristics which make these oocytes

so amenable to channel research. Mature oocytes have a characteristic large round shape; they are approximately

one millimiter in diameter on average. This large size enables easy manipulation and a large surface area upon

which millions of channels can be embedded. Furthermore, they have a distinct darkly pigmented animal pole,

and lighter colored vegetal pole. The pigmented animal pole has a low level of autofluorescence; this low

background fluorescence is extremely useful for the types of sensitive fluorescence measurements as those

described in the TIRF section below. While the size, shape and color all help to make the Xenopus oocyte easy to

manipulate and use, this model system is particularly useful for studying ion channels due to its capacity to

synthesize large quantities of protein in short time as well as a relatively simple culturing system.

The general procedure for heterologously expressing ion channels in Xenopus oocytes is to inject

complementary RNA (cRNA) that encodes the channel gene or genes of interest (Soreq and Seidman, 1992).

Translation of injected cRNA can occur quite rapidly; while Xenopus oocytes possess maternally transferred

mRNAs, these mRNAs are translationally restricted through various mechanisms involving the polyadenylated

tails (Sheets et al., 2017). Exogenously introduced cRNA via microinjection can therefore produce large

33 quantities of protein without having to compete with endogenous transcripts for the access to the ribosomal

machinery (Gurdon et al., 1971). Injected oocytes are typically allowed to incubate in culture solution for several

hours or days depending on how the oocytes will be used (Figure 2-1). In general, incubating oocytes for longer

periods of time will allow for more channel expression; this property of Xenopus oocytes is beneficial in many

cases but can be undesirable depending on the goal of the experiment. An example of a situation in which a large

number of channels can be undesirable is the use of Xenopus in TIRF microscopy, as will be discussed below.

Figure 2-1. Injection of Xenopus oocyte with ion channel encoding cRNA. Altering the type of RNA and the duration of the incubation allows for flexibility in the types of experiments which can be performed using the Xenopus oocyte. Incubating for longer periods of time allows for more channel expression, which enables robust macroscopic currents that can be measured through TEVC. Alternatively, eggs should be incubated for shorter periods of time in order to allow resolution of individual fluorescently labelled ion channels in TIRF.

Other heterologous expression systems can also be used to study ion channel physiology, including

mammalian cell culture lines such as Human Embryonic Kidney (HEK) cells (Thomas and Smart, 2005).

Although cell culture systems have their advantages relative to Xenopus oocytes, namely faster clamping due to their smaller size, Xenopus electrophysiology was preferable in generating this Thesis due to a considerable ease of manipulation. Another key aspect of Xenopus oocytes that make them ideal for the experiments described in this Thesis is the fidelity with which translation of the microinjected RNA occurs. Protein synthesis shares a strong linear relationship with the amount of injected cRNA over a wide range of cRNA dilutions (Gurdon et al.,

1971; Buller and White, 1988). This property is useful for the RNA-titration type experiments as described in

34 Chapter 3, wherein mixtures of RNAs at several different ratios were used to determine the functional stoichiometry of heteromeric Kv2.1/Kv6.4 channels.

Preparation of Xenopus oocytes for cRNA injections

In order to utilize the Xenopus oocyte model system, oocytes must first be processed appropriately.

Ovaries from adult female specimens can be purchased through licensed vendors such as Xenopus 1. Ovaries are initially rinsed in calcium-free media, which is composed of: 98 mM NaCl, 2 mM KCl, 1 mM MgCl2, buffered with 5 mM HEPES and set to a pH of 7.2 using NaOH. Ovaries are then mechanically separated using sterilized forceps and placed in a collagenase digest, which consists of calcium-free media supplemented with 0.3-0.5 mg/ml Type II Collaenase (Sigma Aldrich) for 1 to 2 hours. Digested, defolliculated oocytes are then rinsed with calcium free media and incubated in culture media. Culture media consists of the following: 98 mM NaCl, 2 mM

KCl, 1.8 mM CaCl2, 1 mM MgCl2, 5 mM HEPES, 2.5 mM Na-pyruvate, 100U/ml penicillin, 100 μg/ml streptomycin, pH 7.2. Old and infected oocytes, characterized by depigmentation of the animal pole, loss of membrane integrity, and various “pock” marks, should be removed from the batch, as these can adversely affect the health of all other oocytes.

When picking oocytes for cRNA injection, it is important to select the large stage V/VI Xenopus laevis oocytes. Stage V/VI oocytes are the most mature oocytes and are characterized by the clearly delineated dark- pigmented animal pole and lightly pigmented vegetal pole (Sheets et al., 2017). This is in contrast to the smaller less developed oocytes that do not have clearly distinguishable animal and vegetal poles; these lower stage oocytes would not be able to adequately express heterologously introduced cRNA. Prior to injection, an injection dish should be created with a grid etched in the bottom to prevent the oocytes from moving as the injection needle is inserted. Glass Injection needles, pulled using the P-1000 Flaming/Brown Micropippette Puller (Sutter

Instruments, CA) or similar pipette-puller, must be filled with mineral oil and loaded onto the injector (NanoJect

II, Drummond Scientific). The tip of the injection needle must then be carefully broken to a size that can easily

35 facilitate cRNA flow but is not large enough to damage oocytes. Once oocytes are injected, they should ideally be incubated at 18°C for the desired length of time to obtain sufficient channel expression.

Two-Electrode Voltage Clamp (TEVC)

There are many types of electrophysiological techniques that can be tailored to the specific gating stimulus of an ion channel (e.g. voltage-gated, mechanogated etc.), but all of these techniques rely on some form of membrane “clamping” (Hille, 2001). That is to say, a cell membrane, which has some ion channel population, has a particular gating stimulus held (or “clamped”) at a constant value by the experimenter. The reason why the membrane needs to be clamped in these experiments is because the movement of the ions through the channels can affect the gating of the channels. Clamping the membrane is then a way to mitigate the influence of ionic movement on the channel state. The classic electrophysiologic “clamp” technique is the Two Electrode Voltage

Clamp (TEVC). Fundamentally, TEVC works by using one electrode to measure the voltage of the cell, and a second electrode to inject current to maintain the membrane voltage at a predetermined level (Guan et al., 2013).

TEVC is often used in conjunction with sophisticated bath perfusion systems to enable great control over the contents of the extracellular environment. This allows for the study of ion channel sensitivity to various types of biologic and synthetic molecules (Figure 2-2). When combined with standard molecular cloning and mutagenesis techniques to manipulate the injected channel encoding cRNA, TEVC can be used to interrogate which portions and residues of voltage-gated and voltage-sensitive ion channels are important for channel responses to specific environmental stimuli.

36

Figure 2-2. Schematic of Two-Electrode Voltage Clamp (TEVC) using a Xenopus oocyte. As the name suggests, TEVC makes use of two sharp electrodes that are impaled through the oocyte membrane. One electrode measures the membrane potential of the oocyte; this information is then relayed back to the computer, which compares the actual membrane potential to the desired Command potential as set by the experimenter. Current is then injected by the clamp to keep the membrane potential at the Command potential. These experiments are done in bath solutions (depicted by the +, - and o symbols) as determined by the type of experiment one wishes to perform. Sophisticated perfusion systems allow for rapid switching of this bath to allow for testing of various compounds on channel function.

One drawback to TEVC is that it can only resolve large currents that have a relatively slow time course.

That is to say, rapidly inactivating or small transient currents typically would not be detectable. Cut-Open

Vaseline Gap (COVG also known as Cut-Open) Voltage Clamp is better suited for these types of currents (Stefani and Bezanilla, 1998). An additional drawback to TEVC is that one can only infer the presence of multiple closed states of channels through subtle differences in channel opening kinetics, which may or may not be resolvable.

Patch clamp would be more appropriate to more definitively demonstrate the presence and estimate the stability of these closed-states (Sakmann and Neher, 1984; Zei and Aldrich, 1998). TEVC is still the appropriate technique to use for the experiments presented in this Thesis due to the fact that most of the channels expressed produce relatively large and robust currents. Moreover, the experiments performed require rapid switching of the bath solution in order to test the effects of various compounds on the ion channels. This is more difficult to accomplish in patch clamp and is virtually impossible in Cut-Open due to geometric constraints. Finally, most of the models tested do not necessarily require the ability to distinguish between different types of closed states at this time.

37 How to perform Two-Electrode Voltage Clamp

Acquisition of TEVC data depends on three key components that must be set prior to experimentation:

recording solution, glass electrodes, and clamp circuitry. The first step in obtaining TEVC data is making the

appropriate recording solutions for the desired experiments. Recording solution can be made by mixing

appropriate amounts of 1 M salt and buffering solutions. The pH should then be adjusted with careful titration

using either sodium hydroxide or methanesulfonic acid depending on the starting pH and the desired pH. Once the

desired pH is reached, sufficient UltraPure water must be added to adjust the solution volume to the correct

amount. In this Thesis, most solutions were composed of 96 mM NaOH, 2 mM NaCl, and 2 mM KCl; 5 mM

HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) is used as the buffering agent. This base solution was adjusted to the appropriate pH using methanesulfonic acid. Additional salt solutions, such as CaCl2 or MgCl2,

were added to the base solution prior to recording as the volume used of these salts would not significantly

influence the concentrations of the other ions.

Glass electrodes can be pulled using a micropipette puller in a manner similar to the injection needles

described above, taking care to use glass of the correct diameter. These glass electrodes, provided they contain a

filament, can be back-filled with 3M KCl solution to prevent the entry of bubbles into the tip of the electrode;

once sufficient KCl solution has entered the tip of the electrode, the electrode must be filled roughly halfway with

3M KCl and loaded onto the electrode holders. Electrodes should ideally be between 0.5 and 2 MΩ.

In order to correctly setup the clamp circuitry, the first step is to create a salt bridge between the perfusion

chamber and the bath headstage. This is necessary to prevent leaching of chemicals from the bath electrodes that

can damage oocyte membranes during recording while maintaining strong electrical contact with the membrane.

Once the salt bridge is setup and the bath electrodes are in place, the clamp circuitry can be switched on and the

data acquisition software (e.g. Clampex) should be opened and switched to the appropriate protocol.

Once the TEVC rig is correctly set up, adequate flow of the recording solution into the chamber must be

ensured prior to introduction of an injected oocyte. Flow of recording solution that is too fast can prevent proper

impalement of the oocyte by the electrodes due to motion. Recording solution that is flowing too slowly on the

38 other hand reduces the speed at which solutions can be switched and After the oocyte has been placed into the

recording chamber, the manipulators holding the electrodes should be moved into position in order to impale the

oocyte membrane. Once the oocyte is impaled the clamp can be switched on and the recording can begin. It is

imperative that the clamp is not running when the electrodes are in the solution but not in the oocyte as this can

create a dangerous short-circuit that can harm the experimenter and ruin the clamp.

Analysis of TEVC Data

An outline of the two main ways to analyze TEVC data is shown in Figure 2-3. The simplest way to

analyze TEVC data is to simply measure the amount of current generated at a specific voltage. By measuring the

currents at several voltages, one can generate what is known as a Current-Voltage plot, or I-V plot. The I-V plot is

useful in determining basic characteristics of ion channels, such as the membrane voltage at which the channels

typically open. However, changes in the I-V characteristic can also signify some type of change occurring in the

gating of the channels being studied, as will be shown in Chapter 5 when I discuss the study of the Plant K+

channel family. While I-V plots are useful, they are not always appropriate for in-depth analyses of voltage

sensor-pore coupling. For this, electrophysiologists can exploit the rapid clamping ability of TEVC to measure what are known as tail currents. The idea behind a tail current is that as the membrane is rapidly hyperpolarized following a depolarizing pulse, there is a lag period before channels that opened during the depolarizing pulse can fully close. By measuring the amount of current remaining in this tail, one can get an idea of the proportion of channels that are opened by the depolarizing pulse. There is necessarily a saturation point in the tail current, past which more current cannot be obtained. This makes intuitive sense as the number of channels embedded within the membrane is finite. The maximum size of the tail current is simply termed Gmax; one can then normalize the

tail currents at each voltage pulse preceding the one that gives Gmax in order to generate a G-V curve. Implicit in

the measurement of these so called “isochronal” tail currents is the assumption that there are two states which the

channels can occupy—a closed state and an open state—and that movement of the channel between these two

39 states is purely dependent on the membrane voltage. These assumptions allow for fitting of G-V curves with the

Boltzmann function, which can give an estimate of both the midpoint of activation (V50) and the coupling efficiency between voltage sensor and pore (z). Changes in these parameters can then be used to determine the significance of specific experimental conditions on channel function.

Figure 2-3. Example analysis of Two Electrode Voltage Clamp Data. A) The topmost traces show the current responses in an egg that is expressing channels made from wild-type human Kv12.1 RNA. The middle traces show the voltage protocol that is applied. Boxed regions indicate where measurement of current, conductance and voltage typically occur for generation of the current-voltage (I-V) and conductance-voltage (G-V) graph displayed in (B) and (C) respectively. B) Example I-V relationship of a depolarization-gated ion channel with outward rectification properties. As the membrane is depolarized, channels will begin to open and allow for net outward ionic flow. Ionic flow can increase exponentially as membrane potential increases. C) Example G-V relationship of a depolarization-gated outward rectifying channel. While the I-V relationship can theoretically increase to infinity, G-V relationships are restricted by the number of channels that can be embedded onto the membrane. G- V plots can be used to estimate the proportion of channels opening at a given membrane potential relative to the maximum number of channels that can be open.

40 Total Internal Reflectance Fluorescence (TIRF) Microscopy

Total Internal Reflectance Fluorescence (TIRF) microscopy is a sophisticated technique that allows for determination of ion channel stoichiometry, among many other uses (Ulbrich and Isacoff, 2007; Axelrod, 2008;

Nakajo et al., 2010). This form of microscopy makes use of laser light that is reflected against a thin coverslip upon which an adherent cell that is heterologously expressing a fluorescently labelled protein. The reflection of the laser beam generates an evanescent wave of light that can penetrate only a few hundred nanometers past the coverslip barrier. This evanescent wave illuminates only the cell membrane in contact with the coverslip, which greatly reduces background fluorescence from a cell’s cytoplasm and allows resolution of individual protein complexes at the cell membrane when used in combination with a high magnification objective (Axelrod, 2008).

Use of the laser over a period of minutes leads to photobleaching of the fluorescent labels; the number of these photobleaching steps can then be counted for a number of spots. If one assumes photobleaching is a stochastic process with a binomial distribution, the distribution of photobleaching steps can determine the number of subunits per protein complex (Ulbrich and Isacoff, 2007). One could use Förster Resonance Energy Transfer

(FRET)-based techniques to determine ion channel stoichiometry (Zheng et al., 2002; Kerschensteiner et al.,

2005). However, because TIRF allows for single molecule resolution, it can identify rare heteromeric channel populations which are likely not detectable by FRET. Indeed, in Chapter 3 of this Thesis, TIRF was used to identify the presence of a minor 2:2 Kv2.1:Kv6.4 heteromer. At the level of channel expression required to form this stoichiometry, it likely would not have been detected by FRET.

Performing TIRF Microscopy Experiments

A general schematic of how to carry out TIRF experiments is shown in Figure 2-4. The first step in performing TIRF experiments is the removal of the vitelline envelope surrounding Xenopus oocytes. The presence of the vitelline envelope can interfere with the resolution of channel complexes at the membrane. The envelope is mechanically removed through the use of sharp forceps. Incubating the oocyte in a hyperosmotic culture solution

41 (ND96 + 200 mM sucrose) for several minutes can help separate the vitelline envelopes from the oocyte membrane, facilitating its removal. Once the vitelline envelope is removed, the oocyte must be quickly placed onto an imaging chamber with a coverslip that has been preloaded with culture solution and some type of fluorescent molecules such as TetraSpeck beads (ThermoFisher) to help visualize the plane of the membrane.

Care must be taken to ensure the oocyte is placed with the animal pole down as the vegetal pole can be a source of significant autofluorescence. However, once a devitellinized oocyte has been loaded onto the coverslip, it should not be disturbed as further manipulation can rupture the oocyte and release immature channels onto the coverslip.

Once the oocyte has been placed in the imaging chamber, the imaging chamber must be loaded onto an inverted microscope that has been fitted with a high numerical aperture, high magnification objective. The objective must then be focused onto the membrane to visualize the diffraction-limited spots which represent fluorescently-tagged ion channels. The angle of the laser used for illumination can also be manipulated to accommodate the geometry of the oocyte. Once diffraction-limited fluorescent spots are identified, movies can be recorded on a camera at a minimum rate of 5 frames per second. Recording of the fluorescent signal for longer periods of time ensure that all originally identified fluorescent spots completely lose fluorescence; 4 minute movies were used in Chapters 3 and 4, for a total of 1200 frames per movie.

Figure 2-4. Using Total Internal Reflectance Fluorescence Microscopy (TIRFM) to identify ion channel stoichiometry. Oocytes from Xenopus laevis are prepared similar to TEVC recordings, with a few major differences. In order to make subunit counting feasible, cRNA that encodes for GFP tagged subunits must be injected, as untagged subunits will not be detected. Oocytes are only incubated for 12-18 hours to prevent

42 overexpression; too many channels in the membrane prevent precise measurement of isolated channel spots. The vitelline envelope of the oocyte must also be mechanically removed prior to imaging, as it is too thick for the shallow laser light—which is necessary for the phenomenon of total internal reflectance—to excite the fluorescent labels on the ion channels.

Analysis of TIRF data

The number of photobleaching steps per diffraction limited spot can be counted in a semi-automated

manner using free or inexpensive software and custom scripts. The specific algorithm and code used to count

photobleaching steps for the TIRF data in this Thesis is found in (Chen et al., 2014). To give a brief overview of

how this code was used, movie files are loaded onto the correct folder within the program, to show an overview of

all spots present in the movie (Figure 2-5). Individual spots are then selected with a 7 pixel diameter circle; larger

15 pixel diameter circles close to the selected spot being are used to generate the background fluorescence

waveform. The background fluorescence is then subtracted from photobleaching spot. Photobleaching spots can

be accepted into the dataset if they demonstrate clear, step-wise bleaching without subsequent increases in

fluorescence intensity (Figure 2-5).

Figure 2-5. Example photobleaching data acquired by Total Internal Reflectance Fluorescence Microscopy. This data was acquired from a Xenopus oocyte injected with GFP-Kv2.1 that was incubated for 12 hours. The trace on the right represents a filtered (0.75 Hz low-pass Bessel) version of the background subtracted fluorescence intensity of the spot circled in green on the left. The four steps (arrowheads) represent the photobleaching of four independent GFP tags within the diffraction-limited spot. Because there can only be one GFP tag per subunit, this would imply there are four GFP tagged subunits in this particular spot. The probability of detection of an individual fluorescent tag is dependent on the efficiency of protein folding; in Chapters 3 and 4, the single GFP

43 photobleaching detection efficiency is estimated at 0.69. Photobleaching of all fluorescent tags within a protein complex can be thought of as independent and identically distributed and would follow a simple binomial distribution.

44 References

Axelrod, D. 2008. Chapter 7: Total internal reflection fluorescence microscopy. Methods Cell Biol. 89:169-221. Buller, A.L., and M.M. White. 1988. Control of Torpedo acetylcholine receptor biosynthesis in Xenopus oocytes. Proc Natl Acad Sci U S A. 85:8717-8721. Chen, Y., N.C. Deffenbaugh, C.T. Anderson, and W.O. Hancock. 2014. Molecular counting by photobleaching in protein complexes with many subunits: best practices and application to the cellulose synthesis complex. Mol Biol Cell. 25:3630-3642. Guan, B., X. Chen, and H. Zhang. 2013. Two-electrode voltage clamp. Methods Mol Biol. 998:79-89. Gundersen, C.B., R. Miledi, and I. Parker. 1983. Voltage-operated channels induced by foreign messenger RNA in Xenopus oocytes. Proc R Soc Lond B Biol Sci. 220:131-140. Gurdon, J.B., C.D. Lane, H.R. Woodland, and G. Marbaix. 1971. Use of frog eggs and oocytes for the study of messenger RNA and its translation in living cells. Nature. 233:177-182. Hille, B. 2001. Ion channels of excitable membranes. 3rd.ed. Sinauer, Sunderland, Mass. xviii, 814 p. pp. Kerschensteiner, D., F. Soto, and M. Stocker. 2005. Fluorescence measurements reveal stoichiometry of K+ channels formed by modulatory and delayed rectifier alpha-subunits. Proc Natl Acad Sci U S A. 102:6160-6165. Nakajo, K., M.H. Ulbrich, Y. Kubo, and E.Y. Isacoff. 2010. Stoichiometry of the KCNQ1 - KCNE1 ion channel complex. Proc Natl Acad Sci U S A. 107:18862-18867. Sakmann, B., and E. Neher. 1984. Patch clamp techniques for studying ionic channels in excitable membranes. Annu Rev Physiol. 46:455-472. Sheets, M.D., C.A. Fox, M.E. Dowdle, S.I. Blaser, A. Chung, and S. Park. 2017. Controlling the Messenger: Regulated Translation of Maternal mRNAs in Xenopus laevis Development. Adv Exp Med Biol. 953:49- 82. Soreq, H., and S. Seidman. 1992. Xenopus oocyte microinjection: from gene to protein. Methods Enzymol. 207:225-265. Stefani, E., and F. Bezanilla. 1998. Cut-open oocyte voltage-clamp technique. Methods Enzymol. 293:300-318. Thomas, P., and T.G. Smart. 2005. HEK293 cell line: a vehicle for the expression of recombinant proteins. J Pharmacol Toxicol Methods. 51:187-200. Ulbrich, M.H., and E.Y. Isacoff. 2007. Subunit counting in membrane-bound proteins. Nat Methods. 4:319-321. Zei, P.C., and R.W. Aldrich. 1998. Voltage-dependent gating of single wild-type and S4 mutant KAT1 inward rectifier potassium channels. J Gen Physiol. 112:679-713. Zheng, J., M.C. Trudeau, and W.N. Zagotta. 2002. Rod cyclic nucleotide-gated channels have a stoichiometry of three CNGA1 subunits and one CNGB1 subunit. Neuron. 36:891-896.

45

Chapter 3

S6 gate divergence in regulatory subunits restricts the stoichiometry of heteromeric Shaker family channels

In this chapter, I will discuss a project that has been submitted to the Journal of General Physiology in

2018 that is aimed at identifying mechanisms by which obligatory heteromeric Shaker family potassium channels assume their stoichiometry. These regulatory subunits are known to predominantly contribute only one subunit in a functional tetramer. This poses a dilemma as the only portion of the subunit that is currently known to have a role in channel formation, the T1 tetramerization domain, is not sufficient to exclude a second silent subunit from being incorporated into a channel. The central question we wanted to answer in this project is: what other portions of the obligatory heteromeric subunit Kv6.4 are required to limit the stoichiometry of the Kv2.1-Kv6.4 heteromer to only 3:1? Using a combination of molecular mutagenesis, electrophysiology and Total Internal Reflectance

Fluorescence (TIRF) microscopy, we determined that the distal S6 region, which makes up the activation gate, is critical in regulating this stoichiometry. I performed the majority of the cloning and electrophysiology experiments with guidance from my advisor Tim Jegla. Xiaofan Li performed patch clamp experiments shown in

Figure 3-8E-F; these experiments help demonstrate the loss of current in my current titration experiment (Figure

3-8A-C) is not due to changes in single channel conductance. Jose Chu-Luo cloned the Kv6.4-PIPIIV-Kv2.1CT chimera, and Greg Busey cloned the Kv2.1 I383C, Kv6.4 M422C and Kv6.4-PIPIIV-Kv2.1 CT M422C mutants. I cloned the other mutants and chimeras. I acquired the TIRF data (Figures 3-4 and 3-5) alongside Keith

Mickolajczyk and Will Horton. I analyzed the TIRF data myself using Matlab code generated by Keith

Mickolajczyk. Damian van Rossum, Andriy Anishkin and Sree Chintapalli performed molecular dynamics simulations (Figure 3-10) that identified a potential mechanism by which the S6 gate region could serve to restrict the stoichiometry. My advisor, Tim Jegla, generated the evolutionary model in Figure 3-11.

