An Atomic Scale Measurement from the Voltage Sensor in Herg Channels Using Lanthanide-Based Resonance Energy Transfer

An Atomic Scale Measurement from the Voltage Sensor in hERG Channels Using Lanthanide-Based Resonance Energy Transfer

by Danielle Jeong B.Sc., Simon Fraser University, 2014

Thesis Submitted in Partial Fulfillment of the Requirements for the Degree of Master of Science

in the Department of Biomedical Physiology and Kinesiology Faculty of Science

 Danielle Jeong 2016 SIMON FRASER UNIVERSITY Summer 2016

Approval

Name: Danielle Jeong Degree: Master of Science Title: An Atomic Scale Measurement from the Voltage Sensor in hERG Channels Using Lanthanide-Based Resonance Energy Transfer

Examining Committee: Chair: Dr. William Cupples Professor

Dr. Thomas Claydon Senior Supervisor Associate Professor

Dr. Peter Ruben Supervisor Professor

Dr. Glen Tibbits Supervisor Professor

Dr. Jenifer Thewalt External Examiner Professor Department of Physics

Date Defended/Approved: June 16, 2016

Ethics Statement

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Abstract

The cardiac human ether-a-go-go related gene (hERG) channel is a voltage- gated potassium (Kv) channel that plays a fundamental role in cardiac repolarization.

The importance of the hERG channel derives from its unusually slow activation

(opening) and deactivation (closing) processes. Like other Kv channels, structural reconfigurations of the hERG voltage sensor upon membrane depolarization and repolarization underlie channel activation and deactivation, respectively. However, specific rearrangements of the voltage sensor that may dictate the unique slow activation and deactivation in hERG channels remain unclear. Lanthanide-based resonance transfer (LRET) is a spectroscopic technique that has previously demonstrated an ability to provide quantitative description of voltage sensor dynamics in an archetypal Kv channel. In this report, we outline a rationalized approach to applying

LRET to examine hERG channel voltage sensor dynamics that may be of physiological significance. As a result, we report the first distance measurement from voltage sensors across the hERG channel pore.

Keywords: hERG channels; lanthanide-based resonance energy transfer; voltage- gated potassium channels; voltage sensor

Acknowledgements

This project would not have been possible without the selfless assistance of many individuals. As such, I would like to first show my appreciation for those who have notably contributed to its progress.

With respect to the donor chelate synthesis, Dr. Rikard Blunck from University of Montreal and Dr. Mattieu Starck, a post-doctorate fellow in Dr. Blunck’s lab, are to be recognized for their generosity in sharing the recipe they have developed. For graciously accommodating me to use their lab space and equipment for synthesizing the chelate, I would like to thank Dr. David Vocadlo from SFU Chemistry department and the members of his lab. Deep gratitude is specifically directed toward Hong Yee Tan, a PhD candidate from Dr. Vocadlo’s lab, and Hongwen Chen, a mass spectrometry specialist at SFU, for all the advice and patience they have provided throughout the synthesis troubleshooting phase.

When it comes to addressing technical aspects of the LRET setup, the talent of Pawel Kowalski, an electronic technologist at SFU, is to be admired and appreciated. He has demonstrated utmost ingenuity through various tasks, which includes installing the interlock system, designing and building the delay generator, re-engineering the PMT gating mechanism, and extending the lifespan of the laser used in this project. On matters concerning the optics, I was also fortunate to have an inexhaustible access to the expertise of Dr. Eric Lin, a post-doctorate fellow in Dr. Glen Tibbits’ lab. I cannot be more thankful for his involvement in the alignment of LRET system apparatuses, as well as continued guidance in the maintenance of optical components. As well, I gratefully acknowledge the work of the former post-doctorate fellow in Dr. Tom Claydon’s lab, Dr. Stanislav Sokolov, in initiating the LRET project and establishing a foundation for me. It is upon his shoulders I stand.

There are others who have offered more than a helping hand. Alison Li, a PhD candidate in Dr. Tibbits’ lab, is such an example. She not only has proactively arranged multiple sessions for me to use the MBB department spectrofluorometer and taught me how to operate the equipment, but has done so with persistent enthusiasm. The

kindness of Shyam Panchapakesan, a PhD candidate in Dr. Unrau’s lab, should also be noted in making the lab’s Nanodrop 2000 available for me to obtain the absorbance spectra recordings. Furthermore, I owe a heartfelt word of thanks to Ji Qi, a lab technician in Dr. Tom Claydon’s lab, for all her work in molecular biology. Her indefatigable ability to meet the demands to make all the constructs and cRNA used in this project is worthy of the highest praise.

Every step forward taken through this project could not have been achieved without the gifts of trust and encouragement from my supervisor, Dr. Tom Claydon. I cannot thank him enough for instilling within me a love and passion for scientific research when I was an undergraduate student, and for making my decision to pursue graduate studies to be un-regrettable. To have Dr. Claydon as a role model in science and leadership has been the greatest source of inspiration for me to become a better science student and person. I know that he will continue to have such an effect on me in the years ahead. On the same note, I would like to show my whole-hearted appreciation for the members of my supervisory committee, Dr. Peter Ruben and Dr. Glen Tibbits, for their fatherly support since my undergraduate years. It is unbelievable how privileged I have been to receive their wise advice (though I seldom took them), and to have mentors who would accept me just as I am- neurotic and obstinate. I am also overwhelmed with gratefulness when I think of members of the Molecular Cardiac Physiology Group. From those who have religiously checked on my happiness quotient to those who have always lent an ear, I could not have asked for better lab mates.

Most importantly, a special recognition should be given to my family for the relentless support they have provided behind the scene. I would like to specifically thank my dad for passing on his notorious stubbornness, for the same stubbornness in me was crucial in dealing with roadblocks encountered during the research process. I am also indebted to my mom for praying for me throughout the entire journey. The pillars of prayers that she had built were what had kept me standing strong. I will never be able to adequately express how grateful I am for my sister, Janelle, for injecting bursts of silliness back into my life whenever I was caught up in my work. Lastly, I would like to express my respect for my brother, Samuel, whose considerate nature often reminded me to look around and realize the people I have been blessed with during the duration of this project.

Table of Contents

Approval ...... ii Ethics Statement ...... iii Abstract ...... iv Acknowledgements ...... v Table of Contents ...... vii List of Tables ...... x List of Figures...... xi List of Acronyms and Symbols ...... xii

Chapter 1. Introduction ...... 1 1.1. Voltage sensing in voltage-gated ion channels ...... 1 1.1.1. Structural components of voltage-gated ion channels associated with voltage sensing ...... 2 1.1.2. The principle behind the voltage sensor ...... 3 1.1.3. Models of voltage sensor movement ...... 5 1.2. Techniques used to reveal voltage sensor movement ...... 6 1.2.1. Description of voltage sensor dynamics from electrophysiological approaches ...... 8 1.2.2. Structural insight on the voltage sensor ...... 9 1.2.3. Fluorescence spectroscopy reveals conformational changes of the voltage sensor ...... 10 1.3. Lanthanide-based resonance energy transfer (LRET) ...... 12 1.3.1. Principle of LRET ...... 12 1.3.2. Advantages of LRET ...... 14 1.3.3. Properties of the lanthanide and donor complex ...... 16 Terbium (Tb3+) ions ...... 16 Lanthanide coordinating complex ...... 17 1.3.4. Energy transfer in LRET ...... 18 1.3.5. Applications of LRET to study structural reconfiguration of the voltage sensor in voltage-gated ion channels ...... 20 1.4. Importance of the cardiac hERG channel ...... 21 1.4.1. Physiological role of the cardiac hERG channel...... 22 1.4.2. Voltage sensing in the hERG channel ...... 24 1.5. Overview and objectives ...... 25

Chapter 2. Materials and Methods ...... 26 2.1. Mutant channel preparation ...... 26 2.1.1. hERG1a monomer mutagenesis ...... 26 2.1.2. hERG1a dimer construction ...... 26 2.1.3. Shaker IR monomer construction ...... 27 2.2. Expression system ...... 29 2.2.1. Xenopus laevis oocyte preparation and injection ...... 30 2.3. Tb3+-DTPA-cs124-BMPH donor preparation ...... 30 2.3.1. DTPA-cs124-BMPH properties ...... 30

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DTPA ...... 31 Carbostyril 124 ...... 31 BMPH ...... 32 2.3.2. Chemicals reagents and materials ...... 33 2.3.3. DTPA-cs124-BMPH synthesis ...... 33 Reaction 1 ...... 34 Reaction 2 ...... 34 Precipitation, harvesting and storage ...... 34 Mass spectrometry result and notes on side products ...... 36 2.3.4. Tb3+ and DTPA-cs124-BMPH assembly ...... 37 2.4. Lanthanide-based resonance energy transfer ...... 39 2.4.1. Labeling ...... 39 2.4.2. LRET set-up ...... 40 2.4.3. LRET data acquisition...... 41 2.5. Two-electrode voltage clamp ...... 43 2.5.1. TEVC set-up and experiment condition ...... 43 2.5.2. Experimental protocols ...... 43 Ionic current recordings of the voltage-dependence of activation ...... 43 Simultaneous LRET and ionic current recordings ...... 44 2.6. R0 calculation ...... 47 2.6.1. Tb3+-DTPA-cs124-BMPH emission spectrum ...... 48 2.6.2. Acceptor absorbance and emission property ...... 49 TMRM ...... 49 MTSR ...... 50 Alexa 546 ...... 51 2.7. Data analysis ...... 53 2.7.1. Conductance-voltage relationship ...... 53 2.7.2. LRET signal lifetime decay ...... 53 2.7.3. Distance determination ...... 54

Chapter 3. LRET Experiment Approach and Results ...... 55 3.1. hERG dimer-of-dimer approach ...... 55 3.2. Tb3+ donor-only measurements ...... 61 3.2.1. Concentration dependence of Tb3+ emission ...... 61 3.2.2. Specificity of the Tb3+ donor complex signal ...... 64 3.3. RD at the resting membrane potential ...... 67 3.4. Voltage-dependent change in distance ...... 71

Chapter 4. Discussion ...... 76 4.1. Examining donor-only properties ...... 76 4.2. Interpreting sensitized emission data ...... 77 4.2.1. Evaluation of the time constants ...... 77 4.2.2. Comparison of LRET distance measurements to crystal structures ...... 79 4.2.3. Considerations for measured distances ...... 80 Movement of the donor and acceptor probe ...... 80 Temporal resolution ...... 80 Measurements from multiple sites ...... 82

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4.3. Future directions ...... 83 4.3.1. Other donor-acceptor pairs ...... 83 4.3.2. Detecting vertical distance ...... 83 4.4. Significance of the study ...... 84

References ...... 85 Appendix A. Side-products from DTPA-cs124-BMPH synthesis...... 101 Appendix B. Mass spectrum of DTPA-cs124-BMPH synthesis sample ...... 102

List of Tables

Table 1. J and R0 values for donor and acceptor pairs...... 51 Table 2. Tb3+-DTPA-cs124-BMPH properties at each labeling site...... 63

Table 3. RD from diagonally located voltage sensors in hERG and Shaker channels at resting membrane potential...... 69

Table 4. RD from diagonally located voltage sensors in hERG and Shaker channels at hyperpolarizing and depolarizing membrane potential...... 75

List of Figures

Figure 1. Structure of voltage-gated ion channels...... 3 Figure 2. Three structural models of voltage sensing...... 7 Figure 3. Principle of LRET and FRET when applied to ion channels...... 13 Figure 4. A representation of sensitized emission from a time-gated LRET experiment...... 15 Figure 5. The position of lanthanides in the periodic table...... 16 Figure 6. A Jablonski diagram of the lanthanide sensitization process...... 19 Figure 7. The role of hERG channel in cardiac repolarization...... 23 Figure 8. Gating of hERG channels...... 24 Figure 9. Schematic plasmid maps for hERG1a monomers and tandem dimer with their distinct patterns of restriction enzyme sites...... 28 Figure 10. Synthetic reaction scheme for DTPA-cs124-BMPH complex...... 35 Figure 11. Structure of the donor, Tb3+-DTPA-cs124-BMPH...... 38 Figure 12. LRET set-up and light path diagram...... 42 Figure 13. Voltage protocol used to examine for voltage-dependence of activation in hERG dimer-of-dimer channels...... 44 Figure 14. Voltage protocols applied to determine voltage or time dependent sensitized emission change...... 46 Figure 15. Emission spectrum of Tb3+-DTPA-cs124-BMPH...... 48 Figure 16. Normalized spectral data of the donor and acceptor...... 52 Figure 17. Multiple sensitized emission time constants result from channels composed of monomers...... 57 Figure 18. Single distance is determined from channels composed of dimer- of-dimer...... 58 Figure 19. hERG dimer-of-dimer channels demonstrate wild-type like function...... 60 Figure 20. Concentration-dependent properties of Tb3+-DTPA-cs124-BMPH...... 62 Figure 21. Specificity of Tb3+-DTPA-cs124-BMPH as a donor...... 66 Figure 22. LRET raw traces from the top of S4 in hERG dimer-of-dimer channels...... 70 Figure 23. LRET raw traces from hERG L520C dimer-of-dimers at hyperpolarizing and depolarizing potentials...... 73 Figure 24. Helical wheel of hERG channel S3-S4 linker...... 74

List of Acronyms and Symbols

A Relative amplitude of fit

A Absorbance at peak wavelength

Alexa 546 AlexaFluor546-maleimide

B-MPA ß-maleimidopropinic acid

BMPH N-ß-maleimidopropionic acid hyrazide c Concentration of the fluorophore

CS124 Carbostyril 124

DO Donor-only condition (in the absence of acceptors)

DTPA Diethylene-triamine-pentaacetic-acid

E Efficiency of energy transfer between donor and acceptor

ƐA(λ) Acceptor absorbance at a specific wavelenth

-1 -1 Ɛmax Molar extinction coefficient at peak absorbance, M cm

FD(λ) Donor emission at a specific wavelength

G-V Conductance-voltage hERG Human ether-à-go-go-related gene

IR N-type inactivation removed

J Normalized spectral overlap between donor emission and acceptor absorbance, M-1cm-1nm4

Ƙ2 Geometric orientation factor between donor and acceptor

Kv Voltage-gated potassium channel l Light path length

LRET Lanthanide-Based Resonance Energy Transfer

MTSR Sulphorhodamine methanethiosulphonate

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n Refractive index

ND96 Extracellular recording solution with 3 mM K+

QD Quantum yield of the donor in absence of the acceptor

R Distance in Å

R0 Förster distance when the efficiency of energy transfer is 50%

SE Sensitized emission

Tb3+ Terbium ion

TEVC Two-electrode voltage-clamp

TMRM Tetramethylrhodamine-5-maleimide

V1/2 Voltage at half maximal activation

VCF Voltage-clamp fluorimetry

VG Voltage-gated

WT Wild-type

λ Wavelength in nm

τ Time constant

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Chapter 1.

Introduction

1.1. Voltage sensing in voltage-gated ion channels

Voltage-gated (VG) ion channels are transmembrane protein complexes that detect changes in the membrane potential (a process referred to as voltage sensing), and respond by opening and closing an ion conducting pathway across the membrane.

The interplay of such channels selective for potassium (Kv), sodium (Nav), or calcium

(Cav) ions is responsible for physiological processes such as nerve impulse generation and propagation, neurotransmitter release and initiation of muscle contraction

(Armstrong & Hille, 1998). The crucial role that VG ion channels play in producing biological signals thus accentuates the importance of understanding the mechanism of the process that underlies and initiates all VG ion channel function: voltage sensing.

As such, immense research effort using various techniques has been directed to elucidate the structural and functional underpinnings of voltage sensing. Identification of the component in VG ion channels that is relevant to voltage sensing (the voltage sensor), as well as the general process involved is now agreed upon; however, consensus remains to be reached on the details of voltage sensor dynamics. These are described in the following sections, and key experimental evidence is presented with respect to the techniques used. In particular, spectroscopic approaches are highlighted as a powerful tool that correlates structural changes of the voltage sensor with its

function. Lanthanide-based resonance energy transfer is then specifically introduced as a milestone spectroscopic technique that can additionally provide quantitative descriptions of structural reconfiguration. The potential that LRET has in contributing to the knowledge of voltage sensing is realized when an unparalleled, thorough account of the voltage sensor movement in an archetypal VG potassium channel was provided with this technique. Finally, a specific VG ion channel of clinical significance, the human ether-a-go-go (hERG) channel, is discussed in terms of the significance of insights that will be gained from applying LRET to examine its voltage sensor dynamics.

1.1.1. Structural components of voltage-gated ion channels associated with voltage sensing

A discussion on VG channel voltage sensing begins with the structure. The structural composition of all VG channels is similar; Kv channels are made of four separate subunits (Fig.1) while Nav and Cav channels have four covalently connected domains, all of which ultimately assemble to form a central pore (Long, Campbell, &

Mackinnon, 2005; Yu & Catterall, 2004). Each subunit or domain, respectively, consists of six alpha-helical transmembrane segments (S1-S6), where the first four segments form a voltage-sensing domain (S1-S4), and segments S5 and S6 form the pore domain

(Catterall, 1988). The hallmark feature of VG channels and the segment of interest in investigating how electrical potential change is detected in VG channels is the S4, also referred to as the voltage sensor. This segment is characteristically identified by a conserved sequence motif of four to seven positively charged residues (arginine and lysine) spaced at three amino-acid intervals, known as the gating charges (Noda et al.,

1984). In addition, a number of negatively charged residues in S1-S3 segments have been noted to facilitate the voltage sensing process by stabilizing the positive charges of

the S4 segment in the hydrophobic milieu of the lipid bilayer (D. M. Papazian, Schwarz,

Tempel, Jan, & Jan, 1987).

Figure 1. Structure of voltage-gated ion channels.

A. A cartoon representation of a Kv channel subunit. Cylinders are helical segments. The voltage sensing domain is comprised of S1-S4 segments, with key charged residues indicated. The pore domain is formed by S5 and S6 segments. B. A ribbon representation of the Kv1.2 crystal structure (top view, extracellular side) is presented as an example of the tetrameric assembly formed by four subunits in Kv channels. Each subunit is shown in different colour. (B. from (Long et al., 2005). Reprinted with permission from AAAS.)

1.1.2. The principle behind the voltage sensor

The basis of voltage sensing is based upon the fact that the electric field across the lipid bilayer can be detected by electric charges or dipoles that change their position

or reorient according to changes in the field (Bezanilla, 2007). Before any knowledge on the structure of VG channels, let alone the existence of a distinct voltage sensing segment, was made available, this mechanism was first predicted by Hodgkin and

Huxley in 1952. They posited that Kv and Nav channels possessed four charged

“particles” residing in the membrane electric field that move between activated and deactivated positions in response to changes in voltage. Membrane depolarization would increase the probability for all four voltage sensors to be in the activated position, which in turn is necessary for channel opening (Hodgkin & Huxley, 1952). Direct evidence supporting this model was eventually provided by recordings of gating currents from the movement of gating charges across the membrane electric field (Armstrong & Bezanilla,

1973; Gilly & Armstrong, 1980).

In light of this, the conserved sequence of basic residues in the S4 segment noticed in the first cloned VG ion channel became a prime candidate for the voltage sensor (Noda et al., 1984). Now with the support of findings from many types of biophysical experiments, it is has been shown and widely accepted that the voltage sensor is pulled into the membrane at rest by the internally negative membrane electric field, and then moved outward by electronic repulsion upon depolarization (D. Papazian

& Bezanilla, 1997). This movement is then propagated to the pore domain for a concerted opening of the channel activation gate (Baker, Larsson, Mannuzzu, & Isacoff,

1998; Bezanilla, 2000; Horn, 1998; Larsson, Baker, Dhillon, & Isacoff, 1996; N. Yang &

Horn, 1995; Yusaf, Wray, & Sivaprasadarao, 1996). A dynamic description of voltage sensor movement, however, is still the subject of active debate.

1.1.3. Models of voltage sensor movement

A summary of the biophysical information and the discrepancies related to the voltage-dependent structural reconfiguration of the voltage sensor can be presented by three main, proposed models (Fig.2). In all cases, different combinations and amounts of

S4 rotation and translation are suggested, along with an explanation of how the thermodynamic barrier from having charged residues transit through low dielectric constant lipid bilayer is addressed (Parsegian, 1969).

Fig.2A shows the helical screw model, which consists of simultaneous S4 segment rotation along its axis and translation outward across the membrane in response to depolarization. As a result, the exposure of the voltage sensor charges is changed from the intracellular to the extracellular solution. In the original version of the model, S4 was proposed to rotate 60° and move 5 Å outward, whereas succeeding studies indicated 180° rotation and a distance change of 13 Å (Ahern & Horn, 2004;

Catterall, 1986; Durell, Shrivastava, & Guy, 2004; Gandhi & Isacoff, 2002; Guy &

Seetharamulu, 1986; Keynes & Elinder, 1999). Of note in the helical screw model is that the rotation accompanying the translocation allows a changing subset of the positively charged residues in S4 segments to make and break salt bridges with negatively charged residues in the voltage sensor domain (DeCaen, Yarov-Yarovoy, Scheuer, &

Catterall, 2011; D. M. Papazian et al., 1995). Further elaboration on the molecular basis of rotation has led to resurfacing of the 310 helix hypothesis: a notion that the voltage sensor transitions from an alpha helix to a 310 helix during translocation (Noda et al.,

1984). This idea has gained some favour, as a canonical 310 helix structure disposes residue side chains at every 120° (as opposed to 100° in an alpha helix); consequently, gating charges of the S4 would be aligned along the same side, and aptly placed within

a hydrophilic groove formed by S1, S2 and S3 helices (Jiang et al., 2003; Long, Tao,

Campbell, & MacKinnon, 2007a; Payandeh, Scheuer, Zheng, & Catterall, 2011).

Fig.2B depicts the transporter model. At resting position, the S4 gating charges are in a water crevice in continuum with the intracellular solution. Upon depolarization, the charges are in another water crevice that is in contact with the extracellular solution.

Such a change in the environment of the S4 was claimed to be achieved by a tilt and some rotation of the voltage sensor with minimal vertical translation (2-3 Å), along with rearrangements of the voltage sensing domain (Bezanilla, 2002; Chanda, Asamoah,

Blunck, Roux, & Bezanilla, 2005a; Starace & Bezanilla, 2004).

Fig.2C represents the paddle model, inspired by the crystal structure of the bacterial VG potassium channel, KvAP (Jiang et al., 2003; Jiang, Ruta, Chen, Lee, &

MacKinnon, 2003). This model is in stark contrast with the other aforementioned models; gating charges in the S4 are exposed to the hydrophobic milieu of the lipid bilayer without the protective enclosure formed by the voltage sensing domain. In this case, the

S4 segment and the extracellular portion of S3 (S3b) form a paddle that works as a unit and lies in the periphery of the voltage sensor domain at rest. Upon depolarization, it is postulated to move substantially across the lipid bilayer (20 Å) to reach the outer membrane surface (Ruta, Chen, & MacKinnon, 2005).

1.2. Techniques used to reveal voltage sensor movement

The available models of voltage sensor operation were proposed and shaped by experimental data from a variety of techniques. This was warranted since an understanding of the voltage sensor requires not only functional and structural

characterization, but also correlation between the two. In appreciation of the unique insights and details that each individual technique provides, some key approaches have been highlighted below.

Figure 2. Three structural models of voltage sensing.

A. Helical screw model, B. transporter model, and C. Paddle model. A single voltage sensor domain is presented for simplicity. White cylinders represent the S4 segment unless otherwise indicated. Not all gating charges on the voltage sensor are shown. Proteins that surround the S4 segment (S1-S3) are depicted as a dark gray structure. The hyperpolarized state is depicted on the left, while the depolarized state is shown on the right.

1.2.1. Description of voltage sensor dynamics from electrophysiological approaches

Exploration of voltage sensor movement in VG channels, in terms of the extent of exposure of gating charges to the intracellular or extracellular medium, has been achieved through accessibility experiments. An example is cysteine scanning mutagenesis, which tests the state-dependent ability of a water-soluble, cysteine reacting moiety (e.g. Methanethiosulphonate, MTS reagents) to attach to a cysteine engineered within the voltage sensor (N. Yang & Horn, 1995). When applied to human skeletal muscle sodium channels and Shaker potassium channels, a consensus was reached that the outermost charges were exposed to the outside upon depolarization and the innermost to the inside upon hyperpolarization (Baker et al., 1998; Larsson et al., 1996; N. Yang & Horn, 1995; Yusaf et al., 1996).

Similar findings were acquired via histidine scanning mutagenesis, which also tests exposure of S4 gating charges to solutions by exchanging individual charges for a histidine that can be titrated in a pH range more tolerable by the expression system. The idea is that if the histidine is not exposed to the solution, or if it does not move in response to the change in electric field, it would be un-titratable from either side and no change in gating currents is observed. In Shaker potassium channels, the first four charges were found to be accessible from both sides depending on the membrane potential (Starace, Stefani, & Bezanilla, 1997; Starace & Bezanilla, 2001; Starace &

Bezanilla, 2004). Such a high degree of accessibility observed in all cases, however, can be interpreted as either membrane-translocating movements of the S4 (as the helical screw model would imply) or rearrangements of crevices in the protein that provide a

pathway for MTS reagents to access the residues (as would be the case for the transporter model).

1.2.2. Structural insight on the voltage sensor

A three-dimensional structure of the channel, even in a single conformation, is considered invaluable as a structural framework for findings from electrophysiological studies. However, much dispute arose when the first solved crystal structure of a VG ion channel (KvAP) in the open state presented the S4 segment, along with the upper part of the S3 (S3b, Fig 2C), appeared to lie parallel across the bilayer (Jiang et al., 2003).

Such a conformation and the resultant paddle model raise an obvious issue of energetic cost from moving S4 gating charges through the hydrophobic milieu without any compensatory mechanism (Bezanilla, 2007). In response to critiques that argued for closer positioning of the S4 against the rest of the protein as per evidence from previous work, the subsequent crystal structure of the Kv1.2 channel presents the S4 segment almost perpendicular to the plane of the membrane (Li-Smerin, Hackos, & Swartz, 2000;

Long et al., 2005). The question of how the thermodynamic barrier is overcome remains to be addressed.

Another observation made from crystal structures of VG ion channels lends support for the helical screw model. The 310 helical structure of varying degree has been detected in the S4 segment. For example, in the open state crystal structure of Kv1.2/2.1 chimera, the upper part of the voltage segment carrying the first four positively charged residues (R0-R3) was found as an alpha helix (Long, Tao, Campbell, & MacKinnon,

2007b). Of particular interest was the region below R3, which adopted a 310 helix conformation and involved R4, K5 and R6 forming hydrogen bonds with negative charge

clusters in the voltage sensing domain. Based on this observation, the authors postulated a wave-like transition of S4 into 310 helix secondary structures during activation (Long et al., 2007b). This would also be reflected as a vertical stretching of the voltage sensor, since the rise per residue is 2.0 Å in a 310 helix and 1.5 Å in an alpha helix.