46 Abstract

The Shaker family of voltage-gated K+ channels comprises four functionally independent gene subfamilies, Shaker (Kv1), Shab (Kv2), Shaw (Kv3) and Shal (Kv4) that regulate distinct aspects of neuronal excitability. Subfamily-specific assembly of tetrameric channels mediated by the N-terminal

T1 domain segregates Kv1-4, allowing multiple channel types to function independently in the same cell. Typical Shaker family subunits can form functional channels as homotetramers, but a group of mammalian Kv2 subfamily genes encode subunits that have a “silent” or “regulatory” phenotype characterized by T1-self incompatibility. These channels are unable to form homotetramers, but instead heteromerize with Kv2.1 or Kv2.2 to diversify the functional properties of these delayed rectifiers.

While T1 self-incompatibility predicts that these heterotetramers could contain 1 or 2 regulatory subunits, experiments show predominance of a 3:1R stoichiometry in which heteromeric channels contain a single regulatory subunit. Substitution of the self-compatible Kv2.1 T1 into the regulatory subunit Kv6.4 does not alter the stoichiometry of Kv2.1:Kv6.4 heteromers. To identify other channel structures that might be responsible for favoring the 3:1R stoichiometry, we compared the sequences of these mammalian regulatory subunits to independently-evolved regulatory subunits from cnidarians..

The most universal sequence feature of regulatory subunits are atypical substitutions in the highly- conserved consensus sequence of the intracellular S6 activation gate of the pore. We show here that two amino acid substitutions in the S6 gate of the regulatory subunit Kv6.4 restrict the functional stoichiometry of Kv2.1:Kv6.4 to 3:1R by limiting the formation and function of 2:2R heteromers. We propose a two-step model for the evolution of the asymmetric 3:1R stoichiometry that begins with evolution of T1 self-incompatibility to establish the regulatory phenotype, followed by drift of the activation gate consensus sequence under relaxed selection to limit stoichiometry to 3:1R.

47 Introduction

Shaker family voltage-gated K+ channels regulate neuronal excitability including many aspects

of action potential repolarization and timing. The Shaker family consists of four functionally

independent gene subfamilies which provide a diverse array of depolarization-gated K+ currents: Shaker

(Kv1), Shab (Kv2), Shaw (Kv3) and Shal (Kv4) (Wei et al., 1990; Covarrubias et al., 1991). Many of the delayed rectifier and transient A-type currents observed in neurons are encoded by Shaker family genes, and some of their most notable roles are described below. Kv1 channels localize to the axon initial segment and juxtaparanodes of mammalian neurons where they participate in axonal action potential repolarization (Wang et al., 1993; Dodson et al., 2002; Ogawa et al., 2008; Trimmer, 2015).

They appear to underlie the classical delayed rectifier of the squid giant axon (Rosenthal et al., 1996).

Kv2 channels encode the majority of somatodendritic delayed rectifiers (Tsunoda and Salkoff, 1995b;

Murakoshi and Trimmer, 1999; Du et al., 2000; Malin and Nerbonne, 2002; Misonou et al., 2005), but

they can also be found in ankyrin-free zones of the axon initial segment in mammalian neurons (King et

al., 2014). Mammalian Kv3 channels underlie rapid high threshold delayed rectifiers that facilitate high

spike rates in fast-firing neurons (Wang et al., 1998; Lau et al., 2000; Rudy and McBain, 2001; Lien and

Jonas, 2003). Kv4 channels, in contrast, encode classical somatodendritic A-currents found in many

mammalian and invertebrate neurons (Tsunoda and Salkoff, 1995a; Malin and Nerbonne, 2000, 2001;

Carrasquillo and Nerbonne, 2014), although it should be noted that Kv1 subfamily channels can

contribute a kinetically distinct component to somatodendritic A-type currents in at least some

mammalian neurons (Malin and Nerbonne, 2001; Carrasquillo and Nerbonne, 2014).

Shaker family channels are tetrameric (MacKinnon, 1991; Long et al., 2005a) with each subunit

containing a canonical voltage-gated cation channel core motif of six transmembrane domains (S1-S6).

S1-S4 comprise the voltage sensor domain (VSD), while S5-S6 comprise the pore domain (PDs) with

48 the K+ selectivity filter formed on the extracellular side by a highly-conserved loop (Jiang et al., 2003;

Long et al., 2005a, b). Each channel has four spatially-independent VSDs, but a single pore formed by

extensive intersubunit contact between the PDs. The unique and defining feature of Shaker family

channels relative to other voltage-gated K+ channels is the presence of a cytoplasmic N-terminal domain,

T1, which promotes assembly of tetramers and forms another large intersubunit interface (Shen and

Pfaffinger, 1995; Xu et al., 1995; Kreusch et al., 1998; Long et al., 2005a). Figure 3-1A-D summarize

the structural layout of tetrameric Shaker channels, including the two major intersubunit interfaces in T1

and the inner pore. T1-mediated tetramer assembly requires physical interaction between neighboring

T1 domains and is subfamily-specific because T1s from distinct subfamilies are not compatible and do

not interact (Shen and Pfaffinger, 1995; Xu et al., 1995). The T1 domain therefore plays a key role in maintaining functional segregation of the Kv1, Kv2, Kv3 and Kv4 subfamilies.

Figure 3-1. Tetrameric Shaker family K+ channels have two major intersubunit interfaces. A) Schematic cartoon depicting subunit domain arrangement in Shaker family channels. Two diagonally-opposed subunits of the tetrameric channel are shown. The pore domains (PD, red) from each subunit surround a central ion-conducting pore, while the voltage sensor domains (VSD, blue) are physically isolated at the periphery. The conserved N-terminal cytoplasmic assembly domains (T1, black) form a ring beneath the pore. B) A more detailed cartoon of the tetrameric ion conducting pore from an extracellular perspective, with adjacent subunits differentially shaded. Transmembrane helices S5 and S6 are depicted with cylinders and a selectivity filter helix is shown as a black ribbon. The pore-lining S6 helix forms a major intersubunit interface (arrows) and the intracellular side comprises the activation gate. C) Cartoon of the cytoplasmic T1 ring with adjacent subunits differentially shaded. There are major intersubunit contacts between neighboring, but not diagonally-opposed, T1s (arrows). Helices are depicted with

49 cylinders and β-sheets are depicted with rectangles. D) Amino acid alignment of mouse Kv2.1 and Kv6.4 with residue numbers given at the right margin. Identical residues are shaded black and conservative substitutions are shaded gray. The T1 domain is underlined in red, transmembrane domains S1-S6 are underlined in dark blue and the pore domain is underlined in light blue. The alignment was produced with the CLUSTALW algorithm as implemented in MEGA7 (Kumar et al., 2016) and unconserved N- and C-termini have been trimmed. E) T1 self-incompatibility in regulatory subunits (blue) theoretically allows formation of heterotetramers with a single (3:1R) or two diagonally-opposed (2:2R) regulatory subunits because 4 compatible T1 contacts (+ signs) remain in each case. However, T1 self-incompatibility rules out formation of 2:2R with adjacent regulatory subunits, 1:3R and 4R tetramers, all of which have at least one incompatible intersubunit contact (minus signs).

Kv1-4 subunits can typically assemble as functional homotetramers, and the isolated T1 domain is itself able to self-assemble into a tetrameric structure (Pfaffinger and DeRubeis, 1995; Kreusch et al.,

1998). A notable exception to this rule of assembly is a group of ten “silent” or “regulatory” subunits discovered in mammals, Kv5.1, Kv6.1-6.4, Kv8.1-8.2 and Kv9.1-9.3 that are self-incompatible and thus not able to form homotetramers, but can form functional heteromeric channels with novel biophysical properties when co-expressed with Kv2.1 or Kv2.2 (Post et al., 1996; Patel et al., 1997; Salinas et al.,

1997a; Salinas et al., 1997b; Kramer et al., 1998; Zhu et al., 1999; Ottschytsch et al., 2002). While the names of these channels imply that they belong to distinct gene subfamilies, phylogenetic analysis places them within the Kv2 subfamily (Jegla et al., 2012; Li et al., 2015b), thus preserving the principle of subfamily-specific assembly. In the absence of Kv2.1, Kv6.4 regulatory subunits do not assemble into tetramers and do not traffic to the plasma membrane (Ottschytsch et al., 2002; Ottschytsch et al., 2005).

Mammalian regulatory subunit T1s also do not oligomerize in vitro, suggesting T1 self-incompatibility plays a significant role in generation of the self-incompatible regulatory subunit phenotype (Salinas et al., 1997b; Kramer et al., 1998; Ottschytsch et al., 2002).

Crystal structures of Shaker channels show that a T1 domain only physically interacts with T1s from neighboring subunits; there is no interaction between T1s of diagonally-opposed subunits (Kreusch et al., 1998; Long et al., 2005a). T1 self-incompatibility of regulatory subunits therefore predicts that

50 regulatory subunits could combine with Kv2.1 in two possible channel stoichiometries: 1) an

asymmetric stoichiometry with three Kv2.1s and a single regulatory subunit (3:1R), or 2) a symmetric

stoichiometry with two Kv2.1s and two diagonally-opposed regulatory subunits (2:2R) (Figure 3-1D).

T1 self-incompatibility rules out 1:3R channels and 2:2R channels with adjacent regulatory subunits, but in theory would not be able to distinguish between a 3:1R or diagonal 2:2R stoichiometry. However,

FRET-based determination of the Kv2.1:Kv9.3 stoichiometry suggests that heteromers form predominantly in the asymmetric 3:1R stoichiometry (Kerschensteiner et al., 2005). Measurements of

gating charge components attributable to Kv6.4 and Kv2.1 supports a 3:1R stoichiometry for

Kv2.1:Kv6.4 heteromers (Bocksteins et al., 2017), and therefore suggests that the 3:1R stoichiometry

might be shared among mammalian Kv2 family regulatory subunits.

We reasoned there must be additional factors beyond T1 self-incompatibility of regulatory

subunits that bias formation of functional channels in the 3:1R stoichiometry. There is indeed evidence

suggesting that T1 self-incompatibility is not the only factor guiding assembly of regulatory subunits.

For instance, substitution of the Kv2.1 T1 into Kv6.4 is not sufficient to restore assembly of functional

homomeric channels (Ottschytsch et al., 2005), and N-terminus/C-terminus interaction may play a role

in blocking the ability of Kv6.4 to assemble with Kv3 channels (Bocksteins et al., 2014). To find

channel regions that might be responsible for determining regulatory subunit stoichiometry, we used an

evolutionary approach to look for regulatory subunit sequence signatures near the two major subunit

interfaces in Shaker family tetramers, the T1 contacts and the PD contacts. T1-containing Shaker

channels can be traced to a common ancestor of all extant metazoans (Li et al., 2015b), and

diversification of voltage-gated K+ channels (including the Shaker, KCNQ and EAG families) into the

functional classes, or subfamilies, found in vertebrates occurred prior to the divergence of bilaterians

and cnidarians (Jegla and Salkoff, 1994; Jegla et al., 1995; Jegla and Salkoff, 1997; Sand et al., 2011;

51 Jegla et al., 2012; Martinson et al., 2014; Li et al., 2015b; Li et al., 2015c). Kv1-Kv4 channels can all be found in cnidarians and the functional properties of all four subfamilies, including subfamily-specific assembly, are highly-conserved between cnidarians and bilaterians. Most importantly for this study, the regulatory subunit phenotype evolved independently in cnidarians in the Kv1, Kv3 and Kv4 subfamilies

(Jegla and Salkoff, 1997; Jegla et al., 2012; Li et al., 2015b), providing a rich data set for identification of regulatory subunit sequence signatures.

We show here that alteration of the highly conserved sequence of the pore’s intracellular activation gate is a universal signature for evolution of the regulatory subunit phenotype. The activation gate forms part of the interface between subunits (Figure 3-1B) and extensive mutagenesis studies of

Shaker subfamily channels have demonstrated that various substitutions in the gate can block channel maturation, interfere with gating and alter conductance (Hackos et al., 2002; Kitaguchi et al., 2004; Pau et al., 2017). We tested the hypothesis that these gate substitutions could play a role in determining the stoichiometry of heteromeric channels containing regulatory subunits using a combination of Total

Internal Reflection Fluorescence (TIRF) microscopy and electrophysiology.

Methods

Molecular Cloning

Mouse Kv2.1 and Kv6.4 cDNA were isolated from whole brain mRNA using RT-PCR, cloned into pOX plasmid (Jegla and Salkoff, 1997) using HindIII/XbaI (Kv2.1) or EcoR1/Xba1 (Kv6.4), and sequence confirmed to code proteins identical to Refseq (Kv2.1, NP_032446; Kv6.4, NP_080010).

Kv6.4-Kv2.1T1, Kv6.4-PIPIIV-Kv2.1CT and Kv6.4-PIPIIV chimeras were generated from two PCR fragments using standard overlap PCR techniques and cloned into pOX using EcoR1/Xba1. For the

52 Kv6.4-Kv2.1T1 chimera, the following primers were used: piece 1 (Kv2.1 T1), 5’-

TATAGAATTCCACCATGCCGGCGGGCAT-3’ (sense) and 5’-

AGCCTCCTCCCGGCGCAGCGTCTCAGCC-3’ (antisense), and piece 2 (Kv6.4 core to C-terminus),

5’-AGGCTGAGACGCTGCGCCGGGAGGAGG-3’ (sense) and 5’-

TCTCTCTAGAGTGGTGGGAGTTACAT-3’ (antisense). Kv6.4-PIPIIV-Kv2.1CT was generated with the following primers: piece 1 (Kv6.4 upstream of gate), 5’-TCTCGAATTCCACCATGCCCATGTCT-

3’ (sense) and 5’-GTTATTGACGATAATTGGAATTGGGA-3’ (antisense), and piece 2 (Kv2.1 gate through C-terminus), 5’-CTTATCATGGCTTTCCCAATTCCAATT-3’ (sense) and 5’-

TCTCTCTAGATTCAGATACTCTGATCC-3’ (antisense). Kv6.4-PIPIIV was generated in the same manner except using a piece 2 containing the Kv6.4 C-terminus generated with the following primers:

5’-TCATGCCTTTCCCAATTCCAATTATCG-3’ (sense) and 5’-

TCTCTCTAGAGTGGTGGGAGTTACAT-3’ (antinsense). Kv6.4-Kv2.1CT, Kv6.4- PITIIV-Kv2.1CT,

Kv6.4-PIPIIF-Kv2.1CT and Kv6.4-PITIIF-Kv2.1CT were generated by cloning PCR fragments into

Kv6.4-PIPIIV-Kv2.1CT using an internal BamH1 site in S6 upstream of the gate and Xba1. Fragments were generated using the antisense primer 5’-TCTCTCTAGATTCAGATACTCTGATCC-3’ and the following sense primers: 5’-

AGCGGGATCCTTATCATGGCTTTCCCAGCCACATCCATCTTCCACACCTTCTCCGAGTTCTA

C-3’ (Kv6.4-Kv2.1CT);5’-

AGCGGGATCCTTATCATGGCTTTCCCAATTACAATTATCGTCAATAAC-3’ (Kv6.4-PITIIV-

Kv2.1CT); 5’-AGCGGGATCCTTATCATGGCTTTCCCAATTCCAATTATCTTCAATAACTTCTC-

3’ (Kv6.4-PIPIIF-Kv2.1CT); and 5’-

AGCGGGATCCTTATCATGGCTTTCCCAATTACAATTATCTTCAATAACTTCTC-3’ (Kv6.4-

PITIIF-Kv2.1CT). Pore cysteine mutants for Cd2+ block experiments were constructed using the

53 QuikChange protocol (Wang and Malcolm, 1999) with the following primers: Kv2.1 I383C, (sense) 5-

GTTGGTTACGGAGACTGCTACCCTAAGACTCTCCTG-3’ and (antisense) 5’-

CAGGAGAGTCTTAGGGTAGCAGTCTCCGTAACCAAC-3’; Kv6.4 M422C (sense)

GTGGGCTATGGGGACTGCGTCCCTCGCAGCGTCCCG and (antisense)

CGGGACGCTGCGAGGGACGCAGTCCCCATAGCCCAC. Enhanced green fluorescent protein

(Zhang et al., 1996) (called GFP here for simplicity) tags were cloned onto the N-terminus of Kv2.1 or

Kv6.4-base constructs using NheI/HindIII or NheI/EcoRI, respectively. The flexible linkers between the

GFP tag and channel N-termini were QQQAST for Kv2.1 and QQQNST for Kv6.4. Oligonucleotide

primers were obtained from IDT (Integrated DNA Technologies, IL) and all constructs were confirmed

by sequencing.

cRNA synthesis and Xenopus oocyte preparation

For all constructs, capped cRNAs were made by run-off transcription from NotI-linearized

templates using the T3 mMessage mMachine kit (Life Technologies). Prior to injection, cRNA was

purified using lithium chloride precipitation and analyzed by gel electrophoresis for integrity. cRNAs

were stored at -80°C and diluted to the desired concentration for injection in a 1:20 mix of the RNAse

inhibitor SUPERase-In (Invitrogen) and nuclease-free water. Mature stage V/VI Xenopus laevis oocytes

were injected with 50 nL of cRNA using a NanoJect II injector (Drummond Scientific). Defolliculated

oocytes were isolated from Xenopus laevis ovaries (Xenopus 1) using collagenase digestion (Type II

Collagenase; Sigma Aldrich), as previously described and maintained in an oocyte culture solution

consisting: 98 mM NaCl, 2 mM KCl, 1.8 mM CaCl2, 1 mM MgCl2, 5 mM HEPES, 2.5 mM Na-

pyruvate, 100U/ml penicillin, 100 μg/ml streptomycin, pH 7.2. Oocytes were incubated at 18°C for 12-

54 18 hours prior to TIRF microscopy and 24-72 hours prior to two-electrode voltage clamp and patch clamp.

Electrophysiology

Two-electrode voltage clamp recordings for biophysical characterization of heteromers and

expression ratio titrations use the following bath solution: 96 mM NaOH, 2 mM NaCl, 2 mM KCl, 1

2+ mM CaCl2, 1 mM MgCl2, 5mM HEPES, pH 7 using methanesulfonic acid. For Cd block experiments,

the following base solution was used: 100 mM LiCl, 2 mM CaCl2, 1 mM MgCl2, and 5mM HEPES, pH

7.4. All chemicals were obtained from Sigma Aldrich. Glass electrodes (1-3 MΩ) were pulled using a P-

1000 Flaming/Brown Micropippette Puller (Sutter Instruments, CA) and filled with 3 M KCl. Oocytes

were clamped using a Dagan CA-1B amplifier (Dagan, MN) and the pClamp 10 acquisition software for

data collection and analysis (Molecular Devices, CA). Data was sampled at 10 kHz and lowpass filtered

at 2 kHz using a 4-pole Bessel filter. Voltage-activation (GV) curves were determined from isochronal tail currents recorded after test pulses. Tail currents were fit with a single Boltzmann ( ( ) =

𝐺𝐺 𝑉𝑉 ( )/ +A2), where G is the conductance at voltage V, A1 is the initial value, A2 is the final value, 𝐴𝐴1−𝐴𝐴2 𝑉𝑉−𝑉𝑉50 𝑑𝑑𝑑𝑑 1+𝑒𝑒 V50 is the midpoint and dx is the slope factor. Reported V50 and slope factor values are the mean ±

S.E.M. of fits from individual oocytes, and data were normalized prior to averaging for display. Steady

State Inactivation (SSI) data were determined from peak currents measured during a test pulse to +40

mV following pre-pulses to varying voltages and fit with the same equation. Non-inactivating pedestal

current fractions were defined as A2/A1.

For on cell patch recordings of single channels, vitelline membranes of oocytes were

mechanically removed as previously described (Li et al., 2015a) and transferred to a recording dish

55 filled with internal solution (138 mM KMES, 4 mM KCl, 10 mM HEPES, 5 mM EGTA, pH 7.2 (MES, methanesulfonate)). Patch pipettes (0.4-1 MΩ) were coated with Sticky Wax (Kerr Dental Laboratory

Products, Orange, CA), fire polished and filled with a solution containing 140 mM KMES, 2 mM KCl, 1 mM MgCl2, 0.2 mM CaCl2, 10 mM HEPES, pH 7.2. Ground was isolated with a 1 M NaCl agarose bridge, junction potential was cancelled prior to patch formation, and pipette capacitance was compensated. Data were collected using a Multiclamp 700A amplifier and the pClamp 9 acquisition package (Molecular Devices, Sunnyvale, CA). Data were sampled at 20 kHz and filtered at 1.4 kHz.

TIRF microscopy

Prior to imaging, oocyte vitelline envelopes were mechanically removed as previously described

(Li et al., 2015a). Stripped oocytes were mounted on Number 0 Coverslips (VWR) in a custom-made chamber filled with oocyte culture solution. The chamber was loaded onto a Nikon TE-2000 inverted microscope outfitted with a 60x N.A. 1.45 objective (Nikon). An 80 mW argon ion laser (Spectra

Physics) was used for illumination, and a Cascade 512B EMCCD (Roper Scientific) was used for detection. 240 s movies were acquired at 5 frames per second using MetaVue software (Molecular

Devices). TIRF photobleaching data were analyzed in a semi-automated manner using a custom

MATLAB (Mathworks) script. Briefly, fluorescence intensity over time traces for well-isolated spots were acquired from TIFF stacks, which summed counts in a 7-pixel diameter circle drawn around candidate channels. Fluorescence changes in a larger 15-pixel diameter circle near the chosen spot were used to estimate the background fluorescence in order to correct baseline changes. The number of photobleaching steps was then determined using a published step-finding algorithm (Chen et al., 2014).

Only spots which were stationary and had stable baselines were included in the analysis. Analysis of all

56 observed step dwell times indicated the time constant for a GFP bleaching step to occur was ~68 s, so

>97% of all bleaching events should be captured in 240 s movies.

Sequence Alignments

Sequences were aligned for display using the CLUSTALW algorithm as implemented in

MEGA7 (Kumar et al., 2016). Sequence logos displayed in Figure 3-2 were generated from alignments

using WebLogo (Crooks et al., 2004). Supplemental File 1 contains sequences and accession numbers for all channels included in the sequence logos.

Molecular Modeling

Structural models were based on published open and closed state computational models of Kv1.2

(Pathak et al., 2007) and included the entire conserved T1-S6 regions of Kv2.1 and Kv6.4. Sequence

alignments of Kv2.1 and Kv6.4 to Kv1.2 for structural modeling were generated using MUSCLE, as

implemented by Jalview (Waterhouse et al., 2009); a FASTA file of aligned sequences is provided

(Supplemental File 2). Models were built from the alignments using MODELLER (9v8) (Sali and

Overington, 1994; Marti-Renom et al., 2000; Eswar et al., 2007). For each target (Kv2.1, Kv6.4 and

Kv6.4-PIPIIVN), we generated 30 initial models. MODELLER uses a variety of methods (e.g. Z-DOPE,

molpdf, GA341) to assess the models. These scores are associated with the estimated accuracy of the

model. For each target-based model set we selected the top candidate model guided by the molecular

PDF score (molpdf), which is the sum of all spatial restraints achieved when transferring structural

regions from the template to the target. These candidates also had the lowest Z-DOPE scores of the 30 initial models generated for each template. Assembly and visualization of all the tetrameric models was

57 done in VMD (Humphrey et al., 1996) using visual manipulations and custom-written Tcl scripts. We

generated four tetramer arrangements: Kv2.1 homomer, Kv2.1:Kv6.4 (3:1R), Kv2.1:Kv6.4 (2:2R

diagonal), and Kv2.1:Kv6.4-PIPIIVN (2:2R diagonal). For the first 3 models, local structural clashes in

the models were removed through 1000-step energy minimization using Conjugate Energy algorithm,

followed by relaxation using short (1 ps) Molecular Dynamics simulation in vacuo at 300K, and a 100-

step energy minimization. All the optimizations were done using NAMD (Phillips et al., 2005) with

CHARMM36 force field (MacKerell et al., 1998). For the Kv2.1:Kv6.4-PIPIIVN model, the gate of the final Kv2.1 homomer model was placed into the final Kv2.1:Kv6.4 2:2R model and the model was optimized with a 100-step energy minimization, 1 ps thermal relaxation, then another 100-step energy minimization again with symmetry restraints. The space accessible for the K+ ion center at the pore gate

constriction in the open state models was visualized using “solvent” representation for the probe radius

set to 1.8 Å (potassium ion in CHARMM36 force field (1.76375 Å) rounded to the nearest 0.1A) and all

the protein atoms having their VdW radius derived from CHARMM36.

Results

The highly-conserved S6 activation gate sequence degenerates in regulatory subunits

To determine whether there are sequence signatures that accompany the regulatory subunit

phenotype (i.e. inability to form functional homotetramers), we compared the amino acid sequences of

eight evolutionarily distinct clusters of regulatory subunits comprising 43 proteins from mouse and sea

anemone (Nematostella vectensis) that arose independently as determined by phylogenetic analyses

(Jegla et al., 2012; Li et al., 2015b). We restricted the analysis to these two species because the phenotype for each regulatory subunit cluster has been experimentally verified. In mammals, the 10-

58 gene Kv2 subfamily regulatory subunits can be split into two groups with separate evolutionary origins:

1) Kv5.1, which first appeared in chordates, and 2) Kv6-9, which later arose independently in

vertebrates (Li et al., 2015b). In Nematostella, six additional evolutionarily-independent clusters of regulatory subunits can be found in the Kv1 (4 clusters), Kv3 (1 cluster) and Kv4 (1 cluster) subfamilies

(Jegla and Salkoff, 1997; Jegla et al., 2012; Li et al., 2015b). We found no consistent regulatory-subunit

sequence alterations in the subfamily-specific T1 assembly domain, but 42/43 regulatory subunits

contained unusual substitutions in the C-terminal of S6 (PVPVIV in Drosophila Shaker), a region which

is typically among the most highly-conserved in Shaker family channels (Figure 3-2A-C). This sequence

includes the proline gating hinge and forms extensive intersubunit contacts in the structure of the

mammalian Shaker channel Kv1.2 (Long et al., 2005a). The bold underlined residues, V474 and V478 in Drosophila Shaker, face inward toward the conduction pathway, forming a hydrophobic seal in the closed state (Long et al., 2005a). Access to V474 in Shaker is gated by pore opening, while access to

V478 is possible in both open and closed channels, suggesting that V478 forms the cytoplasmic boundary of the intracellular activation gate (Liu et al., 1997; del Camino and Yellen, 2001; Hackos et al., 2002; Webster et al., 2004; del Camino et al., 2005). To facilitate comparison of gate sequences between channels, in this paper we will number residues by their position within the 6-amino acid

(PVPVIV) gate sequence; in this nomenclature V474 is V2 and V478 is V6. Each independent cluster of regulatory subunits has a distinct pattern of substitutions in this activation gate sequence (Figure 3-2D-

K), with several types of substitutions occurring multiple times. These include loss or displacement of

one of the gating hinge prolines, insertion of polar residues that disrupt gate hydrophobicity, or insertion

of large aromatics that could potentially cause steric conflicts. In the mammalian Kv6-9 cluster, P3 is

lost (9/9 proteins) and replaced with a polar residue (S/T, 8/9 proteins), and an aromatic is inserted in

place of V6 (8/9 proteins; N in Kv8.1) (Figure 3-2D). Site-directed mutagenesis of Shaker has found that

59 hydrophilic substitutions at P3 can destabilize the closed state (Sukhareva et al., 2003), and aromatic substitution of V6 (V6W) results in non-conducting channels (Hackos et al., 2002).

Figure 3-2. The S6 activation gate sequence differs from the Shaker family consensus in evolutionarily- independent regulatory subunit groups. A) Cartoon showing the PD of two diagonally-opposed subunits of a closed Shaker family channel tetramer with the location of a 6 amino acid sequence (PVPVIV) comprising the S6 gating hinge and activation gate overlaid to show its approximate location at the intracellular side of the conduction pathway. (B) S6 consensus sequence logo constructed from all 27 mouse and sea anemone (Nematostella vectensis) Shaker family subunits that can form functional homotetrameric channels. The hinge and activation gate are boxed with a red outline and positions 1-6 as used in the paper are indicated. A highly-conserved section of S6 upstream of the gate is also shown in the logo. Amino acid frequency is encoded in letter height, and colors are used to depict amino acid class (blue, hydrophobic; green, hydrophilic; red, proline; pink, aromatic; orange, acidic; purple, basic). Note the conservative substitution V2I and V4I are found in mouse Kv2.1 and Kv2.2 and are typical for the Kv2 subfamily. C) A similar sequence logo constructed from 43 mouse and sea anemone Shaker family regulatory subunits shows degeneration of the gate PVPVIV consensus sequence; only P1 is found in a majority of regulatory subunits. D) Sequence logo for 9 mouse Kv2 subfamily regulatory subunits with a common evolutionary origin (Kv6.1-6.4, Kv8.1-8.2, Kv9.1-9.3). Asterisks mark positions with unusual substitutions in at least some members of the group. Note the loss of P3 (hydroxyl typical) and V6 (aromatic typical) across the group. E-K) Sequence logos broken out for 7 additional evolutionarily-independent groups of mouse and sea anemone regulatory subunits. Each separate group of regulatory subunits has a distinct pattern of unusual gate substitutions, marked by asterisks. Accession numbers and sequences for channels used to make the sequence logos in (B-E) are given in Supplemental File 1, and amino acid frequencies at each gate position are given in Supplemental File 2.

Since mutations within the Shaker activation gate have been shown to disrupt channel function and expression (Hackos et al., 2002; Kitaguchi et al., 2004), we hypothesized that the unusual

60 substitutions observed in regulatory subunit gate sequences could play a role in restricting the functional

stoichiometry of regulatory subunit-containing heteromers. To test this hypothesis, we chose mouse

Kv2.1/Kv6.4 heteromers as a model system because analysis of gating current components supports a

3:1R stoichiometry (Bocksteins et al., 2017) and heteromeric channels can be functionally distinguished from Kv2.1 homomers based on gating properties (Figure 3-3, Table 1). In oocytes co-expressing Kv2.1 and Kv6.4, currents arising from Kv2.1:Kv6.4 heteromers can be separated from currents arising from

Kv2.1 homomers with a depolarizing prepulse (Figure 3-3C,D), allowing estimates of heteromer:homomer ratios at the level of functional channels.

Figure 3-3. Comparison of gating properties of Kv2.1 homomers and Kv2.1:Kv6.4 heteromers. A) Example currents recorded under two electrode voltage clamp (TEVC) from oocytes expressing Kv2.1 (left) or Kv2.1:Kv6.4 (right) in response to 400 ms depolarizations ranging from -60 mV to +40 mV in 20 mV increments from a holding potential of -100 mV. Tail currents were recorded at -50 mV. Scale bars are included for current amplitude and time and the voltage protocol is shown below the currents. The step to 0 mV is highlighted in red to emphasize the left-shift in voltage activation in Kv2.1:Kv6.4. For Kv2.1:6.4, a 1:10 cRNA ratio was used for the experiments in this Figure 3-to produce a predominantly heteromeric current (see Figure 3-8). B) Normalized conductance voltage (GV) relationships determined from isochronal tail currents recorded at -50 mV after 400 ms steps to the indicated voltages are shown for Kv2.1 (black, n = 7) and Kv2.1:Kv6.4 (red, n = 10). Data points show mean, error bars indicate standard error and smooth curves represent a single Boltzmann fit of the data. V50 and slope values are given in Table 1. The V50 values were significantly different (t-test, p < 0.001).

61 C) Example currents recorded from oocytes for Kv2.1 and Kv2.1:Kv6.4 during a protocol used to determine steady state inactivation (SSI). Oocytes were stepped from a holding potential of -100 mV to voltages ranging from -110 to +30 mV in 10 mV increments for 4 s prior to a 500 ms step to +40 ms to measure current size. For display purposes, here we show currents recorded in response to pre-pulses from -40 mV to +20 mV in 20 mV increments for Kv2.1, and from -100 mV to -40 mV in 20 mV increments for Kv2.1:Kv6.4. The -40 mV sweep is highlighted in red for comparison and represents a voltage at which steady state inactivation is virtually absent in Kv2.1 and near maximal in Kv2.1:Kv6.4. Thus a comparison of currents recorded at +40 mV after pre-pulses to -100 mV and – 40 mV can be used to distinguish homomeric and heteromeric current fractions. Scale bars are given for time and current amplitude and the voltage protocol is shown below the currents. D) Normalized SSI curves for Kv2.1 (black, n = 7) and Kv2.1:Kv6.4 (red, n = 10). Data show mean ± S.E.M., and curves show a single Boltzmann fit of the data. V50 and slope values are given in Table 1. Note steady state inactivation is incomplete for both Kv2.1 and Kv2.1:Kv6.4; residual pedestal current fractions as determined by Boltzmann fitting are given in Table 1.

TIRF microscopy reveals multiple Kv2.1:Kv6.4 heteromer stoichiometries

We used a Total Internal Reflection Fluorescence (TIRF) microscopy single-molecule bleaching

assay to determine the stoichiometry of Kv2.1:Kv6.4 heteromers. TIRF single molecule bleaching

assays have been successfully used to determine the subunit stoichiometry of many different types of ion

channels and other membrane-associated proteins (Ulbrich and Isacoff, 2007; Nakajo et al., 2010; Coste

et al., 2012; Upadhyay et al., 2016). We chose this approach because it allows identification and

quantification of minor heteromer species that would likely be missed by macroscopic analyses such as

intersubunit Förster resonance energy transfer (FRET) and gating currents that have previously been

used to look at Kv2.1:Kv9.3 and Kv2.1:Kv6.4 stoichiometry (Kerschensteiner et al., 2005; Bocksteins et

al., 2017). In this technique, individual membrane surface-expressed channels containing fluorescently- tagged subunits (visible as fluorescent dots) are bleached with a laser and the number of bleaching steps is counted to infer fluorescent subunit numbers (See Methods and (Ulbrich and Isacoff, 2007)).

When we expressed GFP-Kv2.1 by itself in Xenopus oocytes to exclusively form Kv2.1 homotetramers, or Kv2.1 with GFP-Kv6.4 to introduce heteromer formation, we observed fluorescent

62 spots that bleached in discrete steps (Figure 3-4A,B). For GFP-Kv2.1 homomers, fluorescent spots bleached in 1, 2, 3 and 4 steps (Figure 3-4A). From the distribution of channels bleaching in 1-4 steps,

we estimated the probability for detection of an individual GFP fluorophore to be ~69% (Figure 3-4C).

Co-expressing GFP-Kv2.1 with Kv6.4 significantly shifted the distribution towards fewer bleaching steps (p < 0.05, Fisher’s exact test), consistent with a reduction in the average number of Kv2.1 channels/subunit through heteromer formation (Figure 3-4D). Note that while the percentage of channels in this Kv2.1:Kv6.4 co-expression experiment bleaching in 4 steps is significantly reduced, their presence suggests some Kv2.1 homomers are still formed. Residual homomer formation makes it difficult to calculate the stoichiometry of the heteromers from this distribution alone. Therefore, we co- expressed Kv2.1 with GFP-Kv6.4 in a 1:50 ratio to maximize formation of heteromers and to selectively detect heteromers to calculate their stoichiometry. With the GFP tag on Kv6.4 instead of Kv2.1, channels spots predominantly bleached in 1 step (consistent with a 3:1R stoichiometry), but to our surprise, there were also a significant number of spots bleaching in 2 steps (Figure 3-4B,E). Using the

GFP bleaching probability determined from the GFP-Kv2.1 experiment (0.69), 2:2R heteromers were estimated to represent ~30% of all heteromers in this experiment (Figure 3-4F). The absence of spots bleaching in 3-4 steps is consistent with the idea that T1-incompatibility would block assembly of heteromeric channels with more than 2 Kv6.4 subunits. Co-expressing Kv2.1 with GFP-Kv6.4 in a 15:1 ratio decreases 2:2R significantly to 13% (p < 0.05, χ2 test), demonstrating that the proportion of 2:2R

heteromers formed depends on the Kv2.1:GFP-Kv6.4 ratio and is therefore not an artifact of the

bleaching step counting procedure. These results provide evidence that Kv2.1:Kv6.4 2:2R heteromers

can form, but also suggest that their formation is highly inefficient compared to 3:1R heteromers. If the

two stoichiometries formed with equal probability, we would expect 2:2R heteromers to predominate at

the 1:50 RNA ratio of Kv2.1:GFP-Kv6.4.

63

Figure 3-4. Determination of Kv2.1:Kv6.4 heteromer stoichiometry by TIRF photobleaching assay. A) Example TIRF images from an oocyte expressing GFP-Kv2.1 before (left) and after photobleaching (middle). Spots that bleached to background during the photobleaching period (blue circles) were analyzed for number of bleaching steps. Bleach-resistant spots (yellow circles) or spots that bleached but did not have a stable baseline (magenta) were not included in the analysis. Example fluorescence traces for spots in the movie bleaching in 1 (I), 2 (II), 3 (III) and 4 (IV) steps are shown at the right margin. Fluorescent spots (circled) bleachable in discrete steps were observed only in oocytes expressing GFP- tagged channels. B) Example TIRF images and fluorescence traces before and after photobleaching for Kv2.1 co-expressed with GFP-Kv6.4, labeled as in (A). Only spots with 1 or 2 bleaching steps were observed. C) Frequency distribution of bleaching steps for channel spots in oocytes expressing GFP- Kv2.1 (Kv2.1 homomers). Only spots bleaching in 1-4 steps were observed. Assuming all channels are tetramers with 4 GFPs, a binomial fit of the distribution (squares) estimates the probability of detecting GFP fluorescence and bleaching at 69% in our experimental setup. D) Frequency distribution of

64 bleaching steps for channel spots in oocytes co-expressing GFP-Kv2.1 and Kv6.4 compared to GFP- Kv2.1 alone. The addition of Kv6.4 RNA significantly reduces the frequency of 4 steps from 22% to 11% and reduces the frequency of 3 steps from 42% to 32%. There is a corresponding significant increase in the frequency of 2 steps from 25% to 39% and increases the frequency of 1 steps from 12% to 17% (p < 0.05, Fisher’s Exact Test). This shift in step distribution confirms detection of Kv2.1:Kv6.4 heteromer formation. E) Frequency distribution of bleaching steps for channel spots in oocytes injected with Kv2.1:GFP-Kv6.4 in an ~1:50 ratio to maximize heteromer formation. Only spots bleaching in 1-2 steps were observed, and 81% bleach in 1-step. F) Relative frequency of 3:1R and 2:2R Kv2.1:GFP- Kv6.4 heteromers calculated from the bleaching step distribution utilizing the 69% GFP detection probability determined for Kv2.1 homomers.

The S6 gate of Kv6.4 limits formation of 2:2R heteromers

We next examined the role of the Kv6.4 T1 domain and the Kv6.4 S6 activation gate in

restricting the formation of 2:2R heteromers. In keeping with our prediction that T1 self-incompatibility should not be sufficient to block formation of 2:2R heteromers (Figure 3-1), co-expressing Kv2.1 with a

Kv6.4 chimera containing the self-compatible T1 of Kv2.1 (GFP-Kv6.4-Kv2.1T1, Figure 3-5A) did not significantly increase the formation of 2:2R heteromers as measured by TIRF (Figure 3-5B). However, when Kv2.1 was coexpressed with a Kv6.4 chimera containing the Kv2.1 S6 activation gate (PIPIIV) and C-terminus (Kv6.4-PIPIIV-Kv2.1CT, Figure 3-5A), 2:2R heteromer formation was significantly increased to ~60% (Figure 3-5B, p < 0.01, χ2 test). 2:2R formation with the GFP-Kv6.4-PIPIIV-

Kv2.1CT chimera was concentration dependent and higher than 2:2R formation with WT Kv6.4 over a broad concentration range (Figure 3-5C). We included the Kv2.1 C-terminus in this original gate chimaera because the C-terminus of Kv2.1 has previously been implicated in channel assembly

(Mohapatra et al., 2008). A Kv6.4 chimera that retains the Kv6.4 activation gate (PATSIF) but includes the Kv2.1 C-terminus (GFP-Kv6.4-Kv2.1CT, Figure 3-5A) did not increase 2:2R formation (Figure 3-

5D), specifically implicating the activation gate as the key region for increasing 2:2R formation within the Kv6.4-PIPIIV-Kv2.1CT chimera. Furthermore, the chimera Kv6.4-PIPIIV, which contains only the

65 Kv2.1 activation gate, was also able to increase 2:2R formation (Figure 3-5D). Adding the Kv2.1 T1 to

the Kv6.4-PIPIIV-Kv2.1CT (Kv6.4-Kv2.1T1-PIPIIV-Kv2.1CT) did not further increase 2:2R formation

(Figure 3-5D) and did not form functional homotetramers as measured by TIRF (see Figure 3-5 legend for step counts) or outward current recorded at +40 mV (143 ± 14 nA, n = 3). This is consistent with previous observations that the substitution of the Kv6.4 T1 and S6 with Kv2.1 sequences is not sufficient to confer homotetramer formation to Kv6.4 (Ottschytsch et al., 2005). The Kv6.4-Kv2.1T1-

PIPIIV-Kv2.1CT chimera supports our hypothesis that the self-incompatible Kv6.4 T1 is not responsible for preferential assembly of Kv2.1:Kv6.4 3:1R heteromers over 2:2R heteromers, but also suggests that

T1 is not entirely responsible for Kv6.4 subunit self-incompatibility.

We next investigated the role of the two most unusual residues of the Kv6.4 activation gate (T3 and F6, Figure 3-2) in reducing 2:2R formation by introducing them back into the GFP-Kv6.4-PIPIIV-

Kv2.1CT chimera. We chose this chimera instead of GFP-Kv6.4-PIPIIV because it displayed the highest

2:2R formation. When T3 and F6 were re-introduced into Kv6.4-PIPIIV-Kv2.1CT together or individually (Figure 3-5A), the proportion of 2:2R heteromers was statistically identical to the 2:2R proportion we observed for WT Kv6.4 (Figure 3-5D). These results as a whole show that the unusual T3 and F6 substitutions in the Kv6.4 S6 activation gate, and not T1 incompatibility, play a significant role in suppressing Kv2.1:Kv6.4 2:2R heteromer formation.

66

Figure 3-5. The Kv6.4 activation gate but not T1 limits the formation of Kv2.1:Kv6.4 2:2R heteromers. A) Schematic diagram of the chimeric constructs used to test 2:2R formation by TIRF microscopy. The N-terminal T1 domain is labeled and transmembrane domains S1-S6 are shown as rectangles, with a star next to the S6 activation gate. Sequences derived from Kv6.4 are shown in black and sequences inserted from Kv2.1 are shown in blue. Chimera names are given below each diagram. B) Percentages of 3:1R and 2:2R heteromers calculated from TIRF photobleaching assays for Kv2.1 co-expressed with GFP- tagged versions of WT Kv6.4 (red), Kv6.4-Kv2.1T1 (black), and Kv6.4-PIPIIV-Kv2.1CT (blue). The stoichiometry distribution for Kv6.4-Kv2.1T1 was unchanged, but Kv6.4-PIPIIV-Kv2.1CT had significantly fewer 3:1R heteromers and significantly more 2:2R heteromers (p < 0.05, χ2 test). Kv2.1 was expressed with Kv6.4-Kv2.1T1 and Kv6.4-PIPIIV-Kv2.1CT in a 1:80 and a 1:60 cRNA ratio, respectively, to eliminate Kv2.1 homomers and bias 2:2R heteromer formation. C) Expression ratio dependence of 2:2R heteromer formation is shown for WT Kv6.4 and Kv6.4-PIPIIV-Kv2.1CT. 2:2R formation was significantly higher for both constructs at the highest concentration tested relative to lower concentrations (*p < 0.05, χ2 test). Note 2:2R formation is higher for Kv6.4-PIPIIV-Kv2.1CT across a broad range of expression ratios. D) Kv2.1:Kv6.4 2:2R heteromer percentages detected for all Kv6.4 chimeras tested in TIRF photobleaching assays at a highly Kv6.4-biased expression ratio. 2:2R

67 formation was significantly increased relative to WT for the Kv6.4-PIPIIV-Kv2.1CT, Kv6.4-PIPIIV and Kv6.4-Kv2.1T1-PIPIIV-Kv2.1CT constructs (*p < 0.05, χ2 test). The P3T and V6F mutations in the Kv6.4-PIPIIV-Kv2.1CT background eliminate the increase in 2:2R formation. Bleaching step counts and expression rations for the data in D were as follows (construct name, ratio, # 1-steps, # 2 steps): Kv6.4 WT, 1:200, 98, 23; Kv6.4-Kv2.1CT, 1:142, 87, 21; Kv6.4-PITIIF-Kv2.1CT, 1:154, 85, 13; Kv6.4-PITIIV-Kv2.1CT, 1:128, 64, 16; Kv6.4-PIPIIF-Kv2.1CT, 1:334, 67, 15; Kv6.4-PIPIIV-Kv2.1CT, 1:60, 72, 39; Kv6.4-PIPIIV, 1:185, 80, 39; Kv6.4-Kv2.1T1-PIPIIV-Kv2.1CT, 1:78, 63, 32). We did not detect spots bleaching in 3or 4 steps for any GFP-Kv6.4 chimera tested in this study.