1.2.3. Fluorescence spectroscopy reveals conformational changes of the voltage sensor

By contrast to the above techniques, site-directed fluorescence has proven to be a powerful tool that can follow conformational changes in a protein. In the case of examining voltage sensor movement in VG channels, the general approach is to mutate the site of interest to a cysteine (often on top of the S4 segment) and label it with a fluorophore that has a thiol-reactive group. Resultant changes in fluorescence emission time course induced by imposed voltage pulses (voltage-clamp fluorimetry technique,

VCF) can infer qualitative insight on voltage sensor movement (Cha & Bezanilla, 1997;

Mannuzzu, Moronne, & Isacoff, 1996). For instance, the first direct correlation between displacement of most of the gating charge and voltage sensor movement was offered via this method in Shaker potassium channels (Cha & Bezanilla, 1997; Loots & Isacoff,

1998; Mannuzzu et al., 1996). Further detail was noted in that the kinetics of the fluorescence changes in the S4 segment are slower than around the S1-S2 linker, implying that there may be conformational changes preceding the main conformational changes associated with the voltage sensor (Cha & Bezanilla, 1997). Moreover, the capacity of voltage-clamp fluorimetry technique to track distinct voltage sensor reconfiguration was truly illustrated in Nav1.5 and Cav1.2, when fluorescence report of

the S4 from each domain displayed domain specific kinetics (Pantazis, Savalli, Sigg,

Neely, & Olcese, 2014; Varga et al., 2015).

In interpreting VCF results, however, some precautions should be taken. To detect fluorescence change in this specific application of tracking the voltage sensor, the conformational change involved must be substantial enough to change the environment around the fluorophore. Furthermore, it is not straightforward to interpret the observed change in emission, as it can reflect a strict change in the surrounding structures rather than the voltage sensor itself, or a movement of the site in question. Finally, the uncertainty associated with the exact location and orientation of the fluorophore further complicates data interpretation (Bezanilla, 2007).

The question on the extent of S4 translation across the bilayer upon depolarization has also been addressed with two variants of fluorescence resonance energy transfer (FRET), which is a technique that permits investigation of proximity between two moieties of interest based on energy transfer between the donor and acceptor fluorophore. In one study, green fluorescent protein was inserted as the donor after the S6 segment of the Shaker potassium channel. An organic fluorophore was attached in the extracellular regions of the channel as the acceptor, allowing computation of change in distance across the membrane between the two sites upon change in membrane potential. Despite inconsistency in distance measurements over a number of sites, the overall perceived change did not exceed 2 Å and thus indicated that there is little transmembrane translation of the S4 (Bezanilla, 2002). Further support to the transporter model was given by a second method that had used an organic fluorophore as a donor on top of the S4 segment, with a hydrophoblic organic anion dipycrilamine as an acceptor that distributes in the edges of the bilayer according to the

membrane potential. Though this approach was largely qualitative, it also suggested that a pure perpendicular motion of the voltage sensor would be minimal (Chanda, Asamoah,

Blunck, Roux, & Bezanilla, 2005b; Fernandez, Bezanilla, & Taylor, 1982).

1.3. Lanthanide-based resonance energy transfer (LRET)

LRET is a specific spectroscopic approach that can offer quantitative, in addition to qualitative, description of protein structural reconfiguration. Formerly, this has been achieved by FRET in wide range biological applications, and it has shown to be a useful tool for deriving the distance between two protein structures of interest with Ångstrom- level resolution (Selvin, 2002). That being said, there has been a recognizable struggle in drawing reliable distances measurements from FRET, where the source of the difficulty pertains to the properties of fluorophores used (section 1.3.2). This has led to modification of this standard technique to LRET, which involves a lanthanide ion and thus luminescence (as opposed to fluorescence) based resonance energy transfer. The technical advantages of LRET are discussed in detail in the succeeding sections, followed by an example of its application in examining voltage sensor movement in VG ion channels.

1.3.1. Principle of LRET

The underlying mechanism of LRET is similar to FRET. The idea behind resonance energy transfer is to label two sites of interest, one with a donor fluorophore and the other with an acceptor fluorophore. When an appropriate combination of donor and acceptor dye is chosen, (donor emission spectrum overlapping the acceptor absorbance spectrum), excitation of the donor probe results in a transfer of energy to the

nearby acceptor (within 10 to 100 Å) in a distance-dependent manner (Fig.3). The efficiency of energy transfer depends on the inverse sixth power of the distance between the dyes, spectral overlap between donor emission and acceptor absorption, and relative orientation of the donor and acceptor. Resultant fluorescence emission from the acceptor carries distance information between the donor and the acceptor (Selvin,

2002).

Figure 3. Principle of LRET and FRET when applied to ion channels.

A cartoon representation of how LRET and FRET work is shown in the context of an ion channel. LRET results from the distance-dependent energy transfer from an excited donor (D) to an acceptor (A), when attached at a site of interest within the channel. The properties of sensitized emission from the acceptor reflect the degree of energy transfer, and thus the distance from the donor.

1.3.2. Advantages of LRET

Distinct advantages of LRET are highlighted when compared to the specific drawbacks of FRET. The shortcomings of FRET derive from the use of organic fluorophores and their properties. First, the lifetime of typical organic fluorophores is a few nanoseconds. Since both donor and acceptor are organic fluorophores in FRET, sensitized emission also decays in nanoseconds, limiting the accuracy of lifetime measurements. As such, analysis of FRET data involves measurement of change in either the donor or acceptor intensity, which can also be less accurate due to artifacts

(Selvin, 2002). Second, the signal-to-noise ratio in sensitized emission signals is often poor in FRET. This is often due to contaminating fluorescence from the donor (due to substantial spectral overlap between donor and acceptor emission), or acceptor that has been directly excited by the light source (due to similar lifetimes of both donor and acceptor) (Selvin, 2002). Third, absolute distances are difficult to determine with confidence because of the uncertainty in the relative orientation of two dyes used

(Selvin, 1996).

The above limitations are resolved in LRET simply by replacing the donor with a lanthanide-based (typically terbium, reasons are provided in section 1.3.3) probe.

Lanthanides by nature have long lifetime decay in the range of milliseconds. Therefore, their use as a donor results in sensitized emissions also with lifetime decays that are milliseconds long, enabling accurate lifetime decay measurement that is less sensitive to artifacts than intensity measurements (Fig.4). Additionally, signal-to-noise ratio in LRET sensitized emission is incomparably better. This is attributed to the fact that there is a temporal separation between sensitized emission and fluorescence emission from acceptors that have been directly excited by the light source. The latter component can

therefore be removed from the sensitized emission signal by a time gate (Fig.4).

Lanthanide probes also have an emission spectral property that poses less of an overlap with the emission spectra of typical acceptor organic fluorophores. Hence, any donor emission that did not participate in energy transfer can be spectrally removed from the sensitized emission signal. Lastly, the lanthanide emission has been determined to be unpolarised. Moreover, since the sensitized emission occurs over a course of several milliseconds, the acceptor probe can undergo random rotational movement that can reduce the error imposed in distance measurements due to the unknown orientation factor (Selvin, 2002).

Figure 4. A representation of sensitized emission from a time-gated LRET experiment.

In LRET, sensitized emission is recorded after a delay in time (usually 50- 100 µs) to allow for elimination of interference from scattered light and autoflurorescence. Donor emission is spectrally removed by recording sensitized emission at wavelengths where donor emission is minimal. Due to the long lifetime decay property of the lanthanide-based donors, resultant sensitized emission is also several milliseconds (approximately 5 ms) long to allow for measurements of its decay time course.

1.3.3. Properties of the lanthanide and donor complex

The beauty of LRET can only be fully appreciated by understanding the properties of the lanthanide and its coordinating complex. Though a complete picture entails essential but dense concepts from physics and chemistry, key features relevant to LRET are mentioned here for focus and simplicity.

Terbium (Tb3+) ions

It is clear that various advantages of LRET are attributed to the long lifetime decay of the lanthanide-based donor probe. In turn, the lanthanide ion within the donor is responsible for the long lifetime decay. This property is innate to all lanthanide ions and their electronic configurations, where 4f orbitals are being filled with electrons (Fig. 5).

Within 4f orbitals, there are different energy levels over which electrons can be distributed (Werts, 2005).

Figure 5. The position of lanthanides in the periodic table.

The red box marks the location of the lanthanides in the periodic table relative to other elements. Terbium is indicated in grey.

With respect to the LRET application, Tb3+ ions are a common choice due to the optimal size of the energy gap for the main luminescent transition within the 4f orbitals

(from 5D to 7F energy states, Fig.6), and because emission is within the visible spectrum.

Other lanthanide ions are deemed inappropriate due to emission in the ultra-violet (Gd3+) or near-infrared (Ho3+, Er3+, Yb3+) range (Werts, 2005). Having said that, the electronic transition from 5D to 7F state is forbidden (referred to as Laporte forbidden rule). A direct consequence of this restriction is the low absorption coefficient of Tb3+ ions. This is the reason why the donor used in LRET is not simply the Tb3+ ion as it cannot be directly excited, but an entire complex that coordinates the lanthanide ion. A part of this complex is an antenna component that absorbs energy from the excitation source and transfers it to the lanthanide ion to overcome the restrictive barrier (Thibon & Pierre, 2009). Another direct consequence from the aforementioned transition is the long luminescence lifetime.

As a result of the forbidden nature, the luminescence that occurs from the transferred energy is prolonged and is ironically an asset for applications in resonance energy transfer technique (Bunzli & Piguet, 2005).

Lanthanide coordinating complex

The lanthanide donor complex is of interest as it overcomes the low absorptive nature of the lanthanide ion (<1 M-1 cm-1) and enables the lanthanide to luminesce in the first place. To maximize luminescence, there has been extensive design and production of lanthanide coordinating complexes (Bunzli & Piguet, 2005; J. Chen & Selvin, 1999;

Ge & Selvin, 2003; Selvin & Hearst, 1994; Thibon & Pierre, 2009). Some key features in the design of these complexes include: i) photostability as in resistance to photoinduced destruction of the excited complex, ii) incorporation of a protective structure for the lanthanide to minimize non-radiative de-excitation from solvent

interaction as this would diminish the quantum yield of the lanthanide, iii) strong absorption of excitation energy, and iv) efficiency of intramolecular energy transfer from the excited antenna component to the lanthanide, which in turn depends on the efficiency by which the triplet excited state of the antenna is populated (Bunzli & Piguet,

2005; Thibon & Pierre, 2009). Incorporation of these features in donor complex synthesis is exemplified in section 2.3.

1.3.4. Energy transfer in LRET

The summary of the LRET process is presented in Fig.6. It is a multi-step process that begins with absorption of a photon by the antenna molecule in the donor complex from excitation by a UV-light source (337 nm). As a result, there is a transition from the ground state (S0) to a higher electronic singlet state and its vibrational levels

-15 -12 (S1), in the order of 10 s. Excess energy is dissipated via vibrational relaxation (in 10 s), and the molecule assumes the lowest vibrational level of the excited singlet state. By the process of intersystem crossing, the triplet state (molecular electronic state where the excited electron has the same spin as the ground state electron) is populated from the excited singlet state (molecular electronic state where the excited electron has an opposite spin as the ground state electron). Both singlet and triplet states may transfer energy onto the lanthanide ion; however, due to the short lifetime of excited singlet state compared to the triplet state, energy transfer from the singlet state is considered negligible. From the triplet state, energy is transferred to a Tb3+ ion coordinated as a part of the donor complex. Excited electrons in the 5D energy level of the lanthanide ion then transfers energy to an acceptor in the vicinity (within 15 to 100 Å), which results in sensitized emission (over 10-3 s).

Figure 6. A Perrin-Jablonski diagram of the lanthanide sensitization process.

For simplicity, only the processes pertinent to LRET are shown. The multiple electronic states of molecules involved in the LRET process is shown via horizontal lines, each of which is split into a number of vibrational and rotational energy levels. Excitation of the organic antenna molecule in its ground state (S0) by a light source (solid grey arrow, 337 nm) results in populating a higher electronic state (S1). Excited electrons then undergo intersystem crossing (ISC), which yields electron in the triplet state (T1). Energy is transferred (intramolecular energy transfer, IET) to a Tb3+ ion nearby. Excited electrons in the 5D energy state can transfer energy to an acceptor nearby (lathanide-based resonance energy transfer, LRET), resulting in sensitized emission (dashed acceptor fluorescence arrow). Black arrows represent vibrational relaxation from the highest to lowest vibrational level within an energy state. Other potential processes that may occur (antenna fluorescence, antenna phosphorescence and lanthanide luminescence) are also shown as dashed grey arrows.

1.3.5. Applications of LRET to study structural reconfiguration of the voltage sensor in voltage-gated ion channels

As mentioned, there are a large range of distances that have been proposed for the voltage sensor to undergo, which demands for a technique with an ability to provide a more reliable measure of the distance change. LRET offers such an opportunity, as discussed in the previous section. This has been exemplified in a number of biological applications as well, such as measuring the degree of DNA bending from DNA-protein interactions, and the distance between dystrophin and actin in the muscle cell during interactions (Heyduk & Heyduk, 1999; Root, 1997).

The feasibility of applying LRET to VG ion channels, especially in examining voltage sensor dynamics, was shown in a number of studies using Shaker potassium channels. Distance measurement provided by LRET was first validated when the measurement of distance between residues in the pore region gave an agreement to within 1 Å to analogous residues in the KcsA crystal structure (Cha, Snyder, Selvin, &

Bezanilla, 1999). When donor and acceptor were positioned on top of the S4 segments, the distance between inter-subunit voltage sensors was around 50 Å. A small change in the radial distance (3 Å) was detected between closed and open state of the channel, which prevented completely ruling out the notion of translation across the bilayer (Cha et al., 1999). Additional recordings from three consecutive sites in the S3-S4 linkers further demonstrated a possible rotation and a tilt of the voltage sensor, lending support for the helical screw model (Cha et al., 1999).