The Kv6.4 gate blocks function of 2:2R heteromers

While our TIRF results suggest that 2:2R Kv2.1-Kv6.4 heteromers do form, and that the

efficiency of formation is increased by substitution of Kv2.1 gate residues into Kv6.4, they do not

answer the question of whether 2:2R heteromers incorporating either WT Kv6.4 or Kv6.4-PIPIIV-

Kv2.1CT are functional. To examine functionality, we first verified that currents generated by

Kv2.1:Kv6.4-PIPIIV-Kv2.1CT heteromers, like currents from Kv2.1:Kv6.4 WT heteromers, can be

distinguished from Kv2.1 homomer currents based on steady state inactivation (Figure 3-6, Table 1). In

oocytes expressing a 1:10 ratio of Kv2.1:Kv6.4-PIPIIV-Kv2.1CT, the V50 of steady state we measured

was -82.32 ± 1.43 mV, compared to -62.56 ± 0.58 mV for oocytes expression Kv2.1:Kv6.4 WT. Thus a

depolarizing prepulse can also be used to distinguish the current fraction contributed by Kv2.1:Kv6.4-

PIPIIV-Kv2.1CT heteromers from the Kv2.1 homomer current fraction. We then measured total current

size and calculated heteromer current size in oocytes injected with a constant amount of Kv2.1 cRNA

titrated against increasing amounts of either Kv6.4 WT or Kv6.4-PIPIIV-Kv2.1CT cRNA to gain insights into the functionality of 2:2R heteromers. Assuming that tetrameric channels form as a dimer of dimers, as has been previously suggested (Tu and Deutsch, 1999), the availability of Kv2.1 homodimers required for 3:1R formation should drop as the expression of Kv6.4 increases. This will limit the absolute number of 3:1R channels that can be formed and favor 2:2R formation (Figure 3-7A,B). If 3:1R

68 channels are the only functional species, then current size should drop towards 0 as the Kv6.4 expression

ratio is increased, eliminating 3:1R channels (Figure 3-7A). In contrast, if both the 3:1R and 2:2R

heteromers are functional, the current size should drop off much less with increasing Kv6.4 (Figure 3-

7B) due to current flowing through the 2:2R heteromers that preferentially form at high Kv6.4

expression ratios.

Figure 3-6. Biophysical properties of heteromeric currents from oocytes co-expressing Kv2.1:Kv6.4- PIPIIV-Kv2.1CT. A) Example currents recorded from an oocyte expressing Kv2.1:Kv6.4-PIPIIV- Kv2.1CT in 1:10 ratio in response to 1 s voltage steps ranging from -80 to +40 mV in 20 mV increments from a holding potential of -100 mV. The voltage protocol is shown below the currents, the 0 mV trace is highlighted in red, and scale bars are given for time and current amplitude. B) Normalized GV relationship for Kv2.1+Kv6.4-PIPIIV-Kv2.1CT determined from isochronal tail currents recorded at -50 mV after 1 s steps to the indicated voltages. Data points show mean ± S.E.M (n = 10), and the solid blue curve represents a single Boltzmann distribution fit of the data (V50 and slope are included in Table I). Dashed red and black curves show the Boltzmann fits for Kv2.1 homomers and Kv2.1:Kv6.4 WT, respectively. C) Example current traces for the SSI protocol for an oocyte expressing Kv2.1:Kv6.4- PIPIIV-Kv2.1CT in a 1:10 cRNA ratio. The voltage protocol is indicated below the current; the oocyte was held at -100 mV and current traces recorded in response to 4 s pre-pulses ranging from -120 mV to - 40 mV in 20 mV increments are shown, the -40 mV trace is highlighted in red. Pre-pulses were followed by a 500 ms step to +40 mV to show current availability. (D) Normalized SSI relationship determined for Kv2.1:Kv6.4-PIPIIV-Kv2.1CT from peak current amplitudes recorded at +40 mV following 4 s pre- pulses to the indicated voltages. Data show mean ± S.E.M. (n = 10), the blue curve represents a single Boltzmann fit (parameters in Table 1), and the dashed red and black curves show Boltzmann fits for Kv2.1 and Kv2.1:Kv6.4 WT, respectively.

69 Comparison of current sizes for titrations of WT Kv6.4 and Kv6.4-PIPIIV-Kv2.1CT cRNA

against a constant amount of Kv2.1 cRNA are shown in Figure 3-8A,B. During electrophysiological

recordings, oocytes were held at -100 mV for 500 ms, followed by a 4 second hyperpolarizing prepulse

to -120 mV to relieve closed state inactivation, and then pulsed to +40 mV for 500 ms to measure current size. With Kv6.4 WT, total current elicited by a +40 mV depolarization dropped rapidly as

Kv6.4 cRNA levels were increased and approached 0 at a Kv6.4:Kv2.1 100:1 ratio (Figure 3-8C).

Current levels also decreased with increasing Kv6.4-PIPIIV-Kv2.1CT titration but were significantly

higher across almost the entire titration, with 43 ± 3.7% of the control current level remaining at a 1:50

ratio of Kv2.1:Kv6.4-PIPIIV-Kv2.1CT, the highest expression level we tested (Figure 3-8C). cRNA ratios differed slightly between the two titrations, so we statistically compared means from the closest cRNA ratios (Figure 3-8C, brackets). The ~2-fold increase in 2:2R formation that we observed in our

TIRF experiments for Kv6.4-PIPIIV-Kv2.1CT compared to Kv6.4 WT is not sufficient to explain the large difference in current size between the titrations, but the difference could be explained if 2:2R heteromers significantly contribute to outward currents only in theKv2.1:Kv6.4-PIPIIV-Kv2.1CT

titration.

70

Figure 3-7. Predicted effect of Kv2.1:Kv6.4 expression ratio on the number of functional channels for models in which (A) only 3:1R heteromers conduct or (B) both 3:1R and 2:2R heteromers conduct. 3:1R heteromers require adjacent Kv2.1 subunits which will become increasingly rare as Kv6.4 expression levels are increased. Therefore, if only 3:1R heteromers conduct (A), then few functional channels will form at highly Kv6.4-biased expression ratios. Current size would thus be expected to approach zero for a constant amount of Kv2.1 titrated against increasing concentrations of Kv6.4. However, if both 3:1R and 2:2R heteromers conduct (B), then the number of functional channels at highly Kv6.4-biased expression ratios, where more 2:2R heteromers form, will remain significantly higher, and current size would not be predicted to approach zero.

We do not think that differences in single channel conductance Kv2.1:Kv6.4 WT and

Kv2.1:Kv6.4-PIPIIV-Kv2.1CT 3:1R heteromers alone could explain the differences in the titrations.

First, we compared single channel conductance for Kv2.1 homomers and Kv2.1:Kv6.4 heteromers

expressed in Xenopus oocytes using on-cell patches. We observed channels with a conductance of 8.3 ±

0.7 pS in oocytes expressing only Kv2.1 (Figure 3-8D,F). In oocytes expressing Kv2.1 and Kv6.4, we observed channels with conductance statistically identical to these Kv2.1 homomers and one additional group of channels with an ~1/3 smaller conductance which presumably represent Kv2.1:Kv6.4 heteromers (Figure 3-8E,F). This small single channel conductance drop is not sufficient to explain the near complete loss of current at high Kv6.4 cRNA concentrations. However, it could be explained if the channels we detected represent 3:1R heteromers and 2:2R heteromers are either non-conducting or have

71 a drastically reduced conductance. Second, if the difference we observed in whole cell current size in the

Kv2.1:Kv6.4 WT and titrations simply reflected a difference in single channel conductance between

3:1R heteromers incorporating Kv6.4 WT or Kv6.4-PIPIIV-Kv2.1CT, then we would expect that the fold difference in current size across the titration curves to remain constant. Instead, current size in the

Kv2.1:Kv6.4-PIPIIV-Kv2.1CT titration compared to the Kv2.1:Kv6.4 WT titration increased from

1.06 ± 0.22-fold (mean ± SD) at low expression ratios to 4.22 ± 0.60-fold (mean ± SD) at high ratios

(Figure 3-8G), consistent with concentration-dependent introduction of an additional conducting species in the Kv2.1:Kv6.4-PIPIIV-Kv2.1CT titration. We did not pursue single channel analysis for

Kv2.1:Kv6.4-PIPIIV-Kv2.1CT because we reasoned that the introduced gate changes might lessen differences in single channel conductance, making it difficult to detect multiple heteromer species and unsafe to assign observed conductances to specific stoichiometries. While significant current remains at high concentrations of Kv6.4-PIPIIV-Kv2.1CT cRNA, the current level does diminish >50%. This drop off could be due to remaining inefficiencies in 2:2R formation (we observed a maximum of ~60% 2:2R heteromers in TIRF) and/or a modest reduction in the single channel conductance of the 2:2R heteromers relative to 3:1R heteromers and Kv2.1 homomers.

72

Figure 3-8. Kv2.1:Kv6.4 expression ratio titrations predict that 2:2R heteromers have negligible conductance for Kv6.4 WT and conducting for Kv6.4PIPIIV-Kv2.1CT. A) Example currents from oocytes expressing Kv2.1 alone (left, black) or Kv2.1:Kv6.4 WT (red) at 9:1, 1:1 and 1:20 ratios. Kv2.1 cRNA level was kept constant, and currents were recorded at +40 mV following a 4 s prepulse to -120 mV to relieve SSI. Note the dramatic decrease in current size as Kv6.4 cRNA level is increased. Scale bars indicate current amplitude and time. B) Example currents from a similar titration of Kv2.1 vs. Kv6.4-PIPIIV-KV2.1CT; Kv2.1 control is shown in black and cRNA ratios shown are 3:1, 1:1 and 1:20. More current is readily apparent at the 1:20 ratio for Kv6.4-PIPIIV-KV2.1CT than Kv6.4 WT. C) Current amplitude vs. expression ratio titrations are shown for Kv2.1:Kv6.4 WT (red) and Kv2.1:Kv6.4- PIPIIV-KV2.1CT (blue). Expression ratio is given as -log(Kv2.1fraction); 1.0 = a 1:10 Kv2.1:Kv6.4 cRNA ratio. Kv2.1 cRNA amount remained constant while the Kv6.4 cRNA species amount was varied to achieve the given expression ratio. Data were normalized to the amplitude of control Kv2.1 homomeric currents. Data points show mean ± S.E.M. (n = 10-18 eggs per ratio), and asterisks indicate a significant difference (p < 0.05, t-test) between the indicated pairs of data points. D) Example single channel current traces from an on-cell patch from an oocyte expressing Kv2.1 alone. Traces were

73 recorded at -20 mV, 0 mV and +20 mV, the dashed blue line indicates the average closed baseline, and the dashed red lines indicate 1X and 2X the average open channel amplitude for the trace. E) Traces labeled as in (D) for an on-cell patch containing 2 channels from an oocyte co-expressing Kv2.1 with Kv6.4. In this example the dashed red lines indicate 1X and 2X the average open channel amplitude of the largest channel recorded in the trace. Red arrows point to examples of single or double channel openings with smaller than expected amplitudes, indicating a 2nd channel with a lower conductance. F) Plots of single channel current amplitude vs. voltage are given for Kv2.1 homomers expressed in isolation and for presumed homomers and heteromers from oocytes expressing Kv2.1 with Kv6.4. Single channel conductance values shown on the graph were calculated from linear fits; n was 6 for the two Kv2.1 homomer measurements and 5 for the Kv2.1:Kv6.4 heteromers. The Kv2.1:Kv6.4 heteromer conductance was significantly smaller (p < 0.05, t-test). Note there is a small offset in reversal potential between the experiments conducted on oocytes expressing Kv2.1 alone vs. Kv2.1 + Kv6.4 that alters single channel amplitude but does not interfere with conductance measurements; these experiments were conducted at different times with distinct solution batches. G) Normalized current size ratio for Kv2.1:Kv6.4-PIPIIV-Kv2.1CT vs. Kv2.1:Kv6.4 WT increases ~4-fold as the Kv6.4 species cRNA amount is increased. Data show mean ± S.D. and are derived from the data point pairs statistically compared in (C). Kv2.1:Kv6.4 ratios were similar but not identical between the titrations, and data is plotted using the average of the Kv2.1:Kv6.4 ratio of the data pair. Standard Deviation for the ratios was calculated by propagation of error from the original measurements in (C). E) Normalized total current vs. heteromeric current for the Kv2.1:Kv6.4 WT co-expression titration shown in (C). To determine the amplitude of the heteromeric current, we selectively inactivated heteromers with a 4 s prepulse to -40 mV prior to measuring current amplitude at +40 mV. The amplitude of the heteromeric current was determined by adjusting the current fraction removed by the -40 mV prepulse by the fractional inactivation determined from SSI analysis (Figure 3-3D, Table 1). In this case, the inactivating fraction was multiplied by 1.38 to account for the observation that only 27.8 % of heteromeric channels are not expected to inactivate during the prepulse. F) Normalized heteromeric vs. total current amplitude for Kv2.1:Kv6.4-PIPIIV-Kv2.1CT; heteromeric fraction was calculated as in (D) except using a multiplication factor of 1.44 to match the 30.4% pedestal observed for Kv2.1:Kv6.4-PIPIIV-Kv2.1CT in SSI analysis. Note in both titrations the current approaches 100% heteromer at a 1:10 Kv2.1:Kv6.4 species expression ratio (-logKv2.1 fraction = 1).

The titration results might also be explained if Kv6.4 WT was translated far more efficiently than

Kv6.4-PIPIIV-Kv2.1CT, offsetting the current size reduction in the Kv2.1:Kv6.4-PIPIIV-Kv2.1CT titration. However, we would expect such an offset would also offset the cRNA expression ratio required to achieve a given percentage of heteromeric current. We therefore estimated the contribution of heteromeric current to the total current measured at each point in the titrations using a prepulse to -40 mV to selectively inactivate heteromers. Because steady state inactivation of Kv2.1:Kv6.4 heteromers is incomplete, it leaves a pedestal current of a predicted size (Figure 3-3D, Figure 3-6D, Table 1). We therefore calculated the size of the heteromeric current as the size of the current removed by the -40 mV

74 prepulse plus the predicted size of the corresponding pedestal. Thus, the heteromeric current amplitude

was calculated as 1.38X and 1.44X the size of the current inactivated by the -40 prepulse for Kv6.4 WT

and Kv6.4-PIPIIV-Kv2.1CT, respectively. Plots of total current vs. calculated heteromeric current

(normalized to the homomeric Kv2.1 controls) are shown for the Kv6.4 WT and Kv6.4-PIPIIV-

Kv2.1CT titrations in Figure 3-8E,F. Both titrations approach 100% heteromeric current at a 1:10

Kv2.1:Kv6.4 cRNA ratio, suggesting that there was no significant offset in protein subunit translation that could explain the differences between current size in the titrations. Therefore, we focused on differences in the ability of 2:2R heteromers to conduct as the most likely explanation.

Cd2+ block of cysteine mutants confirms 2:2R functionality of Kv6.4-PIPIIV-Kv2.1CT

To test functionality of the 2:2R heteromers in an independent manner, we took advantage of the

phenomenon of Cd2+ coordination in a Kv2.1 pore cysteine mutant. Rat Kv2.1 I379C is blocked by

external cadmium, and block requires coordination by two I379C residues (Krovetz et al., 1997).

Furthermore, rat Kv2.1 WT:Kv2.1 I379C heteromers are blocked by Cd2+ only if the two cysteines are

present in two adjacent subunits; diagonally-opposed cysteines do not support cadmium block (Krovetz

et al., 1997). We hypothesized that if this mutant (I383C in mouse Kv2.1) was co-expressed with Kv6.4,

3:1R heteromers, which would have adjacent Kv2.1 I383C subunits, should be Cd2+-sensitive, while

2:2R heteromers, which would have only diagonally-opposed Kv2.1 I383C subunits, would be Cd2+-

resistant (Figure 3-9A). If 2:2R heteromers contribute to the heteromeric current, then we would expect

less cadmium block as the Kv6.4 expression ratio increases. We therefore tested Cd2+-sensitivity of

Kv2.1 I383C:Kv6.4 WT and Kv2.1 I383C:Kv6.4-PIPIIV-Kv2.1CT at multiple expression ratios to look for evidence of functional 2:2R heteromers.

75

Figure 3-9. Cd2+ block mediated by adjacent Kv2.1 I383C subunits predicts that 2:2R heteromers contribute to whole cell currents for Kv2.1:Kv6.4-PIPIIV-Kv2.1CT but not for Kv2.1:Kv6.4 WT. A) Channels predicted to form for Kv2.1 I383C (black circle with C) + Kv6.4 WT (left, red) or Kv2.1 I383C + Kv6.4 PIPIIV-Kv2.1CT (right, blue) at Kv2.1-biased or Kv6.4-biased expression ratios. Cd2+- coordination sites between adjacent cysteine-containing subunits are marked with red circles; all channels with Cd2+-coordination sites are predicted to be sensitive to Cd2+ block. Cd2+-resistant 2:2R heteromers (red box) are predicted to contribute to currents Kv6.4-PIPIIV-Kv2.1CT-biased mixes. B) Examples current traces before and after addition of 500 μM Cd2+ for Kv2.1, Kv2.1:Kv6.4 and Kv2.1:Kv6.4-PIPIIV-Kv2.1CT recorded in response to a 500 ms +40 mV step following a pre-pulse of 4 s to -100 mV. Oocytes were held at -70 mV. Experiments were carried out in the presence of 200 μM DTT and a 1:10 ratio was used for co-expression experiments. Scale bars indicate current amplitude and time. C) Fractional current remaining after 500 µM Cd2+ addition determined by measuring the peak current during the +40 mV test pulse using the protocol described in (B). Cd2+ block was significantly greater of both heteromer species compared to Kv2.1 homomers (*p<0.05, ANOVA + Tukey post-hoc, n = 10 for each). D) Examples traces before and after application of 500 μM Cd2+ for Kv2.1 I383C, Kv2.1 I383C:Kv6.4 WT, and Kv2.1 I383C:Kv6.4-PIPIIV-Kv2.1CT. Currents were recorded as in (B), and a 1:10 Kv2.1:Kv6.4 species expression ratio was used. E) 500 μM Cd2+-resistant current fraction (peak current during +40 mV test pulse) is shown for Kv2.1 I383C:Kv6.4 WT (red) and Kv2.1 I383C:Kv6.4-PIPIIV-Kv2.1CT (blue) at the indicated cRNA expression ratios. The solid gray and dashed gray lines mark the mean ± S.E.M. of the Cd2+-resistant current fraction for Kv2.1I383C homomers. Cd2+ block was significantly reduced at high expression ratios for Kv6.4-PIPIIV-Kv2.1CT compared to Kv2.1 the lowest expression ratio used (*, p < 0.05, ANOVA, n = 8-11 for each ratio). F) Kv2.1 I383C homomers (n = 10), Kv2.1 I383C:Kv6.4 M422C heteromers (n = 12) and Kv2.1:Kv6.4 M422C-PIPIIV-Kv2.1CT (n = 9) heteromers, which are all predicted to have 4 Cd2+-coordination sites are highly sensitive to block by 500 µM Cd2+ (**p < 0.01, ANOVA; *p < 0.05, ANOVA).

76 Kv2.1, Kv2.1:Kv6.4 WT (1:10), and Kv2.1:Kv6.4-PIPIIV-Kv2.1CT (1:10) were all blocked <

15% by 500 µM Cd2+ (Figure 3-9B,C) compared to >75% block observed for Kv2.1 I383C (Figure 3-

9D). While Cd2+ block of the WT heteromers was significantly greater than Kv2.1 homomers, the

degree of block was small compared to Kv2.1 I383C and thus does not interfere with detection of

enhanced block in channels containing adjacent Kv2.1 I383C subunits. 200 μM DTT increased Kv2.1

I383C current size by 17.9 ± 2.2 % (n = 9, p < 0.05, t-test), suggesting that partial basal oxidation may

interfere with conduction. Therefore, we included 200 μM DTT in all experiments to remove basal

oxidation as a potential confounding factor for comparisons of Cd2+-sensitivity. We then examined 500

µM Cd2+ block of Kv2.1 I383C:Kv6.4 WT heteromers at varying expression ratios (Figure 3-9D,E).

Cd2+ significantly slowed the activation of heteromeric channels, but we did not explore the mechanism.

We increased test pulse duration to allow currents to reach steady state for current amplitude

measurements. Kv2.1:Kv6.4 WT heteromers were blocked to a similar extent as Kv2.1 I383C

homomers (~75%), and the degree block was insensitive to Kv2.1 I383C:Kv6.4 WT expression ratio. In

contrast, Cd2+-block in Kv2.1 I383C:Kv6.4-PIPIIV-Kv2.1CT was sensitive to expression ratio (Figure

3-9D,E). Block was similar to Kv2.1 I383C homomers and Kv2.1 I383C:Kv6.4 WT heteromers with lower concentrations of Kv6.4-PIPIIV-Kv2.1CT cRNA, but the current became significantly more resistant to Cd2+ at high Kv6.4-PIPIIV-Kv2.1CT cRNA concentrations (Figure 3-9E). Some Cd2+ block

remained even at the most biased Kv6.4-PIPIIV-Kv2.1CT expression ratios, consistent with our TIRF

experiments, which show that ~40% of the channels at these ratios could still be Cd2+-sensitive 3:1R

heteromers. Full block by 500 µM Cd2+ could be restored by insertion of an equivalent pore cysteine

(M422C) into the Kv6.4-PIPIIV-Kv2.1CT chimera (Figure 3-9F), which ensures that all heteromer

stoichiometries produced by co-expression with Kv2.1 I383C would have subunits with adjacent

cysteines. The results of these Cd2+ block experiments provide a strong additional line of evidence that

77 Kv2.1:Kv6.4 2:2R heteromers do not contribute significantly to whole cell currents, while Kv2.1:Kv6.4-

PIPIIV-Kv2.1CT 2:2R heteromers do contribute significantly.

Discussion

Our results agree with previous work on Kv6.4 and Kv9.3 which suggest that the predominant

stoichiometry for mammalian Kv2 family regulatory subunit-containing heteromers is 3:1R

(Kerschensteiner et al., 2005; Bocksteins et al., 2017). However, our use of a single molecule technique

(TIRF photobleaching) here revealed that 2:2R heteromers are also formed. Because 2:2R heteromers

represent only about one quarter of all heteromers even at expression ratios highly biased in favor of

Kv6.4, 2:2R formation appears to be inefficient and would likely not have been detected in previous

studies using macroscopic analysis techniques such as FRET (Kerschensteiner et al., 2005) or gating

current components (Bocksteins et al., 2017). T1 self-incompatibility is a key feature of Kv2 family regulatory subunits that helps prevent homophilic subunit interactions, and should rule out formation of

4R, 1:3R and 2:2R tetramers with adjacent regulatory subunits. However, T1 self-incompatibility should not block formation of 2:2R heterotetramers with diagonally-opposed regulatory subunits, the major

2:2R species that would be expected to assemble from Kv2.1:Kv6.4 heterodimers (Figure 3-1C). Our results agree with these expectations for the role of T1 self-incompatibility in assembly of Kv2.1:Kv6.4 heteromers: only 3:1R and 2:2R heteromers were detected in TIRF. However, the ratio of these two stoichiometries was unaffected when the self-incompatible Kv6.4 T1 was replaced with the Kv2.1 T1,

(Figure 3-5A,D), suggesting that T1 does not play a major role in driving preferential 3:1R assembly for

Kv2.1:Kv6.4 and may not be fully responsible for Kv6.4 subunit self-incompatibility.

78 While we observed formation of 2:2R Kv2.1:Kv6.4 heteromers, our results suggest that only the

more prevalent 3:1R stoichiometry is functional (Figures 3-8 and 3-9). The unusual activation gate of

Kv6.4 and its P3T and V6F substitutions play a key role in limiting 2:2R formation and conduction

(Figures 3-5, 3-8, 3-9). We did not experimentally examine the mechanism through which these

mutations restrict stoichiometry, but we made draft homology models of the pore of Kv2.1,

Kv2.1:Kv6.4 heteromers (3:1R and 2:2R) and Kv6.4-PIPIIV (2:2R) to try to gain insights. Open models

and closed models were based on the Kv1.2 computational models presented by Pathak et al. (Pathak et al., 2007). The pore domain (S5-S6) includes numerous signpost residues universally conserved among

Kv1.2, Kv2.1 and Kv6.4 that allow for precise structural alignment (Figure 3-10A). Views of the closed conduction pathway from an extracellular perspective, color coded by hydrophobicity (Figure 3-10B-E),

suggest a feature of the Kv6.4 activation gate that could potentially impact 2:2R heteromer formation. In

the Kv2.1 homomer model, the hydrophobic inner gates form a vapor lock around the conduction

pathway that excludes water from the intersubunit interfaces and allows for strong hydrophobic

interactions between subunits (Figure 3-10B). A vapor lock securing a closed state of a channel is a

phenomenon common among different unrelated channel families of pro- and eukaryotes, and can be

also observed in model hydrophobic pores (Beckstein and Sansom, 2003; Roth et al., 2008; Anishkin et

al., 2010). Surface tension of water meniscus above and below the “dry” gate is expected to provide a

force contracting the pore walls together, stabilizing the pore assembly. In contrast, a hydrophilic cleft

bisects the gate area in Kv2.1:Kv6.4 2:2R heteromers, breaking the vapor lock (Figure 3-10D). With the

single Kv6.4 subunit in the 3:1R heteromer, hydrophobicity is locally reduced, but there is no trans-pore

hydrophilic cleft (Figure 3-10C). When Kv6.4-PIPIIV is substituted for WT Kv6.4 in the 2:2R model,

the hydrophilic cleft is eliminated and the vapor lock is restored (Figure 3-10E). This suggests that the

hydrophilic cleft in the 2:2R Kv2.1/Kv6.4 heteromers could potentially weaken tetramer stability and

79 may play a role in favoring formation of 3:1R heteromers over 2:2R heteromers even at high Kv6.4

concentrations.

Figure 3-10. Structural homology models of Kv2.1 homomers and Kv2.1:6.4 heteromers suggest the Kv6.4 activation gate might perturb pore stability and function. A) Sequence alignment of the Kv1.2, Kv2.1 and Kv6.4 pore domains (PDs). Transmembrane domains S5 and S6 and the K+ selectivity filter are underlined. Residues identical or conservatively substituted across all three sequences are shaded red and black, respectively. The six amino acid activation gate is boxed with positions 1-6 labeled. Note there are identical signpost residues throughout the PDs allowing precise alignment between Kv1.2 (determined structure, (Long et al., 2005a)), Kv2.1 and Kv6.4 (structural models based on Kv1.2 presented here). B – E) Snapshots of the closed conduction pathway viewed from the extracellular side for structural models of a Kv2.1 homomer (B), a Kv2.1:Kv6.4 3:1R heteromer (C), a Kv2.1:Kv6.4 2:2R heteromer (D) and a Kv2.1:Kv6.4 2:2R heteromer with the Kv2.1 activation gate (PIPIIV) substituted for the Kv6.4 activation gate (PATSIF) (E). The protein backbone (thin tubes) is colored white for Kv2.1 and light-purple for Kv6.4. Side chains (van der Waals representation) are shown for position 2, 3 and 6 of the activation gate and colored according to hydrophobicity index in the Kyte and Doolittle scale with their values ranging from 1.8 for Alanine (green) to 4.5 for Isoleucine (white) (scale provided below panels).Residues which line the conduction pathway (I2, V6 in Kv2.1; A2 and F6 in Kv6.4) are labeled in panel C. Yellow ribbons in all panels highlight the position of the gate backbone. In Kv2.1, there is an expected intersubunit hydrophobic vapor lock at the activation gate. While insertion of a single Kv6.4 subunit (C) is well-tolerated, insertion of two diagonally-opposed Kv6.4 subunits (D) simultaneously introduces a hydrophilic cleft that bisects the gate and increases the distance between diagonally-opposed subunits. Both changes favor disruption of the intersubunit vapor lock and could

80 hypothetically reduce tetramer stability at the gate intersubunit interface. The bottom two rows are the view from the cytoplasmic side - one row for the closed state (panels F-I) and another for the open (panels J-M). Side chains at the gate constriction are shown in space fill and colored by the residues type (Kv2.1: I2, white; P3, cyan; V6 gray; Kv6.4: A2, gray; T3, brown with red hydroxyl; F6, pink). In the closed models, the hydroxyl group of T3 in the Kv6.4 gate faces the neighboring subunit (G, H) and thus contributes to the hydrophilic cleft observed in the extracellular view of the 2:2R tetramer (D). In the open conformations, a dehydrated K+ ion (CHARMM36 radius, 1.76 Å) is depicted with a blue circle in the pore opening at the narrowest point of the conduction pathway in the activation gate region. A red dashed line surrounds the portion of the opening accessible to the center of the dehydrated K+ ion (red dot) as determined by rolling the ion against the sidechains lining the pore. Blue mesh covers the inaccessible region of the pore. The black circle around the ion roughly approximates the radius of the first hydration shell (i.e., K+ radius plus the diameter of TIP3P water molecule in CHARMM36). While not an explicit simulation in all-atom setting, it is nevertheless obvious that F6 in the Kv6.4 activation gate narrows the gate opening proportional to the number of Kv6.4 gates. In the 2:2R conformation, almost complete dehydration of a K+ ion would be needed for passage. Alternatively, more drastic rearrangements of the backbone than can be observed in our short vacuum simulations might reduce the constriction, but still block conduction by disrupting gating as observed for the F6 substitution in Shaker (Kitaguchi et al., 2004). Note that substitution of the Kv2.1 activation gate into Kv6.4 in the 2:2R simulation restores both the hydrophobic vapor lock of the closed state (E, I) and a wide conduction pathway to the open state (M).

In addition, intracellular views of the pore at the activation gate show that the combination of T3

and F6 of the Kv6.4 gate significantly narrows the conduction pathway in the open state, especially in

2:2R heteromers (Figures 3-10F-M). The opening at the level of F6 in the 2:2R heteromer would

require significant dehydration of a K+ ion for it to pass, suggesting that conduction for 2:2R heteromers

could be energetically unfavorable. The conduction pathway diameter in 2:2R Kv2.1:Kv6.4-PIPIIV

heteromer model, in contrast has a similar diameter to that of Kv2.1 homomer model. However, the

V6W mutation in Shaker and Kv1.2 does not completely block conduction but instead may energetically

trap channels in the closed state, propagating structural changes to the selectivity filter which alter gating

or conduction (Kitaguchi et al., 2004; Pau et al., 2017). Shaker V6W channels do conduct in a diagonal

2 WT:2 V6W configuration (albeit poorly), and conduction in homomeric V6W channels can be restored by a compensatory gate mutation P3Q that stabilizes the open state (Sukhareva et al., 2003;

Kitaguchi et al., 2004). It should also be noted that because of the ability of individual pore mutations to propagate structural rearrangements broadly through the pore (Pau et al., 2017), the functional

81 consequences of specific gate substitutions might vary significantly from channel to channel.The

significance of the steric barrier observed in the Kv2.1:Kv6.4 2:2R model thereofore requires

experimental clarification. What the models do show is that the Kv6.4 gate will likely perturb pore

structure, and that the degree of perturbation increases significantly when two Kv6.4 subunits participate

in the gate formation (Figure 3-10).

Because we were not able to reach 100% 2:2R formation with the Kv6.4-PIPIIV-Kv2.1CT or

Kv6.4-Kv2.1T1-PIPIIV-Kv2.1CT chimeras even at highly biased expression ratios, regions of the channel other than T1 and the gate are also likely to play a role in suppression of the 2:2R stoichiometry.

The inability of the Kv2.1 T1 to restore assembly of channels with more than 2 regulatory subunits in the Kv6.4-PIPIIV-Kv2.1CT background further suggests that other channel regions could also contribute to Kv6.4 self-incompatibility. A broader analysis of pore interface residues that differ between Kv2.1 and Kv6.4 might reveal additional factors limiting 2:2R formation. We focused on the gate here because of the consistent and striking evolutionary divergence of the gate sequence among regulatory subunits (Figure 3-2). It is also possible that 2:2R formation could be limited over the expression ratio range we tested if Kv2.1 simply has higher affinity for itself than Kv6.4, independent of any structural perturbations. We did not test higher expression ratios here because the concentrations of cRNA required compromised oocyte health. However, if Kv2.1 homodimerization is preferred to

Kv2.1:Kv6.4 heterodimerization, our data suggests that the T1 domain alone does not account for the affinity difference because we were unable to increase 2:2R formation by substituting the Kv2.1 T1 into

Kv6.4 or Kv6.4-PIPIIV-Kv2.1CT (Figure 3-5D). Furthermore, differential affinity does not contradict the role we found for the Kv6.4 gate in limiting 2:2R formation.

Based on the results presented here, we propose a two-step model for evolution of the regulatory

phenotype in Shaker family channels that predicts degeneration of the highly-conserved activation gate

82 sequence and opens an evolutionary path to a 3:1R stoichiometry (Figure 3-11). In the first step, the

regulatory phenotype, which is characterized by an inability to form homotetramers, is established by a

self-incompatibility mutation, most likely in T1 as depicted here (Figure 3-11B). Both 3:1R and 2:2R

stoichiometries are predicted to be functional because T1 self-incompatibility mutations are unlikely to

affect channel function beyond assembly. The self-incompatible mutated channel produced in Step 1 should remain under positive selection to maintain functional channel numbers. The original self-

incompatibility mutation could in theory occur outside T1, provided it does not disrupt heteromer

function. While the precise nature of the self-incompatibility mutations leading to the mammalian Shab

regulatory subfamily is not clear, the T1 is a likely spot because their T1 domains are self-incompatible

(Salinas et al., 1997b; Kramer et al., 1998; Ottschytsch et al., 2002). Furthermore, a single amino acid change in the T1 of Kv1.3 is sufficient to create T1 self-incompatibility while preserving the ability to form heteromers with WT channels (Robinson and Deutsch, 2005). Nevertheless, our results indicate that Kv6.4 self-incompatibility extends beyond T1. Regardless of the location of the original Step 1 mutation, self-incompatibility limits functional channels to no more than 2 mutated subunits/tetramer

(Figure 3-1). Selective pressure on the gate sequence is therefore reduced, and the gate can now accumulate mutations that can be functionally accommodated in no more than two subunits of a tetramer. In Step 2 of the model, this evolutionary drift of the gate sequence over time could eventually result in mutations that are tolerated only in a single subunit, thus fixing the functional stoichiometry at

3:1R (Figure 3-11B). While some subunits in step 2 may be lost to non-functional stoichiometries as we observe for Kv2.1:Kv6.4, advantageous new biophysical properties could maintain positive selection.

The physiological advantage of a delayed rectifier with a low activation threshold, slower kinetics and closed state inactivation (Figure 3-3) in neurons expressing Kv6.4 remain to be determined. We believe it is unlikely that evolution of the regulatory phenotype would begin with a restrictive gate mutation in

83 Step 1, because gate incompatibility in the presence of intact assembly would tie most of the expressed

subunits up in non-functional tetramers (Figure 3-11C). The gate mutation would therefore serve as a

strong dominant negative and would likely face immediate negative selection from the outset. This may

help explain why the gate is so highly conserved in self-compatible-homomer forming Shaker family subunits.

Figure 3-11. Model for the evolution of Shaker family regulatory subunits adopting a 3:1R functional stoichiometry. A) A pre-condition for the evolution of the regulatory phenotype in Shaker family channels is co-expression of two subunits (A and B) from the same subfamily (Kv1, Kv2, Kv3 or Kv4) in at least a subset of cells. Each subunit forms homomeric channels, and because they have cross- compatible T1 domains, they can also form heterotetramers in 4 possible stoichiometries (3A:1B, 2A:2B adjacent, 2A:2B diagonal, 1A:3B). All channels formed are functional, indicated by the presence of a K+ ion (black circle) in the pore. B) Model 1, our favored model, provides a 2-step path to evolution of the

84 regulatory phenotype (in subunit B in this example) functioning in a 3:1R stoichiometry. In Step 1, mutation(s) in subunit B establish self-incompatibility (shown here as mutations in T1), but do not eliminate cross-compatibility with subunit A. This restricts channel assembly to 3 possible stoichiometries (4A, 3A:1B, 2A:2B diagonal); all assembled channels are functional. Expression of subunit B increases the number of channels formed (relative to expression of subunit A alone) and is therefore critical for maintaining currents at or near their pre-mutation starting levels. Step 1 effectively established the regulatory phenotype for subunit B. In Step 2 of the model, Subunit B accumulates gate mutation(s) tolerated in only a single subunit, disrupting 2:2R assembly, conduction or gating. This establishes 3:1R as the single functional heteromer stoichiometry. Loss of function in 2:2R channels is depicted by replacement of the K+ ion with a black X. Dominant-negative suppression of current will occur at highly subunit B-biased expression ratios as few A-A contacts will form during assembly, but reasonable currents will remain with balanced expression. C) In the alternative Model 2, subunit B gate incompatibility evolves first, while self-compatibility remains. This results in strong dominant negative suppression because most subunits are tied up in non-functional channels. Model 2 is likely a dead end because the gate mutation will engage negative selection to preserve current levels.

This model predicts the gate consensus sequence degeneration we observe in all independent

evolutions of the regulatory subunit phenotype within the Shaker family. A 3:1R stoichiometry could be

the most likely long-term outcome because it is logical that the gate would eventually acquire a mutation that can be tolerated in only a single subunit. While maintenance of two functional stoichiometries might initially be selectively neutral, fixation on a single functional stoichiometry could gain a selective advantage over time as the regulatory subunit accumulates mutations that would functionally differentiate 3:1R from 2:2R channels. Nevertheless, the model does not require that each evolutionary

instance of the regulatory subunit phenotype will fix at 3:1R, or that observed regulatory subunit group

will have reached a point of fixation. We believe 2:2R fixation should be less common because 1) The

regulatory phenotype is most likely first established with a loss-of-function mutation in assembly and 2) it is unlikely that any gate mutations are better tolerated in a 2:2R configuration than a 3:1R configuration. Fixation on a 2:2R stoichiometry is possible within this model if this stoichiometry has unique, positively-selected functional properties, but it might require secondary mutation(s) that favor heterodimer formation during assembly. Once a 2:2R stoichiometry is favored, gate mutations tolerated in only a single subunit would be selected against. A cnidarian Kv4 regulatory subunit jShalγ with only

85 a single gate substitution (PVPIIQ) has been predicted to form functional channels in a 2:2R

stoichiometry based on examination of the stoichiometry of N-type inactivation (Jegla and Salkoff,

1997), but in the absence of TIRF photobleaching analysis, it is unclear if 3:1R heteromers do or do not

form.

Further functional characterization of independent evolutionary instances of regulatory subunits

will be needed to test whether this model predictably describes the evolution of the regulatory subunit

phenotype across the Shaker family. It predicts that self-incompatibility should be a universal feature of

regulatory subunits, and that an exclusive 3:1R functional stoichiometry should be common. It would

also be interesting to see if there is correspondence between the predicted severity of gate mutations and

regulatory subunit stoichiometry. Regardless, degeneration of the highly-conserved activation gate

sequence appears to have high predictive value for identifying evolution of the regulatory phenotype in

Shaker family. Another example of preferential assembly of a 3:1R stoichiometry in the voltage-gated

cation channel superfamily are the heteromeric rod cyclic nucleotide-gated channels (Weitz et al., 2002;

Zheng et al., 2002; Zhong et al., 2002). These channels lack the Shaker T1, but a cytoplasmic coiled-coil

domain in the C-terminus seems to play an analogous role; it is formed by 3 helices, one each

contributed by the CNG1A subunits, while the regulatory CNGB1is excluded and thus limited to a

single subunit/channel. Our results suggest it would be interesting to look for co-evolution of gate

sequences and the regulatory phenotype across the breadth of the CNG channel family.

86 Table 3-1. Boltzmann fit parameters for Kv2.1, and Kv2.1:Kv6.4 heteromers

Channel Activation (GV) Steady State Inactivation (SSI)

a b c V50 Slope V50 Slope Pedestal

Kv2.1d 7.2 ± 1.0 14.1 ± 0.3 -10.5 ± 0.6 7.5 ± 0.4 0.67 ± 0.02

+ Kv6.4 WTe -12.8 ± 1.4 15.1 ± 0.4 -62.6 ± 0.6 7.7 ± 0.2 0.28 ± 0.01

+ Kv6.4-PIPIIV- -16.3 ± 0.5 14.2 ± 0.6 -82.3 ± 1.4 12.2 ± 0.5 0.3 ± 0.01 KV2.1 CTf aMidpoint of Single Boltzmann fit in mV (mean ± S.E.M.) bSlope of Single Boltzmann fit (mean ± S.E.M.) cNon-inactivating pedestal current fraction determined from Boltzmann fit (mean ± S.E.M.) dN = 7 e,fN = 10

87 References

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91 Chapter 4

Activation of invertebrate TRPV heterotetramers by the vitamin B3 metabolite nicotinamide

In this chapter, I will discuss a project that identified the vitamin B3 metabolite nicotinamide as an agonist for a heteromeric TRPV channel in C elegans formed from the OSM9/OCR4 subunits. The role of nicotinamide as a TRPV channel opener was found to be evolutionarily conserved in the Drosophila TRPV orthologs Inactive and Nanchung. Additionally, the stoichiometry of the OSM9/OCR4 heteromeric channel was determined using

Total Internal Reflectance Fluorescence (TIRF) microscopy. To the best of my knowledge, this was the first time currents from a C elegans TRPV channel were recorded in a heterologous expression system; previous attempts at current expression for these channels were not successful and it had been speculated that an additional chaperone protein was necessary. We demonstrated that coexpressing the two TRPV subunits OSM9 and OCR4 was sufficient for heterologous expression and that no additional genes or factors were necessary. This project was published in Nature Communications in 2016. I performed the electrophysiology experiments myself and collected and analyzed the TIRF data in collaboration with Keith Mickolajczyk. Awani Upadhyay, who is first author in the 2016 paper, made the initial discovery of nicotinamide regulating the C elegans TRPV homologs based on her experiments aimed at understanding how excess nicotinamide specifically induces cell death in a number of C elegans cells, including the touch-sensitive Outer Labial Quadrant (OLQ) neurons. While these experiments are elegant, they are beyond the scope of this thesis. I will limit my discussion to our investigations of the invertebrate TRPV channel family and the role of nicotinamide in regulating TRPV activity. The experiments shown in this chapter demonstrate using the heterologous Xenopus system that nicotinamide could potentially be an endogenous agonist to invertebrate TRPV channels. Additionally, they prove that OSM9 and

OCR4 combine in a 2:2 stoichiometry. Figures 4-1, 4-2, 4-3 and 4-4 are taken from Figure 4, Figure 8,

Supplementary Figure 4, and Figure 5 of the 2016 Nature Communications paper respectively (Upadhyay et al.,

2016).

92 Introduction

The transient receptor potential (TRP) family of channels are a diverse group of tetrameric non-specific cation channels that are gated by various types of thermal, mechanical and chemical stimuli. Structurally, each subunit consists of a large family-specific intracellular N-terminus preceding 6 transmembrane domains, with a pore domain composed of the fifth and sixth transmembrane domains along with their intermediary linker (Li et al., 2011). The transmembrane core is then followed by a family-specific cytoplasmic domain. In addition to temperature-sensitivity, mammalian TRPV1 channels are known to be activated by capsaicin, the chemical responsible for the “heat” from chili peppers (genus Capsicum), and other related vanilloid compounds (Caterina et al., 1997). While the presumed mechanism of the capsaicin-TRPV interaction has been inferred for a while on the basis of screening tools and mutagenesis, the nature of the interaction has been further characterized only recently (Yang et al., 2015). Although only TRPV1 channels are known to be sensitive to capsaicin (Caterina et al., 1999), other vertebrate TRPV channels can be be activated by naturally occurring and synthesized compounds

(Watanabe et al., 2002; Xu et al., 2006). Importantly, the activation of mammalian TRPV channels by capsaicin and other compounds has been utilized in heterologous models to test how these channels can be regulated through a structure-function point of view (Voets et al., 2002).

As mentioned in Chapter 1, invertebrate TRPV channels appear to be gated primarily by mechanical stimuli, such as touch or sound, and do not show any temperature-sensitivity (Colbert et al., 1997; Goodman and

Schwarz, 2003; Gong et al., 2004; Zhang et al., 2013). Invertebrate TRPV channels are also notably interdependent, which means at least two different types of TRPV subunits are required to make a functional

TRPV channel (Tobin et al., 2002; Gong et al., 2004). Neurons expressing invertebrate TRPV channels display chemosensory functions, and this is dependent on TRPV function (Colbert et al., 1997; de Bono et al., 2002;

Nesterov et al., 2015). Despite this knowledge of how invertebrate TRPV channels could be regulated, currents from invertebrate TRPV channels have never been heterologously expressed. This precludes the ability to investigate how TRPV channels could have evolved from a functional perspective. If the character of the wild- type TRPV current is unknown, it is not possible to understand how perturbations in primary structure can

93 influence function in a hypothesis-driven manner. The experiments shown here demonstrate that nicotinamide can

be used to activate invertebrate TRPV channels in a heterologous system. The nicotinamide-TRPV channel

interaction is shown to be evolutionarily conserved in the invertebrate TRPV lineage. Additionally, it is shown

that the TRPV channel formed by OSM9 and OCR4 has a 2 to 2 stoichiometry.

Methods

Molecular cloning and cRNA synthesis

cRNAs were made from pGEMHE-OSM9 (gift of Dr. C. Bargmann, Rockefeller University) and pOX-

OCR4 (OCR4 cDNA was a gift of Dr. Y. Kohara, National Institute of Genetics, Japan) using mMessage

Machine kits (Ambion, TX). GFP tags were linked to N-terminus of OSM9 and the C-terminus OCR4 using the

flexible linker QQQGQQA.

Xenopus oocyte recordings

Oocytes recordings were carried as described in Chapter 2. The base recording solutions consisted of the following: 98 mM NaCl, 2 mM KCl, 1 mM MgCl2, 1 mM CaCl2, and 5 mM HEPES (pH 7.5 with NaOH).