More recently, a thorough investigation using LRET involved applying a donor on top of the S4 segment, while the acceptor was anchored to a toxin that blocks the pore of the Shaker channel from the extracellular side. Results confirmed the lack of a large

radial movement of the S4, where at all measured sites, the distance did not change more than 1 Å upon depolarization (Posson, Ge, Miller, Bezanilla, & Selvin, 2005). The extent of vertical displacement was also derived, which resulted in a moderate 5 Å that is inconsistent with a large 15-20 Å or near zero vertical movement purported by the paddle and transporter model, respectively (Posson et al., 2005; Posson & Selvin,

2008). Further argument against the paddle model was presented when the top portion of the S3 segment was observed to move 2 Å in the opposite direction of the S4 segment, suggesting that S3 and S4 segments do not move together as a rigid body

(Posson & Selvin, 2008).

1.4. Importance of the cardiac hERG channel

Due to its clinical significance, a VG ion channel that can offer benefits from better understanding the specific details of its voltage sensor movement is human ether- a-go-go related gene channel (hERG). hERG channels are abundantly expressed in the

+ heart tissue, where they conduct the delayed rectifier K current (IKr) that dictates the termination of cardiac action potential (Fig.7) (Sanguinetti & Tristani-Firouzi, 2006). The crucial role that hERG channels play in cardiac repolarization is highlighted by the suppression of hERG channel function being associated with long QT syndrome 2

(LQT2) (J. Chen, Zou, Splawski, Keating, & Sanguinetti, 1999; Curran et al., 1995).

LQTS is a cardiac repolarization disorder characterized by a prolonged QT interval on the electrocardiogram due to a prolonged action potential duration. LQTS2 is a variant that is specifically linked to reduced IKr conduction via hERG channels, whether as a result of congenital mutations in the hERG channel gene or by drug block (Huang,

Chen, Lin, Keating, & Sanguinetti, 2001; Sanguinetti, Curran, Spector, & Keating, 1996;

P. Yang et al., 2002). As affected individuals are identified with an increased risk of ventricular arrhythmia and sudden cardiac death, there is an emphasis on understanding the specific processes that underlie hERG channel function as a VG ion channel in both health and disease.

1.4.1. Physiological role of the cardiac hERG channel

As implied, the involvement of hERG channels in cardiac repolarization has been attributed to its unique gating properties. For most other voltage-gated potassium channels (Kv), activation (opening) and deactivation (closing) in response to depolarization and repolarization, respectively, occurs within milliseconds (Bezanilla,

Perozo, & Stefani, 1994). In contrast, the hERG channel activates and deactivates slowly over a time course of several hundreds of milliseconds, while inactivation and recovery from inactivation takes place at a much faster rate (Fig. 8) (Sanguinetti, Jiang,

Curran, & Keating, 1995; Smith, Baukrowitz, & Yellen, 1996). The contribution of hERG currents during the depolarization phase of the cardiac action potential is thus minimal, since the inactivation process is more rapid than activation (Fig.7). Robust repolarizing current that ultimately terminates the action potential is then observed upon membrane repolarization when hERG channels are rapidly recovering from inactivation, but are slow to close (Sanguinetti & Tristani-Firouzi, 2006). However, the molecular basis of such unusual, but physiologically essential gating characteristics of the hERG channel remains unclear.

Figure 7. The role of hERG channel in cardiac repolarization.

The effect of suppressing hERG channel function is represented in red with reduced hERG current, prolonged action potential duration and prolonged QT interval in an ECG trace. This is juxtaposed to the condition with normal hERG channel function, shown in black. Dashed line marks the point of cardiac repolarization.

Figure 8. Gating of hERG channels.

A cartoon representation of voltage-dependent conformation changes associated with gating of hERG channels is shown. Transitions between closed and open states are slower than transitions between open and inactivated states.

1.4.2. Voltage sensing in the hERG channel

The hERG channel structure is as expected for a typical Kv channel, which was mentioned in section 1.1.1. As such, its S4 segment is also characterized by a positively charged motif (K525, R528, R531, R534, R537 and K538), and is the main carrier of gating charges that establish electrostatic interactions with counter-charges in the S2 and S3 segments (Zhang et al., 2005).

The interest in hERG voltage sensor movement was raised with the first VCF experiment performed on three consecutive residues on top of the S4 (E518, E519 and

L520). When these sites were individually mutated to a cysteine and labeled with a fluorescent probe (TMRM), resultant fluorescent emissions were interpreted to have a slow component that correlated well with the unique voltage and time dependent properties of hERG ionic current activation and deactivation (Smith & Yellen, 2002). hERG gating current measurements that soon followed also reported that the kinetics of

gating charge movement (with majority assumed to be in the S4) were remarkably slower (τ of approximately 50 ms) than those reported for other Kv channels (Bezanilla,

2000; Piper, Varghese, Sanguinetti, & Tristani-Firouzi, 2003). In sum, these studies suggest that the voltage sensor movement is delayed in hERG channels, and that this slow movement may be crucial to understanding the mechanism behind the physiologically important slow activation in hERG channels. A recent report of VCF, however, has questioned interpretation of the previous VCF data, and instead has associated the slow component with rearrangements that accompany pore opening and closing (Es-Salah-Lamoureux, Fougere, Xiong, Robertson, & Fedida, 2010). Such discrepancies in addition to the potential physiological relevance emphasize the need to further examine the specifics of voltage sensor dynamics in hERG channels.

1.5. Overview and objectives

Application of LRET to examine the structural dynamics of hERG channel voltage sensors was considered in light of the fact that: i) LRET has previously demonstrated in

Shaker potassium channels of its ability to resolve complex, Ångstrom-level voltage sensor reconfigurations, ii) there is an implied physiological significance to understanding the detailed process of hERG channel S4 movement, and iii) the resultant data collected in hERG channels can be compared to those found in Shaker potassium channels. To our knowledge, this is the first instance where LRET is used to examine an aspect of hERG channel function. Therefore, the objective of this thesis was to provide: i) a careful experimental design using LRET that allows for an assessment of hERG channel gating, and ii) a step-by-step verification of the proposed approach.

Chapter 2.

Materials and Methods

2.1. Mutant channel preparation

2.1.1. hERG1a monomer mutagenesis

To resolve distances across the channel with respect to the S4 using donor and acceptor probes covalently linked to a thiol-reactive maleimide, a cysteine mutation was engineered into the S3-S4 linker sites (L520 or G516) that just precede the hERG S4 sequence and have previously shown to robustly report the voltage sensor movement

(Smith & Yellen, 2002; Thouta et al., 2014; Van Slyke et al., 2010). For each site, two extracellular native cysteines (C445 and C449) in the S1-S2 linker were also replaced by valine to ensure site-specific labeling. Mutant constructs were generated via overlap extension polymerase chain reaction, and incorporated into the pSK-XhoI expression vector between BstEII and XhoI sites. All constructs were transformed into DH5α competent cells in order to extract the plasmid DNA with a Qiagen Miniprep Kit. To confirm that errors were not introduced during the PCR cycling step, all constructs were sequenced using GeneWiz (South Plainfield, NJ).

2.1.2. hERG1a dimer construction

To circumvent the difficulty of resolving two distances offered by sensitized emissions from a homo-tetrameric protein (refer to section 3.1), two types of hERG 1a

monomer constructs (Type A and B) were produced and linked to generate a hERG 1a dimer construct (Fig.9A). Type A included a hERG 1a coding sequence with an engineered cysteine in the S3-S4 linker segment and with C445 and C449 replaced with valine. The stop codon and poly-A tail were subsequently removed, and an EcoRV site was added on the 3’ end just before the native EcoRI site of the pSK-XhoI vector. Type

B involved a hERG 1a coding sequence with C445 and C449 replaced with valine, but without any engineered cysteine for fluorophore attachment. In this construct, the stop codon and a poly-A tail on the 3’ end were kept, and an EcoRV site was introduced just before the start codon. Type A and B constructs were then cut with EcoRI and EcoRV, and ligated together to form the hERG dimer construct with an EcoRV site in between

(Fig.9B). XbaI restriction endonuclease was used to linearize the finalized construct, which in turn served as a template for cRNA synthesis with the mMessage mMachine T7

Ultra cRNA transcription kit.

2.1.3. Shaker IR monomer construction

To verify the ability of LRET to correctly resolve inter-subunit distances within a channel, two sites above the S4 (V363 and A359) in N-type inactivation removed Shaker

(Shaker IR, Δ6-46) channel that have been previously examined were considered for comparison (Cha, Snyder, Selvin, & Bezanilla, 1999; Posson & Selvin, 2008). For each cysteine introduced at the mentioned sites, an endogenous cysteine in the S1-S2 linker of Shaker-IR sequence (C245) was substituted with a valine. The mutagenesis was induced via the QuikChange kit (Stratagene), and the resultant mutant construct was inserted into pEXO between two flanking EcoRI sites. Constructs were sequenced as previously described (section 2.1.1). BstEII restriction endonuclease was used to linearize the resultant Shaker mutant monomer construct for cRNA synthesis.

Figure 9. Schematic plasmid maps for hERG1a monomers and tandem dimer with their distinct patterns of restriction enzyme sites.

A. Type A and B hERG 1a monomer constructs. The hERG 1a coding sequence is highlighted in red, with 3’ end represented by the arrow. The stop codon and poly (A) sequence (blue bar) is removed in Type A, whereas they are present in Type B. Engineered cysteine site for maleimide-linked fluorophore attachment was only introduced to Type A. Two native extracellular cysteines were replaced by valine in both constructs. B. The dimer construct as a result of linking Type A and Type B construct. The two hERG 1a sequences are joined with an EcoR V restriction enzyme site in between.

2.2. Expression system

The Xenopus laevis oocyte was the expression system of choice for all LRET experiments. One of the concerns from LRET experiments performed on a heterologous system in a living host is signal contamination due to non-specific labeling of endogenous channels in the membrane. This is in contrast to another approach that involves reconstituting purified proteins in artificial lipid vesicles, allowing for examination of the channel of interest in isolation (Faure, Starek, McGuire, Berneche, & Blunck,

2012; Richardson et al., 2006). However, the approach can be laborious, and more importantly, often results in a low protein yield (Miller, 1986).

In light of this, the Xenopus laevis oocyte expression system was specifically chosen as it expresses endogenous membrane channels and transporters at a much lower level than mammalian cell cultures used in heterologous studies (Goldin, 1991).

Additionally, the oocytes offer another advantage of a high expression of the exogenous channel of interest (Sigel, 1990). As is the case for many spectroscopic studies, such a large expression of the subject protein is imperative for LRET in order to achieve a large signal-to-noise ratio (Selvin, 2002). Reduced background signal is also achieved by imaging from the dark animal pole of the oocyte, as it offers lower light reflection (Goldin,

1991). Xenopus laevis oocytes were also considered suitable for LRET experiments as the fluorophores used in the study are impermeable to the oocyte membrane (Ge &

Selvin, 2003; Ge & Selvin, 2004). Otherwise, auxiliary advantages offered by the oocytes expression system included ease of harvest, maintenance and handling (Goldin,

1991).

2.2.1. Xenopus laevis oocyte preparation and injection

To obtain the oocytes, mature female Xenopus laevis frogs were terminally anaesthetized by immersion in ethyl-3-aminobenzoate methanesulphonate (2 g/L) for 15 minutes, as approved by Simon Fraser University Animal Care Committee in accordance with the Canadian Council on Animal Care guidelines. Ovarian lobes were then extracted and treated with 0.25 mg/mL collagenase type 1A in MgOR2 solution (96 mM

NaCl, 2 mM KCl, 20 mM MgCl2, and 5 mM HEPES) for 45 minutes, for subsequent selection and manual de-folliculation of Stage V and VI oocytes. Removal of the follicular layer was warranted due to i) potential contamination of ionic current recordings and

LRET recordings from endogenous ion channels and transporters present in the follicular layer, and ii) potential cell damage during cRNA injection as a result of the tough, collagenous property of the layer (Bossi, Fabbrini, & Ceriotti, 2007; Goldin, 1991).

The isolated oocytes were incubated in SOS+ media (96 mM NaCl, 2 mM KCl, 1.8 mM

CaCl2, 1 mM MgCl2, 5 mM HEPES, 5% horse serum, 2.5 mM sodium pyruvate, and 100 mg/L gentamicin sulfate, pH 7.4) in 19 °C for 2- 72 hours before injection with 50 nL (5-

10 ng) of cRNA with a Drummond digital microdispenser.

2.3. Tb3+-DTPA-cs124-BMPH donor preparation

2.3.1. DTPA-cs124-BMPH properties

Diethylene-triamine-pentaacetic-acid carbostyril124 N-ß-maleimidopropionic acid hyrazide (DTPA-cs124-BMPH, Fig.11) is a well-established lanthanide coordinating complex that satisfies the requirements mentioned in section 1.3.3. It has also repeatedly proved to be highly soluble in aqueous solutions, low in cytotoxicity, and

similar in size (12.8 x 8.3 x 8.1 Å in dimension) to typical organic fluorophores (Faure et al., 2012; Ge & Selvin, 2003; Selvin & Hearst, 1994; Selvin, Jancarik, Li, & Hung, 1996).

As such, DTPA-cs124-BMPH was the molecule of choice to form the donor lanthanide complex used in the presented LRET experiments.

DTPA

DTPA (Fig.10 and 11) is the chelate that tightly binds to lanthanide (terbium, in this case); titration with 100-fold excess of another chelate compound, ethylene-diamine- tetraacetic-acid (EDTA) was determined unable to displace a measurable amount of terbium (Selvin & Hearst, 1994). Moreover, DTPA has shown to effectively shield terbium from the quenching effects of water by providing 8 coordination sites that accommodate the 9 coordination sites of a Tb3+ ion (i.e. sites of potential bond formation), thus allowing for a high quantum yield of terbium (QTb= 0.48 in H2O) (Selvin &

Hearst, 1994; Werts, 2005; Xiao & Selvin, 2001). Another role of DTPA is to be a scaffold that orients the antenna moiety in the vicinity of the lanthanide (within 2.5 Å) to allow for efficient transfer of energy (Selvin, 2002; Selvin et al., 1996).

Carbostyril 124

Carbostyril 124 (Fig.11) is a chromophore that can act as an antenna in this case

-1 -1 by absorbing the excitation light (Ɛ338= 9600 M cm ) and transferring the resultant energy to the Tb3+ ion (Ge & Selvin, 2003; Selvin & Hearst, 1994). The energy transferring capacity of the antenna permits lanthanide chelate complexes in general to be considered much more photostable than organic fluorophores, as the excitation energy is transferred at a fast rate (Vuojola & Soukka, 2014). One of the criteria for successful, non-contaminating lanthanide excitation and luminescence is for the excited

triplet state of the antenna to be higher in energy than the excited 5D state of Tb3+ (see section 1.3.3 and 1.3.4). If the triplet energy state of the antenna is too high, then direct fluorescence from the antenna can be observed (a single emission peak occurs at 410 nm for cs124). If the triplet energy state of the antenna is lower than the 5D state of the lanthanide, then back energy transfer from the lanthanide to the antenna can ensue with subsequent decrease in lanthanide emission (Thibon & Pierre, 2009). Carbostyril 124 in conjunction with Tb3+ and DTPA has proven to be an antenna with an optimum triplet energy state in relation to the 5D state of a Tb3+ ion; neither direct antenna fluorescence nor substantial antenna-induced non-radiative de-excitation of excited-state Tb3+ has been reported (Li & Selvin, 1995).

BMPH

A maleimide-based thiol-reactive linker (Fig.11) was selected over another commonly used thiol-reactive group, iodoacetamides, as the latter has an additional potential to react with methionine, histidine and tyrosine residues. BMPH has been frequently used for biomolecular applications, notably with organic fluorophores used to track conformational changes in ion channels (Bannister, Chanda, Bezanilla, &

Papazian, 2005; Cha & Bezanilla, 1997; Mannuzzu et al., 1996; Savalli, Kondratiev,

Toro, & Olcese, 2006; Smith & Yellen, 2002; Thouta et al., 2014; Van Slyke et al., 2010).

In this instance, BMPH was covalently linked to the DTPA as a means of orienting the donor complex toward the engineered cysteine at targeted S3-S4 linker locations, as previously done in LRET experiments in Shaker IR channels (Cha et al., 1999; Posson et al., 2005; Posson & Selvin, 2008).

2.3.2. Chemicals reagents and materials

For DTPA-cs124-BMPH complex synthesis, the following reagents were purchased from Sigma-Aldrich: molecular sieves (4 Å beads, 4-8 mesh), diethylene- triamine-pentaacetic-dianhydride (DTPA), diethyl ether (inhibitor free), dimethylsulfoxide

(DMSO, anhydrous form), trimethylamine (TEA), carbostyril 124 (cs-124), tetrahydrofuran (anhydrous and inhibitor free form), and terbium (III) chloride

(anhydrous, powder). N-ß-maleimidopropionic acid hyrazide (BMPH) was purchased from Pierce Biotechnology. All glassware was washed with acetone, rinsed with distilled and deionized water, and completely dried in a bench-top oven prior to use.

2.3.3. DTPA-cs124-BMPH synthesis

A DTPA-cs124-BMPH synthesis procedure was generously provided by Dr.

Rikard Blunck and Dr. Mattieu Starck, which was adapted from previous work on thiol- reactive luminescent chelate complex synthesis (J. Chen & Selvin, 1999; Faure et al.,

2012; Ge & Selvin, 2003). Broadly, the synthesis was a simple two step reaction in which an anhydride group of DTPA dianhydride was first reacted with the 7-amino group of cs124. The remaining anhydride group of the intermediate product was then reacted with the amino group of BMPH.

All reagents and reactions were considered to be either air or moisture sensitive.

As such, complex synthesis was achieved under constant argon pressure. This was set up by inserting an argon filled balloon attached to a syringe and needle through a septum enclosing the vessel in which reaction took place. Solvents and solutions throughout the synthesis experiment were consequently added to the reaction vessel using a syringe and a needle to maintain the inert environment.

Reaction 1

DTPA anhydride (25.0 mg, 70 µmol) was dissolved in 0.5 mL of anhydrous

DMSO in an argon environment. Triethylamine (7.0 µL, approximately 70 µmol) dried over molecular sieves overnight was added to the DTPA-DMSO solution. Solution of cs124 (13.4 mg, 77 µmol) dissolved in 0.5 mL anhydrous DMSO in a separate vessel, was added dropwise via needle and syringe to the DTPA solution, to produce the DTPA- cs124 intermediate (Fig.10). The mixture was constantly stirred with a stir bar, and the reaction was carried out at room temperature for 1 hour under constant argon pressure.

Reaction 2

BMPH powder (22.9 mg, 77 µmol) was prepared in a separate flask and placed under argon pressure. DTPA-cs124 mixture was then added using a syringe and needle.

The resultant mixture was stirred, and the reaction was allowed to continue for another hour at room temperature to form the desired DTPA-cs124-BMPH (Fig.10). The final solution with all components dissolved appeared completely clear.

Precipitation, harvesting and storage

To precipitate the product, anhydrous tetrahydrofuran and diethyl ether were added in alternating order while the solution was constantly stirred, until white precipitate was observed (approximately 20 mL of each solvent in total). Once the precipitate had settled down to the bottom of the flask, most of the supernatant was removed using a needle and a syringe. The remaining precipitate was washed several times with copious amount of diethyl ether, sonicating in between to ensure all traces of DMSO were removed. Approximately 50 mg of precipitate was then dried under nitrogen introduced by a Schlenk line, and subsequently stored in a desiccator at -20 °C kept in the dark.

Figure 10. Synthetic reaction scheme for DTPA-cs124-BMPH complex.

The two step reaction involved in DTPA-cs124-BMPH synthesis is represented above as Reaction 1 and Reaction 2. The schematic was generated with ChemDraw (courtesy of Hong Yee Tan).

Mass spectrometry result and notes on side products

As implied, DTPA-cs124-BMPH complex synthesis is based on random interactions and reactions between individual DTPA, cs124, and BMPH components.

Thus from many different combinations of interactions that can occur, it is plausible for other side-products to form, and for the final precipitate to entail both the desired and undesired products. The structure and molecular weight of predicted side products are presented in Appendix A. Therefore, it is favourable to purify the final precipitate. Other previous chelate complex synthesis efforts have reported using high performance liquid chromatography (HPLC) to separate out the desired product (J. Chen & Selvin, 1999;

Ge & Selvin, 2003). In the present synthesis, however, the feasibility of such a task was determined to be low from discussions with various experts, due to similarity in polarity of all the products involved (refer to structures in Fig. 11 and Appendix A).

In labeling channels with unpurified donor synthesis mix, specific side products of concern with respect to LRET experiments were BMPH-DTPA-BMPH (MW 723.65 g/mol), DTPA-BMPH (MW 705.73 g/mol), and OH-DTPA-BMPH (MW 558.50 g/mol) since they can bind to the engineered cysteine site and reduce the apparent efficiency of labeling. To reduce the chance of various DTPA-BMPH molecules from forming, the first reaction between DTPA and cs124 was hence conducted with cs124 in excess (1:1.1 ratio between DTPA and cs124) in the attempt to drive the formation of molecules with

DTPA and cs124 (refer to section 2.3.3, reaction 1). This was based on the rationale that any side products that form consisting of just DTPA and cs124 are more tolerable since they can be readily removed through washes after labeling. The existence of the desired product, DTPA-cs124-BMPH (MW 714. 69 g/mol) was verified by submitting a sample of the precipitate for mass spectrometry (Bruker micrOTOF) (Appendix B). Additional

confirmations of the lanthanide chelate complex’s function and reactivity (despite not undergoing a purification process) are presented through the emission spectrum (section

2.6.1, Fig.15) and donor luminescence lifetime decay measurements (section 3.2).

2.3.4. Tb3+ and DTPA-cs124-BMPH assembly

Tb3+-DTPA-cs124-BMPH is a donor complex (Fig. 11) that has been well described in previous studies applying LRET to investigate the structural dynamics of ion channels during gating, and was thus used for all LRET experiments on the hERG channel outlined in section 3 (Cha et al., 1999; Faure et al., 2012; Posson et al., 2005) .

For each experiment, a new sample of donor complex was made by introducing

TbCl3 in excess to a 1 mg aliquot of DTPA-cs124-BMPH reconstituted in 30 µL of distilled and deionized water to ensure that all chelates are occupied with a Tb3+ ion. The complex formation between the lanthanide ion and chelate molecule involves entropy and enthalpy changes accompanied by dehydration of both components. Though decrease in hydration yields positive entropy change that favours complexation, dehydration is also endothermic. Contributions from coordination between the lanthanide ion and chelate molecule are deemed inadequate in compensating for this unfavourable energy demand (Bunzli & Piguet, 2005). As such, a total of 3 µL of 1 M of TbCl3 was added to DTPA-cs124-BMPH solution over 30 minutes in 1 µL increments, vortexing in between to facilitate the coordination process. The 33 µL donor complex was further diluted with depolarizing solution (99 mM KCl, 1 mM MgCl2, 5 mM HEPES, 2 mM CaCl2, at pH 7.4) to obtain the working concentrations used for labelling. Total volumes ranging from 500 µL to 5 mL were tested in order to determine optimum relative concentration, since absolute concentration of the chelate complex was unknown (see section 3.2.1).

Figure 11. Structure of the donor, Tb3+-DTPA-cs124-BMPH.

The donor complex used in the LRET experiments is shown, where DTPA is the chelate that coordinates Tb3+ ion, thiol reactive maleimide linker (BMPH), and the antenna (cs124). DTPA-cs124 complex dimension based on resolved crystal structure has been determined to be 12.8 x 8.3 x 8.1 Å (Selvin et al., 1996).

2.4. Lanthanide-based resonance energy transfer

2.4.1. Labeling

Oocytes injected with cRNA were incubated in SOS+ medium (96 mM NaCl, 2 mM KCl, 1.8 mM CaCl2, 1 mM MgCl2, 5 mM HEPES, 5% horse serum, 2.5 mM sodium pyruvate, and 100 mg/L gentamicin sulfate, pH 7.4) at 12 °C for 18 to 24 hours to inhibit trafficking of hERG channels to the membrane. Native membrane cysteine residues were then blocked by 1 hour treatment in 500 μM ß-maleimidopropinic acid (Sigma) in order to increase the specificity of donor and acceptor probe labeling. Following this treatment, oocytes were incubated for 48 hours (Shaker IR homo-tetrameric channels) or 5 to 7 days (hERG dimer-of-dimers) at 19-22 °C to allow for trafficking of exogenous channels to the membrane (Mannuzzu et al., 1996). All labeling was performed on cells exhibiting large expression of the subject channel (approximately -10 to -40 µA of -110 mV peak tail current amplitude for hERG channels and -40 to -400 µA peak current amplitude for Shaker IR channels) to guarantee large signal-to-noise ratios. For the donor-only condition (DO), oocytes expressing Shaker or hERG channels were labelled with approximately 25 μM thiol-reactive terbium chelate complex (Tb3+-DTPA-cs124: synthesized in-house, commercial Tb3+-chelate: Invitrogen) in depolarizing solution (99 mM KCl, 1 mM MgCl2, 5 mM HEPES, 2 mM CaCl2, at pH 7.4) for 1 hour on ice and in the dark. For the donor and acceptor condition, the terbium chelate complex and tetramethylrhodamine-5-maleimide (TMRM) was mixed in 1:1 ratio (25 μM: 25 μM). The labelled oocytes were thoroughly rinsed twice in ND 96 solution (96 mM NaCl, 3 mM

KCl, 0.5 mM CaCl2, 5 mM HEPES, at pH 7.4) prior to LRET recording to wash away

BMPH-less side products from chelate complex synthesis, as well as to remove any non-specifically bound probes.

2.4.2. LRET set-up

The fluorescence lifetime decays were recorded using the set-up shown in

Fig.12. The channel expressing, labelled oocyte was held in a bath chamber with a quartz coverslip on the side facing the objective lens. In both donor only and donor- acceptor labelled cases, lanthanide donors were excited by UV light (337 nm, 3.5 ns pulse) supplied by a nitrogen laser (NL100, Stanford Research Systems), that was reflected off of a 364LP dichroic mirror and passed through a 20X quartz objective of a

Nikon inverted microscope. When the channels were labelled with both donor and acceptor, the resultant sensitized emission was passed through the same objective and dichroic mirror, and then filtered through a bandpass (565WB20) and a longpass filter

(565ALP). The two emission filters were placed in series to ensure that the 545-565 nm donor emission (see section 3.2.2 and Fig.16) was excluded from the sensitized emission signal (sensitized emission effectively measured at 565-585 nm). When the channels were labelled with the donor only, emission from the donor was collected by replacing the emission filters 565WP20 and 565ALP with 485DF22 to capture the first emission peak of the terbium chelate complex (Fig.15). Fluorescence from acceptors or luminescence from donors were both detected by a water-cooled (10 °C) photomultiplier tube (Hamamatsu, R943-02) operated at -1300 to -1400 V. All signals were digitized by

Axon Digidata 1400A A/D converter (Axon Instruments) and processed by a computer with pClamp10.5 software (Axon Instruments) to determine the emission intensity and decay time course. An innate 50 ms delay between the PC input signal and the trigger signal to the laser was in place for laser charging. A 100 μs delay between the laser sync signal and the PMT gate opening was inserted by a delay generator. This time gate served two purposes: i) to allow for fluorescence from directly excited acceptor to decay to zero before sensitized emission was recorded, and ii) to prevent prompt fluorescence

from saturating the PMT. The PMT gate was also held open by the delay generator for at least 10 ms to fully capture the LRET signals, which were typically about 5 ms in duration.