Nicotinamide (gift from Dr. W Hanna-Rose, The Pennsylvania State University) was added to the base recording

solution at the desired concentrations.

TIRF Photobleaching

TIRF photobleaching experiments were carried out as described in Chapter 2 and Chapter 3.

94 Sequence Alignment

Sequences shown in Figure 4-6 were aligned using CLUSTALW as implemented by JALVIEW

(Waterhouse et al., 2009). Alignment was exported to MEGA7 (Kumar et al., 2016) for generation of the

Phylogenetic Tree shown in Figure 4-5.

Results

Nicotinamide is an OSM9/OCR4 channel agonist

OSM9 and OCR4 were expressed individually and in combination in Xenopus oocytes to test nicotinamide-dependent activation using two-electrode voltage clamp (Figure 2-2). Expressing OSM9 or OCR4 by themselves did not produce any discernible nicotinamide-activated currents. However, co-expressing OCR4 and OSM9 resulted in a large, nicotinamide-dependent current that appeared to reverse close to 0 mV (Figure 4-

1A,B). This nicotinamide-evoked current is concentration dependent, with an estimated K1/2 value of 63.3 ± 13.2

µM (Figure 4-1C). Another form of vitamin B3 known as nicotinic acid, produced less than 300 nA of current even at concentrations as high as 130 mM (Figure 4-1D). This pales in comparison to the large currents produced by only 100 µM nicotinamide. Based on these results, it is clear that the nicotinamide form, but not the nicotinic acid form, of vitamin B3 is a potent agonist for the heteromeric TRPV channel formed by OSM9 and OCR4.

Interestingly, nicotinic acid has been shown to regulate vertebrate TRPV channels (Ma et al., 2014; Ma et al.,

2015). It is not clear from this data if nicotinamide acts on the OSM9/OCR4 channel from the extracellular environment or the intracellular environment. Nicotinamide has a partition coefficient (log P) of -0.37 (NCBI), suggesting it is partially hydrophilic but may still be able to cross the cell membrane and act from within the cytoplasmic compartment. Regardless of the site of action, the effect of nicotinamide is direct and concentration dependent. The Hill coefficient, which is a measure of cooperativity of agonist binding, estimated from the affinity curve is estimated at 1.8 ± 0.2. The theoretical upper limit of the Hill coefficient is the number of ligand

95 binding sites present in the protein (Stefan and Le Novere, 2013); a value for the Hill coefficient greater than one

therefore implies multiple nicotinamide binding sites are present within the OSM9/OCR4 heterotetramer,

although the precise number of binding sites is not clear.

Figure 4-1. Nicotinamide activates the heteromeric TRPV channel formed by the co-expression of OCR4 and OSM9 protein subunits in Xenopus laevis oocytes. A,B) Coexpressing OSM9 and OCR4 led to large nicotinamide activated currents. Expression of OSM9 or OCR4 by themselves did not generate such nicotinamide-dependent currents. Currents were recorded in response to 2 s voltage ramps from -100 to +100 mV from a -20 mV hold. Current size was measured at -100 mV (star). C) The half maximal concentration (K1/2) for current activation by nicotinamide estimated by fitting Hill curve is approximately 63 µM. Data in 4-1C shows mean ± S.E.M. (n = 5- 9). D) Current is not induced in OCR4 / OSM9 channel by 5 mM nicotinic acid (-.154 ± .018 µA, n = 6). There is a slight increase in the conduction of the OSM9/OCR4 channel upon exposure to 130 mM NA (-.269 ± .026 µA, n = 4). Though this is a statistically significant increase (p<.05, two-tailed t-test), this high (5 mM) concentration of NA elicited over 200-fold less current compared to only 100 µM nicotinamide (-36.5 ± 1.0 µA, n = 4).

Nicotinamide-evoked activity is conserved in the Drosophila TRPV homolog formed by Inactive and Nanchung

Given the similar gating and formation mechanisms among invertebrate TRPV channels, I wanted to test whether the homologous Drosophila TRPV channel created by the component genes Inactive (IAV, equivalent to

OSM9) and Nanchung (NAN, equivalent to OCR4) is also sensitive to nicotinamide. As with co-expression of

96 OCR4 and OSM9, there is a large current in response to nicotinamide in oocytes co-expressing IAV and NAN

(Figure 4-2A). The estimated K1/2 value for nicotinamide is 14.4 ± 1.0 µM (Figure 4-2B). Again, the agonist

activity of nicotinamide is not shared by the related metabolite nicotinic acid (Fig. 4-2C), and neither NAN (n=6

eggs) nor IAV (n=5 eggs) alone produced nicotinamide-responsive channels. These results suggest that the

agonist activity of nicotinamide on the invertebrate TRPV channel is evolutionarily conserved.

Figure 4-2. Fly TRP channel subunits Inactive (IAV) and Nanchung (NAN) make a heteromeric ion channel that is activated by nicotinamide. A) Example traces of the effect of nicotinamide on the IAV/NAN heteromer. Oocytes co-expressing NAN and IAV demonstrate large nicotinamide activated currents. Currents were recorded using the same protocol shown in Figure 4-1. Current size was measured at -100 mV (star), and the half maximal concentration (K1/2) for current activation by nicotinamide was determined from a Hill plot. (n = 5-13 per nicotinamide concentration). B) Affinity of nicotinamide to the IAV/NAN heteromer appears to have almost a five-fold increase in affinity for nicotinamide compared to the OSM9/OCR4 channel (Figure 4-1C). C) Current is not induced in IAV/NAN coexpressing oocytes by 5 mM nicotinic acid (-.188 ± .020 µA, n = 3). There is a slight increase in current upon exposure to 130 mM nicotinic acid (-.422 ± .040 µA, n = 4)). Though this is a statistically significant increase (p<0.01, two-tailed t-test), this high (130 mM) concentration of nicotinic acid elicited over 150-fold less current compared to only 100 µM nicotinamide (-28.7 ± 2.5 µA, n = 3).

OSM9 and OCR4 form a 2:2 heterotetramer

Because the above experiments suggest that functional channels do form and traffic when OSM9 and

OCR4 are coexpressed, heterologous expression appears to be a viable way of investigating structure-function relationships of invertebrate TRPV channels. The question still remains however, of the stoichiometry of the

OSM9/OCR4 heteromer. The stoichiometry of this channel is important to know in order to design appropriate experiments to answer future questions regarding structure-function relationships. Total Internal Reflectance

Fluorescence (TIRF) microscopy was used to determine this stoichiometry; for a discussion on the specific

97 protocols and equipment, see Chapter 2. Expressing OCR4-GFP by itself (Figure 4-3B), or GFP-OSM9 did not produce any stable photobleaching spots at the membrane, suggesting functional channels do not form from

OCR4 or OSM9 alone. The data shown in Figure 3-4A,B demonstrates that our TIRF protocol could identify

photobleaching of wild-type Kv2.1, a known tetramer, in 4 steps with an estimated GFP fluorescent probability of

0.69. I used this value for the fluorescent probability to estimate the proportion of channels containing 1 or 2

GFP-tagged subunits.

Figure 4-3. TIRF photobleaching demonstrates channels that traffic to the membrane when OSM9 and OCR4 are coexpressed, but not individually. A) Oocytes injected with OCR4-GFP and OSM9 displayed many fluorescent spots on the membrane at the beginning of the observation period (top panel, time 0). Photobleaching is irreversible and is expected to occur in discrete steps. Therefore, channels were only included if the bleaching steps were demonstrable without artifacts or recovery following complete bleaching (see I and II in A). Spots that did not bleach in a clear manner (III), behaved erratically following bleaching (IV) or lost fluorescence in a non- stepwise fashion (V) were not included in the analysis since the number of photobleaching steps could not be reliably counted. In the example figure every spot that was excluded from the analysis is indicated with a yellow outline. Every spot that was included as a final data point is indicated by a blue (2-step) or green (1-step) outline. Most spots were no longer visible at the end of the observation period (bottom panel, time 4 min). B) Oocytes injected with OCR-GFP alone showed few fluorescent spots on the membrane at the beginning of the observation period (bottom panel, time 0) and these spots failed to demonstrate stable photobleaching.

No stable fluorescent spots were observed at the plasma membrane upon injection of OCR4-GFP (Figure

4-3B) or GFP-OSM9 alone. However, when OCR4-GFP was coexpressed with OSM9 (Figure 4-3A) or GFP-

OSM9 was coexpressed with OCR4 at a 1:1 RNA ratio, stable fluorescent spots (Figure 4-3A) that bleached in

98 either one or two steps were observed at the membrane (Figure 4-4A,C). There did appear to be one spot that bleached in three steps for each experiment. Because the frequency of these observations are low and contradictory to a uniform tetrameric stoichiometry, these observations are likely artifacts. Using the above- mentioned GFP detection efficiency value of 0.69 in Chapter 3 along with the distribution of just the one and two steps for each experiment, it is estimated that greater than 90% of OSM9/OCR4 channels adopt a 2:2 stoichiometry (Fig. 4-4B,D). This suggests that the overwhelming majority of channels that reach the cell membrane consist of two of each subunit.

Figure 4-4. OSM9/OCR4 channels predominantly form 2:2 heteromers in Xenopus. A) Frequency of the number of bleaching steps observed for 71 channels from oocytes expressing GFP-OSM9 with OCR4. The inset shows example fluorescent traces for two-step (top) and one-step (below) photobleaching. B) Stoichiometry distribution calculated from (A), assuming a 0.69 GFP detection efficiency as determined by GFP-Kv2.1 (Figure 3-4). C) Frequency of the number of bleaching steps observed for 66 channels from oocytes expressing OCR4-GFP with OSM9. D) Stoichiometry distribution calculated from (C), assuming a GFP photobleaching efficiency as shown in Figure 3-4.

Discussion

It has previously proven difficult to establish channel activity for the C. elegans TRPV channels in a heterologous system (Colbert et al., 1997; de Bono et al., 2002; Jose et al., 2007). However, the results presented

99 in this chapter suggest that other non-TRPV subunits or factors are not required for heterologous current

expression. From these experiments, I have confirmed that the vitamin B3 metabolite nicotinamide is an agonist of

a heteromeric C. elegans TRPV channel that is composed of two OCR4 and two OSM9 subunits. Moreover, these experiments show that nicotinamide-evoked activity is evolutionarily conserved among invertebrate TRPV channels as the compound also activates the orthologous Drosophila IAV/NAN channel. It is not clear from the data above if nicotinamide is activating the OSM9/OCR4 and IAV/NAN channels through extracellular or intracellular domains. The activation of TRPV channels by nicotinamide appears to occur with fast kinetics, but the experiments described above do not have the resolution to distinguish the nature of the binding site due to the membrane permeability of nicotinamide.

Nicotinamide as a potential endogenous ligand for invertebrate TRPV channels

The hypothesis that nicotinamide could activate invertebrate TRPV channels derived from the observation that the mutant osm-9 and ocr-4 strains of C elegans demonstrated resistance to nicotinamide-induced cell death.

It was proposed that opening of the OSM9/OCR4 TRPV channel leads to an influx of calcium that is toxic to the cell (Upadhyay et al., 2016). Because of the high levels of nicotinamide required to induce cell death, it was initially unclear if nicotinamide was an endogenous regulator of invertebrate TRPV channel activity. Indeed, the estimated affinity of nicotinamide to the OSM9/OCR4 heteromer is approximately 63 µM, which is almost six times greater than the estimated concentration of nicotinamide at basal levels (Wang et al., 2015). Based on the affinity curve shown in Figure 4-1C, the level of channel activity at 11 µM nicotinamide is roughly 5-10% of the maximal current. Though this may seem like a small amount of channel activity, it may be sufficient to regulate physiologic responses; this is because very little channel activity is actually needed to exert large effects on cells.

In order to predict what types of physiologic responses require nicotinamide modulation of TRPV channels, it may first be helpful to recognize the biochemical functions of nicotinamide and other pathways it is involved in. Nicotinamide is a form of vitamin B3; vitamin B3 is required to generate nicotinamide adenine

100 dinucleotide (NAD). Although NAD is classically known for its role as a key electron carrier in oxidative

phosphorylation, it can also be consumed in ADP-ribose transfer reactions by enzymes such as poly-ADP-ribosyl polymerases (PARPs) and sirtuins (Belenky et al., 2007; Sauve, 2008). Nicotinamide is a byproduct of NAD consumption by these enzymes. Activity of PARPs and sirtuins is typically upregulated under cytotoxic stresses that lead to DNA damage; these enzymes are therefore important in cytoprotection (Guarente, 2006; Schreiber et al., 2006). Importantly, high levels of nicotinamide can inhibit the activity of these enzymes and can promote apoptosis (Luo et al., 2001; Bitterman et al., 2002). Increased levels of nicotinamide through NAD consumption, likely by PARPs and sirtuins, could therefore serve to prime TRPV channels to respond to milder mechanical stimuli. This type of sensitization could enable more robust responses to stimuli that may not typically be harmful, but could exacerbate a prior injury.

Phylogenetic analysis of TRPV evolution

Phylogenetically, OSM9 is homologous to Inactive, and Nanchung is homologous to the OCR subfamily

(Figure 4-5). All four subunits are also more closely related to each other than they are to vertebrate TRPV

channels. NAN and IAV are the only Drosophila TRPV proteins and are orthologs of OCRs and OSM9,

respectively (Matsuura et al., 2009; Venkatachalam et al., 2014). The presence of multiple OCR paralogs in C.

elegans begs the question of whether all heteromeric combinations of C. elegans TRPV channels will respond to

nicotinamide (Figure 4-5). Channels from the coexpression of OSM9 and OCR2 did not appear to be activated by

nicotinamide. It is possible that OSM9/OCR2, and possibly OSM9/OCR1 and OSM9/OCR3 as well, do not

possess nicotinamide sensitivity. The lack of nicotinamide sensitivity in these other channel combinations is

supported by the functional data as shown by my colleague Awani Upadhyay’s experiments (Upadhyay et al.,

2016). This observation, along with the phylogeny presented in Figure 4-5, suggest that nicotinamide sensitivity

was likely present in the ancestral invertebrate TRPV channel, but was lost in the expansion of the OCR genes.

The 2:2 stoichiometry of the OSM9/OCR4 channel suggests that incorporation of multiple OCR subunits within a

101 heterotetramer may be possible. However, expression of the various OCR subunits is cell-type specific (Tobin et

al., 2002); a TRPV heterotetramer with more than one OCR subunit would therefore not be physiologically

possible.

Figure 4-5. TRPV subunits from the cnidarian Nematostella vectensis cluster with invertebrate TRPV clade. To generate the phylogeny shown, full-length sequences for the listed genes were aligned using CLUSTALW as implemented by JALVIEW. This tree was constructed using the Neighbor-Joining method using the following parameters for computing evolutionary distance: Jones-Taylor Thornton matrix-based method with rate variation modeled using a gamma distribution (shape parameter = 2) and positions with less than 50% coverage being eliminated. Units for the evolutionary distance are in amino acid substitutions per site; overall tree length is 15.61085262. Interestingly, the genes CiTRPV_0100131491 and CiTRPV_0100148845 (highlighted in green) cluster with the OSM9-like and OCR-like lineage respectively. These genes are putative TRPV genes from the sea tunicate Ciona intestinalis, which is a basal chordate. The clustering of these pre-vertebrate sequences with invertebrate TRPV genes suggests that interdependent heteromerization could be a part of the ancestral TRPV phenotype. The genes NvTRPV_10538 and NvTRPV_1170467 (highlighted in yellow), which are from the starlet sea-anemone Nematostella vectensis, cluster with the OSM9/Inactive and OCR/Nanchung clades respectively. This supports the idea that interdependent heteromerization is the ancestral phenotype. This could also suggest that exclusivemechanosensitivity in TRPV channels is ancestral.

102 One other immediate observation from tree in Figure 4-5 is the segregation of CiTRPV_0100131491 and

CiTRPV_0100148845 into the OSM9-like and OCR-like clades. These two subunits are from Ciona intestinalis, which is a basal chordate (Dehal et al., 2002). This suggests that interdependent heteromerization of TRPV channels is the likely ancestral phenotype, and that homomerization among vertebrate TRPV subunits is likely a gain-of-function. Interestingly, two TRPV subunits from the cnidarian Nematostella vectensis, which are highlighted in yellow, segregate with the invertebrate TRPV lineage (Figure 4-5). Cnidarians are regarded as the sister group to bilaterians (Putnam et al., 2007). The fact that N vectensis TRPV genes segregate more closely with the invertebrate TRPV clade suggests that the invertebrate TRPV phenotype is the likely ancestral phenotype to all TRPV channels. This would support the idea that interdependent heteromerization is the ancestral TRPV phenotype given that the two N vectensis TRPV subunits segregate into the OSM9-like and OCR-like portions of the tree. This could imply that temperature sensitivity is another gain-of-function observed in vertebrates, and that

TRPV subunits originally only consisted of osmosensitivity or mechanosensitivity. Nicotinamide sensitivity could also be the ancestral phenotype, though it could also be the case that nicotinamide sensitivity evolved independently within the invertebrate TRPV clade. In order to test the ancestral TRPV phenotype, the cRNA that encodes the full-length subunits of the putative N vectensis TRPV channel could be injected into Xenopus oocytes, and current responses from exposure to various amounts of nicotinamide could be measured. If no nicotinamide-induced currents are observed, this would imply that nicotinamide sensitivity evolved after the cnidarian-bilaterian split and possibly independently within the invertebrate lineage.

The structural basis of interdependent heteromerization

Analysis of the gate residues of the various TRPV subunits may help determine the mechanism by which invertebrate TRPV channels are obligatory heteromers. Structures of vertebrate TRPV1, TRPV2, TRPV4 and

TRPV6 all demonstrate the presence of a common intracellular gate (Liao et al., 2013; Saotome et al., 2016;

Zubcevic et al., 2016; Deng et al., 2018). The key residues that make up the intracellular gate, as shown in Figure

103 4-6A, are I1 and M4. TRPV1, TRPV2 and TRPV6 have an additional extracellular gate that is composed of the

critical selectivity filter TI[G/I][M/L]GD. Although TRPV4 also has the same signature sequence in its selectivity

filter, it does not appear to possess an extracellular gate as the residues are simply too far apart (Deng et al.,

2018). As seen in Figure 4-6A, the intracellular gate is completely intact among the invertebrate and the cnidarian

TRPV subunits. All sequences contain the critical I1 and M4 residues in the appropriate position. A look at the selectivity filter of the non-vertebrate TRPV subunits demonstrates that this region is somewhat more divergent from the conserved selectivity filter of the vertebrate TRPV channels however (Figure 4-6B). In particular, the small glycine residue found in position 5 is often replaced by a bulky aromatic residue.

Figure 4-6. Disruption of the extracellular gate could be the mechanism of obligatory heteromerization among invertebrate TRPV channels. Analysis of the sequence alignment used to generate the phylogenetic tree in Figure 4-5 shows strong conservation of the distal S6 sequence among all lineages (A) but poor conservation of the selectivity filter among the invertebrate TRPV subunits (B). Key positions in the intracellular S6 gate (A) and the selectivity filter (B) are shown with asterisks and numbered 1-4 and 1-6 respectively.

The substitution of position 5 in the selectivity filter is reminiscent of the role of F6 in regulating the stoichiometry of Kv6.4 as presented in Chapter 3. The mechanism by which the aromatic residues in position 5 of invertebrate TRPV subunits prevents homomerization but allow heteromerization is not evident from the

104 sequence alignment alone. All of the “obligatory” heteromeric TRPV subunits in both the OSM9-like and OCR- like lineages shown in Figure 4-6 display the same type of substitution from a small glycine residue to an aromatic tyrosine or phenylalanine. It is possible that subtle differences in the structures of the individual OSM9- like and OCR-like subunits lead to incompatibility in the gate in the homomeric configuration but allow for stable heteromerization. Without structural data, one way to test if this position does lead to the interdependent heteromerization phenotype would be to generate a homology model of the OSM9/OCR4 heteromer using existing vertebrate TRPV crystal structures. It would also be interesting to see if placement of these aromatic residues into a vertebrate TRPV abolishes homomerization and channel function is only rescued when mutants containing either a tyrosine and a phenylalanine are coexpressed. The most likely candidate to perform this homology modeling and mutagenesis would be TRPV4. As mentioned above, TRPV1, TRPV2 and TRPV6 have an extracellular gate, but TRPV4 does not (Deng et al., 2018). It is more likely that the aromatic substitutions would be disruptive enough to affect gating, while still allowing proper folding of the subunits, in the TRPV4 background due to its unique structural differences.

105 References

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107 Chapter 5

Heme modulation of CNBD-containing Ion Channels

In this chapter, I will discuss a project that is yet to be published, which looks at the effect of heme modulation of the diverse Cylic Nucleotide Binding Domain (CNBD) containing family of channels. Despite the name, CNBD containing channels are non-uniformly modulated by cyclic nucleotides such as cAMP and cGMP; that is to say, cyclic nucleotides are required for channel opening of a subset of CNBD channels (the CNG family) and do not seem to influence the gating of other CNBD channels (the EAG family). Therefore, the discovery that heme appears to be an evolutionarily conserved modulator this family is particularly exciting. This project originated from a series of preliminary experiments performed by my advisor Tim Jegla. With the exception of the initial heme affinity curves for human Kv12.1 and N vectensis HCN1 shown in Figure 5-3, which were generated by my advisor Tim Jegla, I acquired all of the electrophysiology data presented here. The wild type plant channel constructs and all histidine mutants tested were cloned by the following students: Emma Baker,

Allan Lin, Fortunay Diatta, and Greg Busey.

Introduction

The Cyclic Nucleotide Binding Domain (CNBD) containing ion channel family is a large group of ion

channels within the 6-TM superfamily that can trace their lineage to prokaryotes. As mentioned in Chapter 1,

members of this channel family are responsible for critical physiologic functions including: cardiac pacemaking,

cardiac repolarization, regulating neuronal excitability, and signal transduction among metazoans, and transport of

nutrients necessary for metabolism among plants. From the discussion in Chapter 1, it is evident that CNBD-

containing channels are influenced by a broad array of molecules and stimuli aside from cyclic nucleotides.

Moreover, cyclic nucleotide sensitivity itself is not conserved across all CNBD-containing channels.

Within the core of the CNBD-containing family, there exists a highly conserved motif consisting of a

histidine and a cysteine residue separated by three variable residues (Figure 5-1A). This HxxxC motif appears to

108 point towards the VSD in all available CNBD-containing channel structures (Figure 5-1B, (Whicher and

MacKinnon, 2016; James et al., 2017; Lee and MacKinnon, 2017; Li et al., 2017; Wang and MacKinnon, 2017))

and may contribute to part of a known VSD-PD interface between the S5 and S1 domains (Lee et al., 2009).

Interestingly, this motif is not present in any Shaker family channels, so the motif is evidently not necessary for the function of all voltage-gated channels. There are presently no known functions for the HxxxC motif; however it is likely that these residues are important to gating due to their position in the above-mentioned VSD-PD interface.

Figure 5-1. The highly conserved HxxxC motif in CNBD-containing ion channels does not have any currently known functions. A) A consensus logo of the motif, found in the S5 transmembrane domain, made from an alignment of >50 sequences from the voltage-gated members of the EAG family, HCN family, and Plant CNBD- containing channel family. The histidine and cysteine residues that bookend the motif are universally, and near universally conserved in all observed sequences. B) The position of the histidine and cysteine residues (shown as stick models) in the recently published rat EAG1 structure (PDB: 5K7L). This motif consistently points towards the VSD, particularly the S1 transmembrane domain.

The HxxxC motif resembles the canonical c-type heme binding motif, which is CxxCH (Allen et al.,

2003). Because of this resemblance, we hypothesized that the HxxxC motif confers the ability to bind heme to

CNBD-containing channels. Heme is a prosthetic group that binds to many different proteins in order to facilitate biochemical reactions related to electron transfer, reduction-oxidation reactions, and diatomic gas binding

(Chapman et al., 1997). There are several different types of heme that differ with respect to the side-chains

109 attached to the main porphyrin ring (Chapman et al., 1997). The most commonly observed heme types are b-type

heme and c-type heme (Fufezan et al., 2008). The major difference between b-type heme and c-type heme is the substitution of vinyl groups in b-type heme to sulfhydryl groups in c-type heme (Figure 5-2); the sulfhydryl groups allow for covalent linkage of the heme moiety to the protein via cysteine residues (Bowman and Bren,

2008). Although b-type hemes are predominantly bind to b-type heme binding motifs, and c-type hemes interact with c-type hemoproteins, interaction between non-synonymous heme types and heme-binding motifs is possible through mutagenesis (Barker et al., 1995; Tomlinson and Ferguson, 2000).

Interestingly, b-type heme has been shown to interact with other members of the 6-TM family, namely the

Kv1.3 and BK channels. This interaction has been shown to occur through c-type heme binding motifs. The effect of heme on these channels is inhibition of inactivation of Kv1.3 channels and inhibition of activation of BK channels (Tang et al., 2003; Sahoo et al., 2013). It should be noted though that the heme-channel interactions occur via residues in the N-terminus of Kv1.3 and the C-terminus of BK (Tang et al., 2003; Sahoo et al., 2013); therefore, the heme-channel interaction takes place intracellularly in these channels. The HxxxC motif in CNBD- containing channels is accessible from the extracellular environment. For this reason, the effect of heme was tested on CNBD-containing channel gating using Two-Electrode Voltage Clamp and bath application of heme.

Because b-type heme was shown to interact with a c-type motif in the channels mentioned above, b-type heme was also used in these experiments.

Figure 5-2. Heme types are determined by differences in functional groups attached to the central porphyrin ring. The two major heme types found in nature, b-type heme and c-type heme, are distinguished by the presence of

110 sulfhydryl groups (red circles) in c-type heme. These sulfhydryl groups allow covalent linkage of the heme to its interacting protein. The types of reactions that can theoretically be facilitated by these two heme types is largely the same. Black lines represent covalent bonds; unless specified, all nodes between bonds represent carbon atoms. All heteroatoms listed are abbreviated as they would be in the Periodic Table of Elements.

Surprisingly, hemin, which is a b-type heme conjugated to a chloride ion, modulated the activity of two distinct CNBD-containing ion channels: human Elk1 (HsElk1) and Nematostella vectensis HCN1 (NvHCN1).

This modulation of activity occured in a concentration dependent manner (Figure 5-3). Hemin inhibited NvHCN1 channels (Figure 5-3A,C-D) and potentiated the activity of HsElk1 channels (Figure 5-3B,E-F) in a voltage- dependent manner. This suggested that in both cases hemin stabilized the outward conformation of the voltage sensor domain. The estimated dissociation constant (KD), which is a measurement of the affinity, was in the micromolar range for HsElk1 (Figure 5-3F) and in the hundreds of nanomolar for NvHCN1 (Figure 5-3D). It should be noted that these measurements of the dissociation constant could represent an overestimate as the true concentration of heme in the plane of the membrane could be much lower due to poor solubility.

111

Figure 5-3. Heme is a potent inhibitor of Nematostella vectensis HCN1 channels and an activator of human Elk1 channels. Example traces from a Xenopus oocyte expressing (A) NvHCN1 channels and (B) HsElk1 channels prior to (Control) and after the application of 10 µM hemin. NvHCN1 channels appear to be inhibited by hemin, whereas HsElk1 channels are potentiated. C) The G-V curve of NvHCN1 channels left shifts upon application of hemin in a concentration dependent manner. D) Using the shift in the V50 as an estimate for affinity, a Hill equation is fit demonstrating sub-micromolar affinity for NvHCN1. E) Like NvHCN1, the left-shift among HsElk1 channels is also concentration dependent. F) Fitting a Hill equation to the concentration dependent change in V50 demonstrates an affinity in the single micromolar range.

Based on these preliminary findings, it was evident that heme appeared to have a significant role in modulating the function of CNBD-containing channels from two distinct metazoan lineages. The mechanism by which this modulation occurs was appears to suggest stabilization of the VSD-out conformation of the channels,

112 owing to the fact that depolarization-gated channels are potentiated and hyperpolarization-gated channels were

inhibited. An alternative mechanism by which this could occur is through direct interaction with the pore,

however this is not as likely as there are no changes in current size in HsElk1 after hemin exposure. Because of

this large shift in channel function, I set out to determine if heme-channel interactions are conserved in CNBD-

containing family, and to determine the mechanism by which heme exerts its effects. Specifically, I wanted to

know: if heme is causing the channels to enter into a metastable state; if divalent cations can disrupt the heme

effect; if heme has an effect on Plant CNBD-containing channels; and finally, to test if heme is exerting its effects

through the highly conserved HxxxC motif.

Methods

Molecular Cloning

KAT1, SKOR and GORK (courtesy of Dr. Sarah Assmann) were cloned into pOX vector using the

InFusion cloning kit (Clonetech) per manufacturer’s instructions. Channel cDNA was inserted within the HindIII

and XbaI cutsite, and the primers used for cloning are as follows. For KAT1:

5’-CGACATCGATAAGCTTCCACCATGTCGATCTCTTGGACTCGAA-3’ and

5’-AGCAGAAACTTCTAGATTAATTTGATGAAAAATACAAATGATACC-3’. For SKOR:

5’-CGACATCGATAAGCTTCCACCATGGGAGGTAGTAGCGGCGGC-3’ and

5’-AGCAGAAACTTCTAGATTATGTTTCAACAGCCAAATACAGTT-3’. For GORK:

5’-CGACATCGATAAGCTTCCACCATGGGACGTCTCCGGAGACGG-3’ and

5’-AGCAGAAACTTCTAGATTATGTTTGATCAGTAGTATCACTGA-3’. HsElk1 H368Q and GORK C151A mutants were cloned using overlap PCR. The primers used for the H368Q mutant are as follows:

5’-TGCAGAAGCTCAGAATAAACGCTC-3’ and

5’-CCAGATACACGCCATCCACTGTGCAAGGAGTGCAAA-3’ (Piece 1), and

113 5’-TTTGCACTCCTTGCACAGTGGATGGCGTGTATCTGG-3’ and

5’-ATGTAGCTTAGAGACTCCATTCGG-3’ (Piece 2). The primers used for the C151A mutant are as follows:

5’-TGCAGAAGCTCAGAATAAACGCTC-3’ and

5’-ATAAATAAGATCCCAAGGGAAGGCACCGATGAAATCCATGAG-3’ (Piece 1), and

5’-CTCATGGATTTCATCGGTGCCTTCCCTTGGGATCTTATTTAT-3’ and

5’-ATGTAGCTTAGAGACTCCATTCGG-3’ (Piece 2). The restriction enzymes used to place the mutagenized fragments into the plasmid were EcoRI and MluI for H368Q, and NheI and EcoRI for C151A. Oligonucleotide primers were obtained from IDT (Integrated DNA Technologies, IL). All mutant constructs were confirmed by sequencing.

cRNA synthesis

All cRNAs used in the experiments described were created using the T3 mMessage mMachine kit, per manufacturers’ instructions (Life Technologies). cRNAs were stored at -80°C until the time of injection. A 1:20 mix of the RNAse inhibitor SUPERase-In (Invitrogen) and nuclease-free water was added to cRNA dilutions to prevent degradation during injections.

Electrophysiology

See Chapter 2 for a discussion on how electrodes were prepared and how oocytes were clamped.

Solutions used in this chapter consisted of the following. HsElk1 recordings: 96 mM NaOH, 2 mM NaCl, 2 mM

KCl, 1 mM CaCl2, 5 mM HEPES, pH 7.5 with methanesulfonate. GORK and SKOR recordings: 96 mM NaOH, 2 mM NaCl, 2 mM KCl, 1 mM CaCl2 5 mM HEPES, pH 7 with methanesulfonate. KAT1 recordings: 96 mM

NaOH, 2 mM NaCl, 5 mM KCl, 1 mM CaCl2 5 mM HEPES, pH 7 with methanesulfonate. Hemin and

Protoporphyrin IX were purchased from Sigma-Aldrich. Zinc Protoporphyrin IX and Cobalt Protoporphyrin IX

114 were purchased from Frontier Scientific. For all heme-related compounds 10 mM stock solutions were prepared

fresh before recording by dissolving in either 30 mM NaOH solution or DMSO.

Sequence Alignments

Sequences were aligned via the CLUSTALW through MEGA7 (Kumar et al., 2016). Sequence logo

displayed in Figure 5-1A was generated using WebLogo (Crooks et al., 2004).

Results

The heme effect in human Elk1 channels is distinct from mode-shift

Outward stabilization of the voltage sensor among metazoan CNBD-containing channels is known to

occur through other known mechanisms, namely the phenomenon known as “mode-shift” (Figure 5-4A). Mode- shift, which is also referred to as voltage dependent potentiation (VDP), is a phenomenon that occurs in many voltage-gated channels when the cell membrane is depolarized for relatively long periods of time (Villalba-Galea,

2017). These prolonged depolarizations allow the voltage sensor domain to enter into a relaxed metastable state while it is still in the outward conformation (Villalba-Galea et al., 2008). Because mode-shifted VSDs are in a metastable state, hyperpolarziation of the membrane is required to return the VSDs into the downward position.

Both EAG family and HCN family channels are known to exhibit mode-shift (Mannikko et al., 2005; Gianulis et al., 2013). In order to test if heme is simply leading to the voltage sensor to enter into a mode-shifted state, I tested to see if heme can further left-shift already mode-shifted channels. Mode-shift was induced by holding oocytes at +60 mV for 2 seconds; this was followed by increasingly hyperpolarizing pulses and a measurement of the tail current at -40 mV (Figure 5-4B). Following the addition of heme, there was a left shift in the mode-shifted

GV of approximately 33 mV (Figure 5-4C), which is somewhat higher compared to the heme-induced left shift in

115 the non-mode shifted GV (ΔV50 of approximately 19 mV, p < 0.05 t-test). These results suggest that the heme

effect and mode-shift are independent processes and are possibly synergistic.

Figure 5-4. Heme and mode-shift are additive processes on HsElk1. A) Mode-shift of VSDs, particularly in CNBD-containing channels, occurs when the membrane is strongly depolarized for extended periods of time. Mode-shifted VSDs require hyperpolarizing pulses in order to return to VSD-in state. B) Mode-shift of human Elk1 channels was induced by holding oocytes to +60 mV for 2 seconds; hyperpolarizing pulses are required to close the mode-shifted channels and isochronal tail currents at -40 mV are used to generate the mode-shifted GV. C) Heme confers an additional left-shift on the mode-shifted GV that is similar in magnitude to the non-mode- shifted GV. V50 of the various GVs are as follows: control, no mode-shift -61.3 ± 0.8 mV; hemin, no mode-shift - 80 ± 4 mV; control, with mode-shift -125.3 ± 3.8 mV; hemin, with mode-shift -158.2 ± 1.3 mV (n = 4 for all experiments).

The left-shift observed in human Elk1 channels is dependent on iron-containing heme moieties and is reversed in the presence of divalent cations

Another potential mechanism by which heme could be exerting its effects on CNBD-containing channels is by creating tension in the lipid membrane through intercalation. Membrane tension itself has been shown to activate Shaker family channels (Schmidt and MacKinnon, 2008; Schmidt et al., 2012). Cobalt Protoporphyrin IX

(Co-PPIX), a type of heme in which the central iron atom is replaced with a larger cobalt atom, did not

116 demonstrate any changes in the GV for HsElk1 (Figure 5-5A). Flowing hemin after Co-PPIX left-shifted the GV

by approximately 10 mV (Figure 5-5B). Hemin induced left-shift without Co-PPIX in the same recording session

was greater than 25 mV (Figure 5-5B). The reduction in the hemin induced left-shift after Co-PPIX exposure

coupled with the lack of a left-shift with Co-PPIX suggests that metal-substituted hemes could occupy the same site as iron-containing hemin, but do not induce the same type of effect. This suggests that membrane stretch is not the mechanism by which heme is exerting its effects on CNBD-containing channels.

Because the character of the metal within the porphyrin ring itself can alter the heme effect, I wanted to test whether other heavy metals can also influence the heme effect. As mentioned in Chapter 1, divalent cations, such as zinc and cadmium, are known to be potent inhibitors of EAG family channels by directly inhibiting voltage sensor movement (Silverman et al., 2000; Zhang et al., 2009). Surprisingly, when oocytes expressing wild-type HsElk1 channels were exposed to heme in the presence of 100 µM zinc, the left shift observed in

Figures 5-3 and 5-4 is not seen. Instead, the inhibition of the channels by zinc appears to be potentiated (Figure 5-

5C). Increasing the concentration of the zinc in the bath media to 1 mM reduces the amount of maximal right-shift observed compared to the heme-free zinc control (Figure 5-5D). These results suggest that heme could be potentiating the effect of divalent cations by increasing the affinity of the channel-divalent cation interaction

(Goodman et al., 2011).

117

Figure 5-5. Iron-containing heme is required for the observed left-shift and divalent cations reverse the heme effect. A) Co-PPIX does not appear to have any effect in the gating of HsElk1 channels by itself. B) Flowing heme after Co-PPIX exposure demonstrates a reduction in the left-shift compared to heme alone (V50 of -57.2 ± 2.8 mV for control, n = 4; V50 of -83.6 ± 3.9 mV, n = 3 with heme alone; and -67.1 ± 1.7 mV with heme after Co- PPIX, n = 4). C) Exposure to heme potentiates right-shift in the presence of 100 µM zinc. Oocytes were iteratively depolarized for 1 second from -100 mV and isochronal tail currents were collected at -40 mV. Red traces show currents elicited by +30 mV pulse. D) Hill plot of heme induced shift of human Elk1 in the presence of 100 µM Zinc (black, n = 4 eggs per hemin concentration) or 1 mM Zinc (red, n = 3 eggs per hemin concentration). Heme accentuates the right-shift of zinc, with a maximal increase in right-shift greater in the presence of 100 µM Zinc (V50 in 100 µM zinc alone: -23.0 ± 1.7 mV; V50 in 100 µM zinc and 30 µM hemin: -4.7 ± 2.4 mV) compared to 1 mM Zinc (V50 in 1 mM zinc alone: 5.0 ± 1.6 mV; V50 in 1 mM zinc and 30 µM hemin: 12.3 ± 1.7 mV).

Hemin leads to stabilization of the VSD-out state of Plant CNBD-containing channels

Hemin was tested on the outward rectifying Plant CNBD-containing channels SKOR and GORK and the inward rectifying Plant CNBD-containing channel KAT1. Hemin greatly increased the current size of SKOR

(Figure 5-6A,B) and GORK (Figure 5-6C,D), and moderately reduced the current size for KAT1 (Figure 5-6E,F).

These results are consistent with the model of VSD-out stabilization that appears to be present in the metazoan channels in Figure 5-3. Because Plant CNBD-containing channels are a separate evolutionary lineage, this suggests that heme modulation of CNBD-containing channels is evolutionarily conserved.

118

Figure 5-6. Outward voltage sensor stabilization by heme is conserved in Plant CNBD-containing channels. Example traces of the Plant CNBD-containing channels (A) SKOR (C) GORK and (E) KAT1 before (Control) and after the application of 10 µM hemin. Hemin increases the current size by (B) 2.3-fold in SKOR at +70 mV (n = 5) and (D) 5-fold in GORK (n = 5). F) Hemin decreases current size to 68% of the peak inward current at - 150 mV (n = 3) for KAT1.

The conserved S5 histidine is necessary, but not sufficient, for the heme effect in EAG family

The only functional mutant of the histidine residue in the HxxxC motif thus far identified is a histidine to glutamine (H->Q) substitution. The homologous mutation in the human ERG1 channel appeared functional but does not contribute to pH sensitivity of the channel (Van Slyke et al., 2012). When the histidine to glutamine mutation is made in human Elk1, the resulting mutant channels traffic to the membrane and appear functional.

119 These channels are right-shifted compared to wild-type HsElk1 channels, having an apparent V50 of approximately -19 mV (Figure 5-7A). When 10 µM hemin is flowed onto H368Q mutants, the channels are left- shifted with a V50 of approximately -38 mV (Figure 5-7A,B). This 19 mV left-shift is two-thirds the size of the left-shift that was observed in wild-type HsElk1 channels at the same hemin concentration (Figure 5-3B,E-F).

Unfortunately, the histidine to glutamine substitution is not functional among any of the plant channels.

However, there are clues of other residues that may be important to the heme-protein interaction among the Plant

CNBD-containing channels. GORK contains a cysteine residue found in the S3 region; this residue, C151 in

GORK, was previously implicated in conferring redox sensitivity in SKOR (Garcia-Mata et al., 2010). When 10

µM hemin was flowed onto GORK C151A mutant, there was a roughly 3-fold increase in current size (Figure 5-

7C,D). Wild-type GORK channels see an increase in current size of roughly 5-fold, so there is a 40% reduction in hemin efficacy in the C151A mutant background.

Figure 5-7. The heme effect can be reduced by mutating key residues in the pore domain of HsElk1 channels and the voltage sensor of GORK channels. A) HsElk1 H368Q channels are functional with greatly right-shifted gating when compared to wild-type channels. B) The heme effect appears to be mitigated in HsElk1 H368Q channels. The V50 of the channels in the control solution is -19.1 ± 1.2 mV; this shift increases to -38.6 ± 5.1 mV when the channels are exposed to 10 µM hemin (n = 4). C) An example trace of GORK C151A expressing oocyte prior to (Control) and after the application of heme. Heme has an effect on C151A mutant channels, (D) however this

120 effect is only a 3-fold increase (n = 3) as compared to wild-type GORK channels, which show closer to a 5-fold increase.

Discussion

The data presented demonstrate that the heme effect is likely not the result of the mode-shift phenomenon

(Figure 5-4B, C). Among EAG family channels at least, interactions between the eag domain and the CNBHD have been shown to be critical for mode-shift to occur (Gianulis et al., 2013; Dai and Zagotta, 2017; Zhao et al.,

2017; Dai et al., 2018). Interestingly, preventing channel-PIP2 interaction also appears to be important for mode-

shift (Li et al., 2015). It would therefore be interesting to see if heme can still exert its effects in mutant EAG

channels that have disrupted mode-shift or when PIP2 levels are depleted.

The heme effect in EAG channels has also been shown to require iron-containing heme. CoPPIX may be

able to interact with the heme binding site but does not elicit the same effects (Figure 5-5A,B). The differing electronic structure of the central cobalt atom leads to differences in the ability of CoPPIX to participate in similar physiological functions; notably CoPPIX is not capable of binding diatomic gases to the same degree as FePPIX

(Yonetani et al., 1972; Blough et al., 1980; Marden et al., 2000). This suggests that the physiological role of heme binding, at least to EAG channels, could be related to gas sensitivity. It would be interesting to see if CoPPIX has the same inhibitory effects in HCN and the Plant CNBD-containing channels.

At least within the EAG lineage, binding of divalent cations appears to be potentiated by the heme- channel interaction (Figure 5-5C,D). Although divalent cations are not known to interact with HCN and Plant

CNBD-containing channels through the same voltage sensor-based mechanism as EAG channels, the residues within the VSD that appear to mediate divalent cation sensitivity are also present within the HCN and Plant

CNBD-containing channel lineage (Zhang et al., 2009). It would therefore be interesting to see if modulation of divalent cation binding is potentiated in the presence of heme in these other channels. Additionally, it would be interesting to see if the residues that mediate divalent cation sensitivity are important for the left-shift observed when heme alone is present.

121 Heme has been shown to interact with Plant CNBD-containing channels (Figure 5-6). These channels represent a divergent evolutionary lineage compared to the animal CNBD-containing channels shown in Figures

5-3 through 5-5, suggesting that the heme-CNBD channel interaction could be evolutionarily conserved. A key difference between the phenotype of the heme effect between metazoan and Plant CNBD-containing channels is that heme radically affects current size of the Plant CNBD-containing channels, whereas heme does not have significant effects on EAG current size. Upon review of Figure 5-3A, heme may have an effect on NvHCN1 current size; this effect on NvHCN1 current size will need to be further examined in light of the observed heme effect in Plant CNBD-containing channels. The most parsimonious explanation for the results shown in Figures 5-

3 and 5-6 is that heme exposure leads to an evolutionarily conserved stabilization of the VSD-out conformation of

CNBD-containing channels. It is possible that heme exerts its effects by manipulating the pore domain directly, and either facilitating or blocking CNBD-containing channels depending on if it is depolarization-gated or hyperpolarization-gated. This is unlikely, however, given the aforementioned lack of current size change in EAG channels following heme exposure.

Stabilization of the VSD-out conformation can happen in one of two ways: heme could directly facilitate the movement of the VSDs, or heme can couple the outward movement of VSD movement more tightly to the pore domain (Figure 5-8). It is not clear from the data which of these two possibilities is correct. HCN and Plant

CNBD-containing channels notably have weaker VSD-PD coupling compared to EAG channels (Zei and Aldrich,

1998; Latorre et al., 2003; Ryu and Yellen, 2012). If the coupling between the VSD and pore were to strengthen, this could explain the measured changes in current size following heme exposure in these two families of channels. In the case of EAG channels, which already demonstrate reasonably strong coupling between voltage sensor and pore, strengthening of VSD-PD coupling could manifest as the hyperpolarized activation that is observed. This would favor the voltage-independent model shown in Figure 5-7, however more direct measurements of VSD movement are need to test the two possibilities.

122

Figure 5-8. Heme-stabilization of the VSD-out conformation in depolarization and hyperpolarization gated channels. Heme can facilitate the VSD-out conformation by affecting voltage sensor movement directly, which is a voltage-dependent transition, or by affecting VSD-PD coupling, which is a voltage-independent transition.

The heme effect appears to require the conserved S5 histidine residue at least among animal depolarization-gated channels (Figure 5-7B). It should be noted that the effect is still present to some degree in the human Elk1 H368Q mutant. The significance of this lingering heme effect is not clear, but it may suggest that other residues also play a role in the heme effect. Based on the allosteric regulation of the heme effect, it would be interesting to see if mutants of any of the known residues that mediate divalent cation binding have alterations in the heme effect. Preliminary results, shown in Chapter 6, suggest that one such residue, a negatively charged aspartate residue in S2, may have some diminished ability to interact with heme. However, these results would need to be replicated before any definitive conclusion can be drawn.