2.4.3. LRET data acquisition

LRET data were acquired at the maximum permitted sampling frequency for each protocol: 250 kHz for measurements at the resting membrane potential and 200 kHz for fluorescence measurements collected during a voltage-clamp protocol. Ionic current recordings were concomitantly collected for all LRET recordings involving a voltage-clamp step, as a means of monitoring channel function and cell health after labeling (section 2.5 for TEVC conditions and protocols). All lifetime data were obtained as averages of consecutive pulse protocols (20 to 40) to optimize signal-to-noise ratio without damaging the cell or photobleaching the probes used, as has been done previously (Hyde et al., 2012). Consecutive pulses were repeated with a pulse interval of

1 s for measurements from the resting membrane potential, and 5 s for voltage-clamp conditions. Extracellular cysteine-removed hERG WT channels that were treated identically to experimental conditions were used as negative controls to provide a measure of background signal (section 3.2.2).

Figure 12. LRET set-up and light path diagram.

The terbium chelate complex (donor) excitation source was supplied by a nitrogen laser (A). The 337 nm UV beam was reflected off of a 364LP dichroic mirror (B1) and passed through a 20X quartz objective (C) before reaching the labelled Xenopus oocyte in the cell bath chamber (D). The emission filter arrangement shown (565WB20, B2; 565LP, B3) was for recording sensitized emission when the cell was labelled with both donor and acceptor. When recording the donor emission only, B2 and B3 were replaced by 485DF22 emission filter (not shown). All emissions were detected by a photomultiplier tube (E). The fluorescence signal was digitized by an A/D converter (G) and processed by a computer with pClamp software (H). A delay generator (I) was also in place to insert a 100 µs delay between the laser sync signal and PMT gate, as well as to keep the PMT gate open for 10 ms. Voltage-clamp of the oocyte was enabled using the two-electrode voltage-clamp technique with an Axoclamp 900A amplifier (F).

2.5. Two-electrode voltage clamp

2.5.1. TEVC set-up and experiment condition

Simultaneous voltage-clamp of the oocyte was achieved via the two-electrode voltage-clamp technique (Fig.12) using the Axoclamp 900A (Axon Instruments) amplifier coupled to the Digidata 1440 interface (Axon Instruments). The recording microelectrodes were made of thin-walled borosilicate glass, with a resistance of 0.4-1.0

MΩ after the tips were manually broken, and filled with 3 M KCl. Computer driven voltage protocols were generated and data acquisitions were obtained with pClamp10.5 software (Axon Instruments). Ionic current data were sampled at 10 kHz and low-pass filtered at 4 kHz with a Bessel filter. All experiments were performed in ND96 solution

(96 mM NaCl, 3 mM KCl, 0.5 mM CaCl2, 5 mM HEPES, at pH 7.4) and at room temperature (20-22 °C).

2.5.2. Experimental protocols

Ionic current recordings of the voltage-dependence of activation

For electrophysiological characterization of hERG WT and mutant channel voltage-dependent activation, the voltage protocol used is shown in Fig.13. Steady-state conductance-voltage (G-V) relationships were determined from peak tail currents recorded during a voltage step to -110 mV, following 4 s depolarizing pulses to +60 mV in 10 mV increments from a holding potential of – 80 mV.

Figure 13. Voltage protocol used to examine for voltage-dependence of activation in hERG dimer-of-dimer channels.

The voltage protocol was applied to collect ionic current recordings from hERG dimer-of-dimer channels in order to verify that their voltage- dependent activation was akin to that of hERG WT channels. Conductance-voltage relationship was determined from the peak tail currents generated at -110 mV.

Simultaneous LRET and ionic current recordings

Voltage-dependent change in distance in hERG channels

To detect voltage-dependent change in distance between the two voltage sensors diagonally located across the pore in hERG dimer-of-dimers, the voltage protocol presented in Fig.14A was applied. Ionic current traces were concomitantly recorded by clamping the oocyte at a holding potential of -80 mV, then stepping to a single depolarizing potential of +60 mV for 2 s, and returning to a -60 mV step.

Sensitized emission was recorded at -80 mV just prior to the depolarizing step, and once

again at 2 s into the depolarizing potential (marked by arrows in Fig.14A), in order to compare the voltage-dependent change.

Voltage-dependent change in distance in Shaker IR channels

Shaker IR mutant channels (Shaker IR C245V V363C and Shaker IR C245V

A359C) were considered as a positive control, to serve as a comparison to what has been reported in the literature. As such, voltage protocols applied to each mutant channel were adapted from the corresponding literature that has reported distance measurements using the same donor-acceptor labeling sites.

For Shaker IR C245V V363C channels, the voltage protocol consisted of holding at -90 mV before stepping down to -120 mV, followed by a step up to +50 mV (Fig.14C).

Voltage-dependent sensitized emission changes were measured from LRET recordings

50 ms into the -120 and +50 mV step (Cha et al., 1999). For Shaker IR C245V A359C channels, the voltage protocol consisted of holding at -100 mV before stepping down to -150 mV, followed by a step up to +25 mV (Fig.14B). Voltage-dependent sensitized emission changes were measured from LRET recordings 50 ms into the -150 and +25 mV step (Posson & Selvin, 2008).

Figure 14. Voltage protocols applied to determine voltage or time dependent sensitized emission change.

A. Voltage protocol used to consider voltage-dependent change in the diagonal distance across the pore of hERG channels. Sensitized emission was recorded at the end of 2 s depolarizing pulse. B. Voltage protocol applied to detect voltage-dependent change in sensitized emission in Shaker IR C245V V359C. C. Voltage protocol applied to detect voltage-dependent change in sensitized emission in Shaker IR C245V A363C. Arrows indicate the time point at which LRET recording was taken.

2.6. R0 calculation

R0 is defined as the Förster distance at which energy transfer is 50% efficient (E=

0.5). It is a constant where the value depends on the spectral properties of the donor and acceptor used to acquire the sensitized emission (Selvin, 2002). It can be derived using the following expression:

2 -4 1/6 R0 = 0.211(Ƙ n QDJ) in Å

Ƙ2 is the geometric orientation factor pertaining to the relative orientation of the transition dipoles of the donor and acceptor, as well as their relative orientation in space. In LRET,

Ƙ2 is determined to be 2/3, based on the assumption that both donor and acceptor rapidly and completely rotate during the millisecond lifetime of the lanthanide donor

(Selvin, 2002). n is the refractive index, which is set to be 1.4 when LRET measurements are taken in an aqueous salt solution. QD is the donor quantum yield in the absence of the acceptor, and was measured to be 0.48 for Tb3+-DTPA-cs124 donor complex (Xiao & Selvin, 2001). J is the normalized spectral overlap of the donor

-1 -1 emission (FD) and acceptor absorption (ƐA in M cm ) with λ in nm. As implied, J was determined for every donor and acceptor combination, based on the integral equation:

4 -1 -1 4 J= ʃ FD(λ)*ƐA(λ)*λ dλ/ ʃ FD(λ)dλ in M cm nm

3+ J and R0 value for three combinations, Tb -DTPA-cs124-BMPH with TMRM,

MTSR or Alexa546, were calculated using EXCEL (Microsoft). The summary of the values and comparison with values previously reported in literature can be found in

Table.1.

2.6.1. Tb3+-DTPA-cs124-BMPH emission spectrum

3+ Donor only emission spectrum, FD(λ), of Tb ion complexed with in-house synthesized DTPA-cs124-BMPH, was measured in a quartz cuvette (Hellma Analytics) using a fluorescence spectrophotometer (Cary Eclipse Varian) at room temperature. The donor complex was reconstituted in depolarizing solution (99 mM KCl, 1 mM MgCl2, 5 mM HEPES, 2 mM CaCl2, at pH 7.4) to a final concentration of 25 µM, as in the donor- only labelling condition stated above (section 2.4.1). The spectrophotometer settings included λex = 337 nm, λem = 450-650 nm, excitation and emission slit width = 5 nm. The resultant emission spectrum is presented in Fig.15. It is notably comparable to previous reports of the Tb3+ emission spectrum when complexed with DTPA-cs124 as well as other lanthanide chelate/antenna moiety, thus confirming proper function of the synthesized DTPA-cs124-BMPH complex (Selvin & Hearst, 1994; Selvin, 2002; Thibon

& Pierre, 2009).

Figure 15. Emission spectrum of Tb3+-DTPA-cs124-BMPH.

Donor emission spectrum was obtained from exciting the complex with a 337 nm light source. The maximum emission peak of Tb3+ luminescence was at 546 nm, akin to the reported value in literature (Selvin, 2002).

2.6.2. Acceptor absorbance and emission property

The absorbance spectra for all acceptors considered for LRET were collected at room temperature using a Nanodrop 2000 UV-Vis spectrophotometer (ThermoFisher).

Acceptors were reconstituted in depolarizing solution (99 mM KCl, 1 mM MgCl2, 5 mM

HEPES, 2 mM CaCl2, at pH 7.4) to a final concentration of 25 µM (c). Path length (l) that the light traveled through was determined to be 1 mm. Therefore, once the absorbance at the peak absorbance wavelength (A) was measured, the molar extinction coefficient

(Ɛ) at each wavelength was calculated from the Beer-Lambert Law:

A= Ɛlc

The emission spectra for all acceptors were recorded in a quartz cuvette (Hellma

Analytics) using a fluorescence spectrophotometer (Cary Eclipse Varian) at room temperature. The spectrophotometer settings consisted of: excitation wavelength at 545 nm (peak wavelength at which the energy transfer occurs during LRET experiments when using a Tb3+-based donor), emission range of 450-700 nm, and excitation and emission slit width at 5 nm.

TMRM

Tetramethylrhodamine-5-maleimide (TMRM; Toronto Research Chemical) was an optimal candidate acceptor with respect to a Tb3+-based donor due to i) its peak excitation wavelength (550 nm) corresponding closely to the peak emission wavelength of the Tb3+ donor (545 nm), and ii) its rising emission being situated where the donor emission is absent (around 565 nm), which allows for a spectral separation of donor emission and sensitized emission (Fig.16A). TMRM also has a high extinction

coefficient (110 000 M-1cm-1 in methanol), and has a maleimide-linker for anchoring the fluorophore at specifically engineered cysteine sites.

The calculated Ɛmax at 550 nm for 25 µM TMRM in depolarizing solution was 87

920 M-1cm-1, comparable to the known value in methanol. The ensuing calculated J value (4.42 x 1015 M-1cm-1nm4) was also similar to the value previously reported for the

Tb3+-DTPA-cs124 and TMRM combination (3.80 x 1015 M-1cm-1nm4) (Posson et al., 2005;

3+ Selvin, 2002). The resultant R0 for this experimental condition between Tb -DTPA- cs124-BMPH and TMRM combination was 56.2 Å (refer to Table.1 for a summary), which closely followed the value described (57 Å) for an analogous donor-acceptor pair

(Posson et al., 2005; Selvin, 2002).

MTSR

Sulphorhodamine methanethiosulphonate (MTSR; Toronto Research Chemical) is also a TMR-based fluorophore with a methanethiosulphonate group (MTS) as the sulfhydrol reagent that readily reacts with cysteine residues. Its measured peak absorbance and emission at 570 and 590 nm, respectively, allowed for it to be considered as an acceptor to be used in conjunction with a Tb3+ donor complex

(Fig.16B).

The calculated Ɛmax at 570 nm for 25 µM MTSR in depolarizing solution was 10

240 M-1cm-1, and the calculated J value was 3.68 x 1014 M-1cm-1nm4. The consequential

3+ R0 for Tb -DTPA-cs124-BMPH and MTSR combination was 37.2 Å (refer to Table.1), which was dissimilar to the value mentioned (60 Å) for an equivalent donor-acceptor pair

(Richardson et al., 2006).

Alexa 546

AlexaFluor546-maleimide (Alexa 546; ThermoFisher) is another notable acceptor that demonstrates strong absorption with extinction coefficient greater than 65 000 M-

1cm-1, recognizable photostability, and a quantum yield much greater than that of TMRM.

The peak absorbance and emission wavelengths of Alexa 546 (555 and 571 nm, respectively) also enable it to be paired with a Tb3+-based donor (Fig.16C).

The calculated Ɛmax at 555 nm for 25 µM Alexa 546 in depolarizing solution was

90 280 M-1cm-1, comparable to the known value in methanol (112 000 M-1cm-1). The

15 -1 -1 4 3+ calculated J value was 3.56 x 10 M cm nm . The R0 for Tb -DTPA-cs124-BMPH and

Alexa 546 combination in this case was 54.2 Å (refer to Table.1), which was similar to the value mentioned (55 Å) for an equivalent donor-acceptor pair (Posson & Selvin,

2008).

TMRM MTSR Alexa 546 Selvin, Richardson Posson Calculated Calculated Calculated 2002 et al., 2006 et al., 2008 J 3.80 x 1015 4.42 x 1015 N/A 3.68 x 1014 N/A 3.56 x 1015 (M-1cm-1nm4)

R0 (Å) 57.0 56.2 60.0 37.2 55.0 54.2

Table 1. J and R0 values for donor and acceptor pairs.

Shaded values highlight J and R0 derived from spectral measurements, and subsequently used for distance calculations. Donor in all cases was Tb3+-DTPA-cs124-BMPH. Known values from literature are also presented for comparison.

Figure 16. Normalized spectral data of the donor and acceptor.

Tb3+-DTPA-cs124-BMPH donor emission spectrum with A. TMRM, B. MTSR, and C. Alexa 546 acceptor absorbance and emission spectra. J is represented by the overlap between donor emission and acceptor absorbance spectra, where energy transfer occurs to result in sensitized emission.

2.7. Data analysis

Data were analyzed using pClampfit 10.3 (Axon Instruments) and SigmaPlot 11

(Systat Software). All data are expressed as means ± S.E. n stands for the total number of cells recorded from for each experiment condition.

2.7.1. Conductance-voltage relationship

In order to describe steady-state conductance-voltage relationship, peak tail current amplitudes were plotted against the test voltages. Peak tail current amplitudes from individual data sets were normalized to the maximum value and fitted with a single

Boltzmann function:

y = 1 / (1 + exp(V1/2 –V) / k)

y is the normalized conductance. V1/2 is the half-activation potential while V is the test potential. k is the slope factor.

2.7.2. LRET signal lifetime decay

The donor-only luminescence decay produced by Tb3+ complexed with in-house synthesized DTPA-cs124-BMPH, as well as sensitized emission from hERG L520C C- less and G516C C-less dimer-of-dimers were best described by a double standard exponential function:

I(t) = Aslowexp(-t/τslow) + Afastexp(-t/τfast) + C

I(t) is the intensity at a given time, while t represents time. Aslow and Afast are the relative amplitudes of the fast and slow components. τslow and τfast are the time constants for the fast and slow components, and C is a constant.

The sensitized emission from Shaker IR C245V V363C and C245V A359C channels formed from monomers was best fit by a triple exponential function:

I(t)= Aslowexp(-t/τslow) + Amedexp(-t/τmed) + Afastexp(-t/τfast) + C

I(t) is the intensity at a given time. t stands for time. Aslow, Amed and Afast are the relative amplitudes of the fast, medium, and slow components. τslow, τmed, and τfast are the time constants for the fast, medium and slow components, while C is a constant.

2.7.3. Distance determination

The efficiency of energy transfer (E) was determined by comparing the donor- only lifetime (τDO) with the lifetime of the sensitized emission (τAD) and applying the relation:

E = 1 - τAD/τDO

Based on the dependence of E on the inverse sixth power of the distance between the donor and acceptor (R-6), the distance (R) between the two probes were computed based on the formula:

-1 1/6 R = R0 (E -1)

The calculated R0 value presented in Table.1 for the appropriate donor-acceptor pair was used for all distance computations.

Chapter 3.

LRET Experiment Approach and Results

3.1. hERG dimer-of-dimer approach

For examining the dynamics of the voltage sensor in VG ion channels in response to membrane voltage change, the simple, standard approach in spectroscopic studies has been to engineer a cysteine in the extracellular S3-S4 linker above the voltage sensor. The subsequent labeling of the site with a fluorophore equipped with a thiol-reactive moiety allowed for observed changes in fluorescence emission to be translated into descriptions of voltage sensor movement, many of which corroborated findings from functional and structural studies (Blunck, Starace, Correa, & Bezanilla,

2004; Cha & Bezanilla, 1997; Es-Salah-Lamoureux et al., 2010; Mannuzzu et al., 1996).

Such an approach was also adopted in the first report of the LRET application on

VG ion channels, when investigating the voltage-dependent distance change in the

Shaker channel voltage sensor (Cha et al., 1999). In this study, the Shaker channels were expressed as tetramers composed of monomers, each with an extracellular engineered cysteine on top of the S4. The caveat of having four labeling sites within a channel was that donor and acceptor probes would be arranged to offer two potential distance measurements: one between adjacent subunits (RA) and another diagonally across the pore (RD), (Fig.17A,B). While this may seem favourable in that additional

information on the channel architecture is provided, it also presents the difficulty of confidently associating the given parameters to the supposed distances.

In an attempt to simplify the interpretation of sensitized emission data, others investigating voltage-dependent S4-S5 linker reconfiguration in KvAP channels have considered generating their channels from dimers (Faure et al., 2012). It was previously demonstrated that dimers assemble in a manner such that identical monomers are at diagonal positions (Liu, Sompornpisut, & Perozo, 2001). A single cysteine per dimer then allows for specific binding of donor and acceptor diagonally across the pore from each other, outputting in a single distance measurement per channel (Fig.18A,B).

This approach is particularly attractive, since other LRET studies that have used a Tb3+-donor complex similar to the one reported here revealed that the donor-only lifetime decay has two components (τDO,slow and τDO,fast), where τDO,slow was attributed to the lanthanide luminescence decay and τDO,fast was deemed to be a non-specific effect of an unknown source (Heyduk & Heyduk, 1997; Richardson et al., 2006). Analysis of sensitized emissions from channels composed of monomers where two distance measurements are possible thus produces four potential components of decay, which introduces further complexity in distance determination (Fig.17C). Previous studies have described that only the slowest component of the sensitized emission deriving from

τDO,slow (τSE slow, from τDO slow in Fig.17C) can be reported with confidence. It was regarded unfeasible to differentiate the two middle components from one another, while the fastest component was too fast to resolve and partially lost in the transient that follows the laser excitation pulse (Richardson et al., 2006).

Figure 17. Multiple sensitized emission time constants result from channels composed of monomers.

A. A tetramer formed by monomers, each with an engineered cysteine, leads to four possible donor or acceptor binding sites. B. Two energy transfer distances between donor (filled) and acceptor (unfilled) arises in a homotetramer: a longer distance diagonally across the pore between two opposite monomers (RD), and a shorter distance between two adjacent monomers (RA). C. A flow chart of four possible sensitized emission components is presented, where the Tb3+ donor intrinsically has two components and there are two distances that are measured. Only the slowest sensitized emission component (τSE slow, from τDO, slow) can be used in confidence to compute RD, since the two middle components (τSE fast from τDO slow, and τSE slow, from τDO fast) cannot be distinguished and the fastest is lost in the transient part of the signal (adapted from Richardson et al. 2006).

Figure 18. Single distance is determined from channels composed of dimer-of- dimer.

A. A dimer is formed by two monomers linked by an intracellular loop. Only one of the monomers carries an extracellular cysteine as a donor or acceptor binding site, while the other is removed of extracellular cysteine. B. A channel from dimer-of-dimer results in only two possible binding sites within a channel, located diagonally across from each other. Only one combination of donor (filled) and acceptor (unfilled) arrangement will provide sensitized emission; energy transfer will not occur in channels labeled with donors only or acceptors only. C. A flowchart shows the two potential components of sensitized emission, due to two innate components of the Tb3+donor. The slow component refers to the distance across the pore (RD), while the fast component is non-specific.

These observations drove the decision to consider the dimer approach here for hERG channels, in order to measure diagonal donor-acceptor distance without a confounding adjacent pair distances (Fig.18C, see also method section 2.1.2 for hERG dimer construction), especially in expectance of two time constants from the donor used.

To ensure that the linkage of two subunits by their intracellular C and N-terminal ends do not affect the proper function of the assembled channel, the voltage-dependence of activation of hERG dimer-of-dimers (with G516C or L520C mutation in the S3-S4 linker of one of the subunits) were compared to that of the hERG wild-type (WT) and WT C- less (extracellular cysteine removed) homo-tetrameric channels.

Fig.19 shows that that the hERG dimer-of-dimers have retained their function that is phenotypically similar to those observed in the WT and WT C-less homotetramers, using the same voltage-dependent activation protocol in all cases. There were no concerning differences with respect to voltage-dependence and ion current kinetics, despite the covalent link between C and N terminus between two subunits, as well as a cysteine mutation introduced in the S3-S4 linker. Therefore, hERG L520C dimer-of-dimer and hERG G516C dimer-of-dimer channels were considered appropriate to be used in the following LRET experiments to resolve the diagonal distance across the pore between two voltage sensors.

Figure 19. hERG dimer-of-dimer channels demonstrate wild-type like function.

A. Ionic current traces were recorded from holding at -80 mV, followed by 4 s depolarizing steps up to +60 mV in 10 mV increments, and a -110 mV step to generate the peak tail currents (Voltage protocol shown in Fig.13). B. G-V relationships were constructed from peak tail current shown in B (See section 2.5.2).

3.2. Tb3+ donor-only measurements

3.2.1. Concentration dependence of Tb3+ emission

Chemical labeling procedures often entail application of an excess of fluorescent probes to ensure saturated labeling. However, this approach can also lead to unspecific labeling of both endogenous and exogenous proteins, interruption of the proper function of the channel in question, or unspecific incorporation of the label into the lipid membrane (Heyduk & Heyduk, 1997; Sandtner, Bezanilla, & Correa, 2007). In recognition of previous reports on the presence of a non-specific component within the

Tb3+-DTPA-cs124 donor luminescence decay (section 4.1) and in the attempt to minimize such an undesirable property, several concentrations of the donor were tested and their characteristics were noted.

Fig.20 shows the concentration-dependent properties of Tb3+-DTPA-cs124-

BMPH donor emission when labelling hERG L520C dimer-of-dimer channels. As expected, Tb3+ luminescence decayed with a bi-exponential time course. The slow component (τslow, 1.07 ± 0.02 ms compared to τfast, 0.15 ± 0.02 ms) was identified as the luminescence originating from the DTPA-cs124-BMPH bound Tb3+ ion, comparable to the reported value (τslow, 1.1 ms) from the same synthesized donor complex (Faure et al.,

2012). The optimal concentration was determined based on the balance between emission intensity, prevalence of the slow decay component, and proper channel function reflected in the ionic current recordings. It is notable that the slow time constant was fairly insensitive to changes in the donor concentration (Fig.20 A). However, at the maximum concentration, the favourable high luminescence intensity and fractional

Figure 20. Concentration-dependent properties of Tb3+-DTPA-cs124-BMPH.

A. Time constant of Tb3+ luminescence decay as a function of Tb3+ donor concentration. The time constant of donor luminescence decay is a property of the donor complex. In this case of Tb3+-DTPA-cs124-BMPH, the slow time constant of 1.07 ± 0.02 ms was identified as the luminescence decay time constant from the donor complex, as previously reported. The time constant was insensitive to the donor concentration until higher concentrations. B. Fractional amplitude of slow time constant as a function of Tb3+ donor concentration. Luminescence decayed with a bi-exponential time course. The optimal donor concentration was one in which the signal was carried mostly by the 1.07 ms component. C. Luminescence intensity as a function of Tb3+ donor concentration. Luminescence intensity increased with increasing amount of donor. D. Raw traces of luminescence intensity for various concentrations of the donor. Data in A, B, and C are shown as mean ± SE. n= 3 to 5.

contribution of the slow component (Fig. 20B to D) were accompanied by a slight deviation of the slow time constant from its characteristic value (Fig.20A, 1.0X dilution factor). This has also been observed in another report of concentration-dependent Tb3+ donor properties, where such a decrease in τslow, along with the non-saturating aspect of

Tb3+ emission intensity, were attributed to the unspecific effect of donor (Sandtner et al.,

2007). Considering all factors above, Tb3+-DTPA-cs124-BMPH reconstituted to 25 µM

(0.5X dilution factor) was chosen as the optimal concentration to use in labeling.

Table.2 summarizes the slow time constant, relative amplitude of the slow time constant and emission intensity obtained from labeling each site of interest with donor only. The similarity in the donor properties regardless of difference in labeling site suggests that the observed properties are intrinsic to the donor complex (depending on the lanthanide, antenna, and chelate used), and independent of the labeling environment.

Emission Labeling Site n Τ (ms) A slow slow intensity (V)

hERG L520C 24 1.07 ± 0.02 0.36 ± 0.02 4.46 ± 0.53

hERG G516C 4 1.04 ± 0.02 0.26 ± 0.03 4.55 ± 0.98

Table 2. Tb3+-DTPA-cs124-BMPH properties at each labeling site.

Values were obtained from labeling with the determined optimal donor concentration, 25 µM. Data are presented as mean ± SE.

3.2.2. Specificity of the Tb3+ donor complex signal

In light of the potential non-specific effects that may arise from donor complex labeling and the fact that the synthesized chelate complex was not purified, specificity of the observed donor-only signal was evaluated in various ways.

Previous work has raised concerns of contaminating signals from excess Tb3+ ion interactions with channels that cause the observed non-specific fast component in the donor luminescence (Hyde et al., 2012; Sandtner et al., 2007). In investigating this in our system, Fig.21A presents the extent of contamination in the donor-only signal when the channels exposed to TbCl3 or chelate complex only. This was evaluated by assessing carboystyril124 emission, since it is excited at 337 nm and emits in the 400 nm range (peak at 410 nm). In both chelate only and chelate with TbCl3 cases, the resultant signals were substantially smaller (0.60 ± 0.05 V, n=4 each) than the luminescence decay obtained from Tb3+ complexed with DTPA-cs124-BMPH (at least

10-fold difference). Moreover, such outcomes also suggest the effectiveness of the donor chelate, where considerable Tb3+ luminescent emission is only observed when the lanthanide ion and chelate complex are coordinated together.

In order to properly use Tb3+-DTPA-cs124-BMPH for distance measurements, it is important to confirm specificity of labeling. To achieve this goal, all labeled cells

(whether donor-only or donor-acceptor) underwent a pre-labelling process with ß- maleimidopropinic acid (B-MPA, Methods section 2.4.1) to block out endogenous cysteines in the extracellular surface of the membrane (Gonzalez, Rambhadran, Du, &

Jayaraman, 2008; Hyde et al., 2012; Posson et al., 2005; Posson & Selvin, 2008). The specificity of the donor complex labeling was implied by comparing cells expressing

hERG WT C-less channels (control condition where all extracellular cysteines are removed from the exogenous channel in question) with and without B-MPA pre- treatment (Fig.21B). The suppression of donor-only signal upon B-MPA application suggests effectiveness of the treatment and targeting of endogenous cysteines and thus increased likelihood that subsequently introduced labels are specifically binding to the engineered cysteine.