123 As mentioned above, the H->Q mutation is not functional in the Plant CNBD-containing channels tested.

This precludes the ability to directly test if the conserved heme-channel interaction necessarily requires the conserved HxxxC motif in all CNBD-containing channels. Interestingly, other residues appear to be important in conferring heme sensitivity among Plant CNBD-containing channels; C151 in GORK has a reduction in the heme effect size compared to wild-type GORK (Figure 5-6C,D and 5-7C,D). This residue is unique to the outward rectifying Plant CNBD-containing channels and has been implicated in modulating redox sensitivity in SKOR

(Garcia-Mata et al., 2010). C151 is not consistently found in any of the metazoan CNBD-containing channels.

This suggests that the heme-channel interaction, while potentially requiring the same key S5 motif, may also require other residues that are lineage dependent. The corollary to this is that heme may exert effects among channels in the metazoan lineage that are unique to metazoan channels. One potential difference between metazoan and plant channels in their heme interactions, the ability to bind diatomic gases, is explored in Chapter

6.

124 References

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126

Chapter 6

Summary, Biological Applications of Findings, and Future Directions

In this chapter, I will give a summary of the results from Chapters 3, 4 and 5. I will then discuss the biological significance of these results as well as potential applications of these findings. Finally, I will speculate on the direction of the research areas presented, with an emphasis for the project presented in Chapter 5, as this is unpublished work.

Summary of Experiments

The development and testing of a novel model for obligatory heteromeric stoichiometry formation

The data presented in Chapter 3 demonstrate that the activation gate itself is necessary to fix the

stoichiometry of an obligatory heteromeric subunit to one subunit in the functional heteromer. The disrupted gate

in Kv6.4 and related subunits likely arose after the loss of T1 self-association. Subsequent mutations in the gate then fix the stoichiometry of the Kv2.1-Kv6.4 heteromer to allow cells greater control over channel assembly.

This gate regulation model may be supported by the data shown in Chapter 4: the stoichiometry of the invertebrate TRPV channel formed by OSM9 and OCR4 is predominantly 2:2. Furthermore, analysis of the gate interfaces shows broad compatibility with homomer forming TRPV subunits in the vertebrate lineage with the exception of one key residue whose significance remains to be seen.

127 Identification of a novel invertebrate TRPV ligand and exploration of invertebrate TRPV assembly

My experiments showed that the hypothesis that nicotinamide is a ligand for invertebrate TRPV channels is correct, and that this ligand activity is evolutionarily conserved. Previous attempts to express invertebrate

TRPV channels heterologously were unsuccessful. With a newly identified ligand, it becomes feasible to study structure-function relationships of invertebrate TRPV channels heterologously. With the ability to heterolgously express invertebrate TRPV channels, it was identified using TIRF microscopty that the OSM9/OCR4 channel has

2:2 stoichiometry. This fundamental property of the channels had been speculated, but never previously been proven.

Characterization of a conserved heme-channel interaction within the CNBD-containing channel family

It was discovered by my advisor, Tim Jegla, that heme appears to potently stabilize the voltage sensor domains of two different CNBD-containing channels from humans and Nematostella vectensis in the outward position. My experiments demonstrate that this effect is independent of voltage dependent potentiation, also known as mode-shift. Additionally, heme appears to potentiate divalent cation binding, suggesting there may be some type of allosteric interaction between the putative heme binding site and the known divalent cation binding site. Moreover, the outward stabilization of the VSD with heme exposure appears to be conserved in Plant

CNBD-containing channels. Finally, it was shown that the heme interaction depends, at least partially, on a universally conserved histidine within the S5 domain, although the nature of the heme-protein interaction may also exhibit lineage-specific interactions.

128 Biological Significance and Potential Applications of Findings

Restricted heteromeric stoichiometry prevents regulatory subunit mixing

The data in Chapter 3 show that the functional stoichiometry of the Kv2.1/Kv6.4 heteromer is three

subunits of Kv2.1 to one subunit of Kv6.4. This is significant, because Kv6.4 is one of several regulatory subunits

in the mammalian lineage (Bocksteins, 2016), many of which display similar divergence in the gate sequence. It

is therefore likely that the 3:1 stoichiometry is conserved among the mammalian obligatory heteromers.

Regulatory subunits are expressed in many different tissue types; additionally, there appears to be overlap in the

tissue expression profile, suggesting that individual cells could express multiple regulatory subunits

simultaneously. The implication of a stoichiometry limited to one regulatory subunit per heteromer is that

different regulatory subunits likely cannot be present in the same heteromer in vivo, even if a cell expresses two or

more regulatory subunits.

One potential application for this knowledge would be the development of targeted therapeutics that can

specifically affect these obligatory heteromers without influencing the behavior of other channels. Dysfunction of

regulatory subunits has been implicated in diseases such as migraine and certain types of hereditary blindness

(Bocksteins, 2016), so targeting of these channels is clinically relevant. In order to modulate a specific

heteromeric current in a diseased cell, one would only have to design a drug that specifically targets the regulatory

subunit within that heteromer. This method of drug design has the added benefit that effects on vital organs, such

as the heart, can be controlled through the choice of subunit.

Invertebrate TRPV channels as an in vivo nicotinamide biosensor

The discovery that invertebrate TRPV channels are activated by nicotinamide contributes to a growing

body of evidence linking vitamin B3 metabolism to TRP channel activity. Nicotinic acid has been shown to activate vertebrate TRPV channels (Ma et al., 2015). Although not in the TRPV lineage, TRPM2 channels can be

129 activated either directly by NAD or through ADP-ribosylation (Perraud et al., 2001; Sano et al., 2001). This suggests that not only do TRP channels serve sensory functions but their activity can also be modulated by the metabolic state of the cell.

One potential application of the nicotinamide-TRPV relation is in vivo measurement of nicotinamide production. As mentioned in Chapter 4, nicotinamide is produced during repair of genotoxic damage; it could therefore be very useful to measure fluctuations in nicotinamide levels during various pathological states. Because invertebrate TRPV channels are calcium-permeable (Nesterov et al., 2015; Upadhyay et al., 2016), they can be used in a system with genetically encoded calcium sensors such as GCaMP (Nakai et al., 2001). In this system, upregulation of nicotinamide would lead to an increase in calcium-induced fluorescence.

The interaction between heme and CNBD-containing channels

Heme has previously been shown to interact with non-CNBD-containing 6-TM ion channels (Tang et al.,

2003; Sahoo et al., 2013). However, the relation between heme and CNBD-containing channel gating characterized in Chapter 5 is novel due to the following reasons. First, it is likely that the heme-CNBD-containing channel interaction is occurring extracellularly due to the fact that heme is being applied extracellularly and is not membrane permeable. This is in contrast to previous investigations of heme-channel interactions that primarily utilize intracellular application of heme (Tang et al., 2003; Sahoo et al., 2013; Burton et al., 2016). Second, as a consequence of the extracellular binding site, heme likely has a more direct interaction with the voltage sensing or gate coupling mechanisms of CNBD-containing channels. Whereas in the heme-channel interactions studied previously, the cytoplasmic domains implicated in heme-binding are not as directly linked to voltage gating.

Finally, the heme-channel interaction has been shown to be conserved in metazoan and plant CNBD-containing channels. This broad conservation of the heme effect suggests that heme-binding was present in the ancestral

CNBD-containing channel. The conservation of heme modulation in the non-CNBD-containing channels mentioned above is unclear.

130 One potential application of the heme-CNBD channel interaction could be the development of drought- resistant crops. Water is a necessary nutrient in plant growth, and it is accepted that roughly 65% of fresh water is used just for agricultural purposes (Postel et al., 1996). Drought-like conditions can severely hamper crop growth through water loss (Schroeder et al., 2001; Korner, 2015). It is therefore desirable to engineer crops to become drought-resistant, particularly given that these types of conditions may become more common through Global

Warming (Seager et al., 2007). The primary mechanism used by plants to regulate evaporative losses of water is through the opening and closing of stoma through guard cells. Guard cells rely on potassium efflux and influx to determine turgour pressure, which in turn controls the cross-sectional area of the stoma (Schroeder et al., 2001).

Figure 6-1 shows the likely outcome of heme application on plant stomatal size, which is that heme exposure would likely lead to stomatal closure. This would theoretically reduce evaporative water loss and help plants conserve water.

Figure 6-1. Predicted effects of heme exposure on plant guard cells. Guard cells are specialized cells used by plants to regulate gas exchange and evaporation. When guard cells are turgid, the stoma remains open; this facilitates gas exchange but also leads to water loss through evaporation. When guard cells are flaccid, gas

131 exchange is restricted as is water loss. Guard cells utilize GORK channels (shown as gray boxes), KAT1 channels (shown as yellow boxes) and aquaporins (shown as blue boxes) to regulate turgour. The net flow of potassium through GORK and KAT1 channels dictates the direction of water flow through the aquaporins, which in turn dictates turgour pressure. Based on the results presented in Chapter 5, heme exposure would lead to increased GORK current and reduced KAT1 current. This would yield a reduction in turgour pressure through water efflux and result in stomatal closure.

Future Directions

Testing the gate regulation model in other 6-TM channels

With regards to the gate-regulation model presented in Chapter 3, it will be interesting to see if other

known regulatory subunits follow the stoichiometry rules as predicted. The are several independent expansions of

the regulatory phenotype found in virtually all clades of animals. Indeed, a number of regulatory subunits from

cnidarian Shal, and Shaw as well as ctenophore Shaker regulatory subunits were identified through previous work in our own lab (Li et al., 2015). The stoichiometry of these channels could also be determined, possibly through the use of TIRF microscopy as was done in Chapter 3. This stoichiometry could then be correlated to the divergence in the gate sequence. Regulatory subunits that do not display significant divergence in the gate sequence, such as the mammalian Kv5 family, would be predicted to have greater rates of 2:2 heteromer formation compared to regulatory subunits that are more highly divergent from homomer forming subunits.

The gate regulation model could be tested on the invertebrate TRPV channel formed by OSM9/OCR4 to determine if interdependent heteromerization is the result of perturbations in the selectivity filter. As shown in

Chapter 4, the OSM9/OCR4 channel has a 2:2 stoichiometry. Additionally, there are few significant differences between the selectivity filter of the homomer-forming vertebrate TRPV lineage and the obligatory heteromer- forming invertebrate TRPV lineage. The selectivity filter forms an extracellular gate in vertebrate TRPV1,

TRPV2 and TRPV6 channels. One potential way to test the gate regulation model using the invertebrate TRPV lineage would be to place the pore domain of OSM9 or OCR4 into a vertebrate TRPV channel background and test if functional homomerization is abolished when the chimeric constructs are expressed by themselves.

132 Functional heteromerization should only be present when vertebrate TRPV channels containing the OSM9 pore

are coexpressed with vertebrate TRPV channels containing the OCR4 pore.

Investigating the origins of nicotinamide sensitivity in TRPV channels

The data presented in Chapter 4 demonstrate that nicotinamide has an evolutionarily conserved TRPV

channel activation property among invertebrates. Initial tests on vertebrate channels showed it likely does not

share the same role, although interestingly enough, another vitamin B3 metabolite, nicotinic acid, has been shown

to activate vertebrate TRPV4 channels. A phylogenetic analysis of TRPV channels shows cnidarian TRPV

subunits clustering with the invertebrate clade (Figure 4-5). This suggests that cnidarian TRPV channels could

also be activated by nicotinamide, however there is not enough resolution in the analysis to prove this. In order to

test if nicotinamide sensitivity predates the cnidarian-bilaterian split, experiments should be designed in a manner similar to what was demonstrated in Chapter 4: NvTRPV_10538 and NvTRPV_1170467 should be expressed individually, or together, in Xenopus oocytes and exposed to varying concentrations of nicotinamide. If currents are produced with nicotinamide exposure, then nicotinamide sensitivity in TRPV channels is the ancestral phenotype. It should be noted though, that the absence of nicotinamide-induced currents does not necessarily mean that nicotinamide-sensitivity evolved independently in the invertebrate clade. Other possibilities that would need to be ruled out would be the cnidarian TRPV subunits are either not properly translated, assembled or trafficked to the membrane. This alternative hypothesis is less likely if the full-length coding sequences of the

TRPV subunits are used, since the invertebrate TRPV subunits did not require any chaperones or cofactors to express into functional channels in Xenopus.

133 Determining the mechanism and potential roles of the heme-CNBD interaction

As mentioned in the Discussion section in Chapter 5, it is unclear from the data whether heme is directly facilitating outward movement of the VSDs or exerts its effects by manipulating VSD-PD coupling. In order to distinguish between these two possibilities, more direct experiments are needed. One potential way to answer this question is to use the sophisticated technique known as Voltage Clamp Fluorometry, or VCF (Figure 6-2A). In

VCF, a mobile cysteine residue in the voltage sensor domain of an ion channel that is accessible from the extracellular environment is labelled using a thiol-reactive fluorescent dye. Movement of the fluorescently labelled residue across the plane of the membrane can be measured using a photodiode or photomultiplier tube

(Gandhi and Olcese, 2008). When used in combination with standard Two Electrode Voltage Clamp, the measured fluorescent changes can be used to generate a Fluorescence-Voltage (FV) relationship; the FV relationship is effectively an assay of voltage sensor movement (Gandhi and Olcese, 2008). One can then predict that if heme induces its effects on CNBD-containing channels by manipulating voltage sensor movement directly, the changes in GV relationship in the presence of heme would also be present in the FV relationship (Figure 6-

2B). However, if heme influences VSD-PD coupling, there likely would not be as much of left-shift in outward

VSD movement in the FV relationship (Figure 6-2C). There may be changes in the rate of voltage sensor return following depolarization in the latter case, but the ability to measure these changes would depend on the speed of the clamp.

134

Figure 6-2. Voltage Clamp Fluorometry could be used to identify the mechanism of channel modulation by heme. A) Voltage Clamp Fluorometry (VCF) works by placing a fluorescent label (green circles) on a part of the voltage sensor domain that can move across the membrane during changes in transmembrane potential. When the VSDs are in the down state, the fluorescent signal is at a minimum. As the membrane depolarizes and the VSDs move extracellularly, and the fluorescent label produces a signal that can be measured by photosensitive equipment. Measuring the changes in fluorescent signal over a range of voltages can be used to generate a Fluorescent- Voltage (FV) curve B) If heme affects voltage sensor movement directly, the measured heme left shift in the GV will be associated with a one-to-one left shift in the FV. C) If heme affects VSD-PD coupling, the left shift in GV with heme exposure will not be associated with an equivalent left shift in the FV. In (B) and (C), Solid lines represent GV, dashed lines represent FV. Black lines depict the control solution and red lines depict the heme shifted solution.

The role of heme-binding to the physiology of these channels must also be explored. Heme is known to have diverse roles as a sensor of soluble diatomic gases such as O2, nitric oxide (NO) and carbon monoxide (CO).

This gas binding allows heme-protein complexes to partake in sophisticated signaling pathways (Shimizu et al.,

2015). It would be particularly interesting to test if heme binding confers NO sensitivity to CNBD-containing channels, as NO has diverse signaling roles in both animals and plants. Among animals, NO is highly active in the

135 nervous system, where it can act as a neurotransmitter produced both pre- and post-synaptically (Garthwaite,

2008). NO is also implicated in regulation of smooth muscle muscle activity (Feletou et al., 2012; Farrugia and

Szurszewski, 2014). Among plants, NO appears to play an important role in growth signaling and reproduction

(Domingos et al., 2015).

Preliminary results suggest NO given off by the small molecule S-Nitroso-N-acetyl-DL-penicillamine

(SNAP, dissolved in DMSO) can completely reverse the heme-induced left shift in human Elk1 channels (Figure

6-3A, B). More experiments are needed to determine if heme is still present on the channels or if it is washing off.

This can be accomplished by simply flowing on SNAP in combination with heme following heme exposure. If wash-off is not sufficient to explain how SNAP reverses the heme effect, one potential explanation for these results could be the residues on S1 that appear to face the HxxxC motif among the published structures of CNBD- containing channels. Among the EAG family, the S1 residues consist of a proline-tyrosine dimotif, which is near universally conserved in EAG channels. A proline-phenylalanine diresidue has been suggested to allow NO- binding specificity to heme II of Cytochrome c554 of Nitrosomonas europaea (Andersson et al., 1986; Iverson et al., 1998; Upadhyay et al., 2006). It is possible that the conserved tyrosine in EAG channels is sufficient to maintain NO-binding, given that it too is aromatic. This hypothesis is supported by the result in GORK where

SNAP exposure largely leaves the heme effect more in-tact (Figure 6-3C,D). The putative residues on the S1 face opposite the HxxxC motif in GORK, based on sequence alignment, are a proline and a methionine. Lack of an aromatic residue next to the proline could explain the partial loss of NO efficacy.

136

Figure 6-3. Nitric Oxide has heterogeneous effects on heme-shifted CNBD-containing channels. A) Example traces of human Elk1 channels in control solution (black), after 10 µM SNAP exposure (blue), and after 10 µM hemin exposure (red). SNAP does not seem to have any effects on its own, however it appears to completely reverse the hemin-induced left-shift. B) The V50 of control Elk1 channels was estimated to be -68.4 ± 0.4 mV. This was shifted to -96.9 ± 1.6 mV upon hemin exposure. The V50 of left-shifted Elk1 channels shifted back to -69.8 ± 1.7 mV after SNAP (n = 3 for all compounds). C) Example traces of wild-type GORK channels in control solution (black), after 10 µM hemin exposure (red), and after 10 µM SNAP exposure (blue). The increase in current size was only partially mitigated by SNAP. Oocytes were pulsed to +60 mV with tails at 0 mV. D) Hemin increased GORK current size by 5.1 ± 0.4-fold at +60 mV compared to control solution. This was reduced to a 4.1 ± 0.3-fold increase compared to a control solution upon exposure to SNAP (n = 3 for all compounds). This is only a reduction in the hemin-induced current change of 20%.

The role of divalent cation binding on the heme effect must also be further elucidated. The data presented in Chapter 5 suggest that heme may allosterically potentiate the effects of divalent cations, at least within the

EAG family. The residues that are necessary for divalent cation inhibition of EAG channels are known (Zhang et al., 2009). It would therefore be interesting to test if these residues contribute to the heme effect. Preliminary results suggest that an aspartate residue in S2 may have a reduced hemin-induced left-shift compared to wild-type

Elk1 channels (Figure 6-4). D1 is known to be important in determining the selectivity of divalent cations that can interact with the voltage sensor (Zhang et al., 2009). Although this experiment needs to be replicated in more physiologic solutions, these results suggest that the aspartate residue, referred to as D1, may be important in

137 conferring the heme effect. The data shown in Figure 6-4 is insufficient on its own to determine whether D1 is an allosteric regulator of the heme effect or is directly interacting with the heme moiety. An analysis of the published structures of human Erg1 and rat Eag1 channels, both of which show the voltage sensor in the outward conformation (Whicher and MacKinnon, 2016; Wang and MacKinnon, 2017), demonstrates that D1 points in the direction of S1, near the locus of the conserved proline-tyrosine motif. Taken together, this would suggest that D1 may be important alongside the S5 HxxxC motif in order to confer the heme effect.

Figure 6-4. Elk1 D261A appear to be less responsive to hemin. A) Example traces of Elk1 D261A before and after exposure to 10 µM hemin. Red traces represent the current elicited at -40 mV. Oocytes were depolarized for 2 seconds and isochronal tail currents were collected at -40 mV. All recordings were performed in 98 mM Na+, 2 mM K+, 1 mM Ca2+, 6 mM Cl, 5 mM HEPES and pH 8. B) The GV of Elk1 D261A before (black) and after (red) exposure to hemin. The V50 of the control Elk1 D261A GV is approximately -48.2 ± 0.6 mV, whereas the V50 of hemin exposed Elk1 D261A channels is -64.2 ± 1.2 mV (n = 4 each).

It was shown in Chapter 5 that the GORK S3 residue C151 appears to be important in facilitating the heme effect in GORK. This residue is not very well conserved among the metazoan CNBD-containing channels.

In fact, it is unclear how C151 may interact with the HxxxC motif, if this is indeed the common pathway for channel-heme interaction, as the S3 transmembrane domain appears to be fairly distant from S5 in all of the published animal CNBD containing channels (Whicher and MacKinnon, 2016; Lee and MacKinnon, 2017; Li et al., 2017; Wang and MacKinnon, 2017). There are no published structures of any of the Plant CNBD-containing data, so it is possible C151 occupies a position closer to the HxxxC motif specifically in Plant CNBD-containing channel lineage. This would be in line with a model in which the conserved HxxxC motif allows for heme binding among all CNBD-containing channels, and lineage specific residues determine the mechanism and physiologic

138 role for the channel-heme interaction. It is also possible that the HxxxC motif is dispensable for the heme effect in

Plant CNBD-containing channels. Mutants of the S5 histidine are non-functional in the Plant CNBD-containing channel family; therefore, coexpression of S5 mutants with wild-type channels must be utilized to distinguish between these two possibilities.

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VITA

Aditya Pisupati

EDUCATION: PhD Neuroscience, The Pennsylvania State University, 2014-Present. Doctoral Thesis Advisor: Timothy Jegla, PhD MD, The Pennsylvania State University, 2012-Present. B.S. Engineering Science with Honors (Minor Bioengineering), The Pennsylvania State University, 2012. Thesis title: “The Minimal Cost of Blood Flow: Murray’s Law Revisited”. Undergraduate Thesis Advisor: Patrick Drew, PhD

PUBLICATIONS: Aditya Pisupati, Keith J. Mickolajczyk, William Horton, Damian B. van Rossum, Andriy Anishkin, Sree V. Chintapalli, Xiaofan Li, Jose Chu-Luo, Gregory Busey, William O. Hancock, Timothy Jegla (2018). S6 gate divergence in regulatory subunits restricts the stoichiometry of heteromeric Shaker family channels. J Gen Physiol. (submitted)

Timothy Jegla, Michelle M. Nguyen, Chengye Feng, Daniel J. Goetschius, Esteban Luna, Damian B. van Rossum, Bishoy Kamel, Aditya Pisupati, Elliott S. Milner, and Melissa M. Rolls (2016). Bilaterian Giant Ankyrins Have a Common Evolutionary Origin and Play a Conserved Role in Patterning the Axon Initial Segment. PLOS Genetics. 12(12):1006457

Awani Upadhyay, Aditya Pisupati, Timothy Jegla, Matt Crook, Keith Mickolajczyk, Matthew Shorey, Laura Rohan, Katherine Billings, Melissa Rolls, William Hancock, Wendy Hanna-Rose (2016). Vitamin B3 / Nicotinamide is an endogenous agonist for a C. elegans TRPV OSM-9/OCR-4 channel. Nature Communications. 12(7):13135.

CONFERENCE PRESENTATIONS AND SYMPOSIA: “Activation gate region influences stoichiometry of heteromeric Shaker family channels”. Aditya Pisupati, William J Horton, Keith Mickolajczyk, Xiaofan Li, Jose Chu, William O Hancock, Timothy Jegla. 2015. Poster. Presented at the 59th Annual Meeting of the Biophysical Society. Baltimore, Maryland.

HONORS/AWARDS: Recipient of the Huck Graduate Research Innovation Grant, 2016. Dean’s List, Fall 2008 through Spring 2012. Recipient of the Undergraduate Summer Discovery Grant, 2011.