Finally, it is crucial to ensure that there is no donor luminescence impinging into the sensitized emission and thereby influencing distance interpretation. Fig.21C indicates that there is a proper spectral separation introduced by the emission filters used, between the donor-only emission and the sensitized emission. Donor-only signal amplitude recorded using a series of 565 bandpass and longpass emission filters (the sensitized emission recording condition, where Tb3+ emission is minimal (refer to Fig.15), was negligible (0.20 ± 0.05 V, n=10), giving confidence that there is minimal spillover of donor luminescence into the sensitized emission. In all subsequent sensitized emissions recorded, the signals were at least 40 times larger than the donor-only signal obtained with 565 emission filters.

Figure 21. Specificity of Tb3+-DTPA-cs124-BMPH as a donor.

A. Signals obtained with 485 emission filters from channels labeled with Tb3+ ions only or chelate complex only. There is a negligible effect of the two individual components on donor-only (DO) signal. B. Effect of pre-treatment with B- MPA on addressing the endogenous cysteines and enabling specific labeling of the donor complex is shown, where the hERG WT C-less B-MPA treated condition represents background signal. C. Donor-only signal taken with a 485 emission filter compared with donor-only signal from 565 emission filters. Clear spectral separation between the donor and sensitized emission is indicated. Presented traces are from cells expressing hERG L520C dimer-of-dimer. Similar findings were observed with other channels as well (data not shown).

3.3. RD at the resting membrane potential

With introduction of the acceptor fluorophore along with the lanthanide donor, inter-subunit distance across the pore (RD) from two sites at the top of the S4 at the resting membrane potential was examined. First observations were made in Shaker IR channels (A359 and V363) to compare to the reported LRET derived distance measurements, and then in hERG channels (L520 and G516).

A summary of resultant distances from the sensitized emissions is presented in

Table. 3. Measurements made from Shaker channel sites (47.6 ± 0.1 and 47.7 ± 0.2 Å) were similar to the reported values (49 and 45 Å for A359 and V363, respectively), offering confidence on the ability to resolve targeted distances (Cha et al., 1999; Posson

& Selvin, 2008).

Fig.22 presents representative sensitized emission traces obtained from labeling diagonally located sites on top of hERG dimer-of-dimer channel voltage sensors. In contrast to the donor-only signal recorded from the same location, the sensitized emission displayed a much faster decay with distinct multiple components. On average, the sensitized emission signal was at least 10 times greater than the background signal.

From both L520 and G516, RD values were found to be ~10 Å larger than analogous measurements made in the Shaker channels (Table.3). In addition to the implied difference from Shaker channel architecture, the measurements in hERG channels were in better agreement with the diagonal distances acquired from the extracellular part of KvAP and NaChBac S4 segments using LRET, which were approximately 60 Å (Richardson et al., 2006).

To further ensure that the distances measured were independent of the specific donor-acceptor pair, distances were also assessed with other acceptor fluorophores available, MTSR and Alexa 546. RD from using Alexa 546 supported the distance computed from using TMRM. The difference of a few angstroms between the two measurements was as previously observed and expected when the degree of error in distance determination was considered. The sources of error in contributing to distance measurements in LRET are further discussed in section 4.2 (Faure et al., 2012;

Richardson et al., 2006).

It is notable that the distance computed when using MTSR was drastically smaller than others. However, such a discrepancy could be associated with an underestimated MTSR R0 value (refer to Table.1) for the following reasons. The known

R0 value of MTSR is 60 Å, comparable to that of TMRM and Alexa 546; this is expected since MTSR shares similar spectral properties (and thereby J value) as the two other organic fluorophores (Fig.16). In this case, the derived J value for MTSR was 10 fold smaller due to the 10 fold difference reflected in the calculated extinction coefficient. All acceptor absorbance measurements and the calculation for the extinction coefficient were based on the assumption that the working stock was reconstituted to a final concentration of 25 µM. In light of the fact that MTSR has a shorter half-life than both

TMRM and Alexa 546, it is conceivable that the MTSR stock used had partially degraded and thus was comprised of a much lower concentration than the assumed 25 µM.

Additionally, the efficiency of energy transfer was comparable in all three donor-acceptor cases: 0.51 ± 0.01 with TMRM, 0.48 ± 0.01 with MTSR, and 0.53 ± 0.02 with Alexa 546.

Hence, if the actual MTSR concentration was determined and corrected for, it is most

likely that distance measurements from using MTSR as an acceptor would better align with those of other donor-acceptor pairs.

Acceptor

Labeling Site TMRM MTSR Alexa 546 Reported value

hERG L520C 56.0 ± 0.4 (20) 37.7 ± 0.2 (5) 53.1 ± 0.7 (4)

hERG G516C 54.8 ± 0.3 (8)

Shaker A359C 47.6 ± 0.1 (4) 49 (Posson et al., 2008)

Shaker V363C 47.7 ± 0.2 (8) 45 (Cha et al., 1999)

Table 3. RD from diagonally located voltage sensors in hERG and Shaker channels at resting membrane potential.

Mean distance across the pore from each labeling site is reported, with sample size in brackets. The known value from analogous measurements in literature is presented for Shaker channel for comparison.

Figure 22. LRET raw traces from the top of S4 in hERG dimer-of-dimer channels.

Donor-only (DO), sensitized emission (SE), and background signals from hERG L520C (A) and G516C (B) obtained from labeling with Tb3+-DTPA-cs124-BMPH and TMRM are shown. SE displayed much faster decay and distinct multiple components compared to DO emission. Slow time constants for DO and SE are indicated, which were used to compute the distance between donor-acceptor pair.

3.4. Voltage-dependent change in distance

An increasing number of spectroscopic studies have provided insight into the voltage-dependent reconfiguration of the voltage sensor by noting the qualitative change in fluorescence emission from the fluorophore anchored to the top of the S4 (Cha &

Bezanilla, 1997; Es-Salah-Lamoureux et al., 2010; Mannuzzu et al., 1996; Smith &

Yellen, 2002). In the case of LRET, a change in the time course of the sensitized emission decay can be translated to a dynamic distance change associated with gating at an Ångstrom level (Cha et al., 1999; Hyde et al., 2012; Posson et al., 2005; Posson &

Selvin, 2008).

As the first step to observe voltage-dependent change in the hERG channel voltage sensor movement, the lifetime decay of sensitized emission recorded from 2 s

(approximate steady-state) voltage steps to –80 mV and +60 mV were compared. The same voltage protocol was also applied to donor-only labeled cells, to assess whether there was any voltage-imposed modification of donor characteristics that might confound the observed distance change. Donor lifetime in the absence of an acceptor remained the same regardless of the voltage (paired t-test, p<0.001; refer to overlapping donor traces in Fig.23), and such insensitivity to voltage was observed at all sites considered for labeling, as previously noted in Shaker potassium channels (Cha et al., 1999). This indicated that there was no significant change in the environment of the terbium ion held within the chelate complex upon imposed voltage change.

Table.4 is a summary that compares RD values obtained from holding the channels at a hyperpolarizing potential and then at a depolarizing potential. In all cases,

there was no voltage-dependent distance change observed (Fig.23). Such an outcome was perhaps not unexpected given the nature of the donor-acceptor arrangement on the channels, which is designed to most aptly detect radial movements of the voltage sensor. Extensive investigations on the voltage-dependent S4 reconfiguration in Shaker channels using LRET have concluded that there is little to no radial displacement of the voltage sensor in response to voltage change (Hyde et al., 2012; Posson & Selvin,

2008). Only a 1.5 Å change was detected at most when channels were exposed to -150 mV, followed by a +25 mV step, and only at a couple sites in the S3-S4 linker demonstrated such a change. It should be mentioned that Shaker A359C was one such site that reported an RA (distance between voltage sensors in adjacent subunits) change from 35.6 ± 0.2 Å (-150 mV, approximately 50 Å in terms of RD) to 34.1 ± 0.2 Å (+60 mV,

48.2 Å in terms of RD) (Posson & Selvin, 2008). By contrast, Shaker V363C was a site at the top of the S4 that did not report a voltage-dependent change in radial distance (Cha et al., 1999).

In consideration of the above, it is possible that hERG L520C and G516C are sites that do not portray voltage-dependent radial reconfiguration of the hERG voltage sensor. This is supported by the relative location of L520 and G516 residues in the S3-

S4 linker of hERG channels (Fig.24). The top of S4 is alpha helical in structure, where residues are splayed out 100° from each other. Such an arrangement places L520 and

G516 residues on the same side of the alpha helix. Therefore, when it comes to reporting measurements across the pore and conferring tilting motion of the voltage sensor, both sites can be predicted to output similar distances. This warrants other residues in the S3-S4 linker on the opposing side of the alpha helix to be used as a

labeling site (S517, E518 and E519); further discussion regarding this is presented in section4.2.3.

Figure 23. LRET raw traces from hERG L520C dimer-of-dimers at hyperpolarizing and depolarizing potentials.

Donor-only and sensitized emission signals from +60 mV are overlaid on top of the signals from -80 mV. At both potentials, similar time constants were reported. The same observation was made from hERG G516C labeling site.

Figure 24. Helical wheel of hERG channel S3-S4 linker.

Alpha helical structure of the S3-S4 linker is represented as a helical wheel, with relative position of G516 and L520 shown. As a turn occurs in an alpha helix every 3.6 residues, G516 and L520 are positioned on the same side of the helix, vertically 5.4 Å apart from one another. The diameter of an alpha helix (with length of the protruding residue side chains averaged) about 12 Å.

Acceptor

TMRM MTSR Alexa 546

Labeling Hyperpolarizing Depolarizing Hyperpolarizing Depolarizing Hyperpolarizing Depolarizing Site potential potential potential potential potential potential

hERG 56.6 ± 0.9 (6) 56.4 ± 0.7 (6) 38.0 ± 0.3 (5) 38.1 ± 0.2 (5) 53.5 ± 0.6 (4) 53.6 ± 0.6 (4) L520C

hERG 56.0 ± 0.8 (3) 56.2 ± 0.6 (3) G516C

Shaker 48.6 ± 0.4 (4) 48.6 ± 0.4 (4) A359C

Shaker 47.5 ± 0.2 (4) 47.5 ± 0.2 (4) V363C

Table 4. RD from diagonally located voltage sensors in hERG and Shaker channels at hyperpolarizing and depolarizing membrane potential.

Sensitized emission recordings from hERG channels were obtained after 2 s at -80 mV or +60 mV. Sensitized emission recordings from Shaker channels were obtained after 50 ms at -150 or -120 mV and +25 or +50 mV for A359C and V363C, respectively, as per reported protocols used in the literature for each of those sites (Cha et al., 1999; Posson & Selvin, 2008).

Chapter 4.

Discussion

As the first known application of LRET on hERG channels, the work here outlines: i) steps taken to confirm proper set-up and function of the technique and its components, and ii) a deliberate experimental design to measure static and dynamic distances in hERG channels, and iii) an attempt to validate and confirm distance measurements by considering sources of error and alternate interpretations of the data collected.

4.1. Examining donor-only properties

Lanthanide-based donor probes have fixed properties that can be well characterized. This is a consequence of entrapping the lanthanide ion in a protective cage-like chelate, which renders it insensitive to the local change in environment (Thibon

& Pierre, 2009; Vuojola & Soukka, 2014). It is helpful to take advantage of this fact and try to fully understand the nature of each of lanthanide donor characteristics, as it would facilitate the interpretation of sensitized emission data.

Previous studies have reported donor-only lifetime decays to have three exponential components, whiles others reported only two (Hyde et al., 2012; Posson et al., 2005). In all cases, the slowest component that was in the millisecond range was identified as the donor lifetime, as expected based on the known property of lanthanide

ions (section 1.3.3). The faster time constant of unknown source, however, varies in the literature and ranges from 100 to 300 µs (Hyde et al., 2012; Posson et al., 2005). Donor- only measurements presented here carried a similar fast time constant of 150 ± 20 µs, despite the effort to reduce any non-specific labeling. Although this fast component is largely dismissed as being a non-specific effect by many, it still may contribute a systematic error that causes slight underestimation of distances (Heyduk & Heyduk,

1997; Posson et al., 2005).

4.2. Interpreting sensitized emission data

4.2.1. Evaluation of the time constants

The analysis approach used to quantify sensitized emission signal was derived based on i) the known number of intrinsic components of the donor signal, and ii) the expected number of distance measurements. This was presented as flow charts in Fig.

17 and 18. Sensitized emission obtained from hERG dimer-of-dimers was well described by a double exponential as expected. The slow component was used to compute the diagonal distance across the pore, while the fast component (150 ± 20 µs) was in good agreement with the notion that it had originated from the non-specific donor fast component.

By contrast, interpretation of the Shaker channel sensitized emission data was not as straightforward, as expected from a channel formed from monomers. According to

Fig.17, four components were anticipated. However, the emission decay was best described by a triple exponential, and this was accepted based on previous reports that the two middle components cannot be distinguished from one another (Richardson et al.,

2006). The resultant three components were: 1.05 ± 0.04 ms, 0.29 ± 0.01 ms, and 0.11 ±

0.01 ms with relative amplitudes of 1.4 ± 0.1 %, 31.8 ± 3.5 % and 66.7 ± 4% (Shaker

A359V data).

Two approaches were taken in interpreting the three parameters. One was to simply apply the rationale presented in Fig.17 and associate the slowest component,

1.05 ms, with RD. This outputted a distance of 109 Å between two diagonally opposing voltage sensors, which exceeds even the homologous measurements made in the KvAP crystal structure (refer to section 1.2.2 and 4.2.2) (Bezanilla, 2007).

The second approach was based on an observation that the slowest time constant is akin to the slow time constant from donor-only measurements. Thus, it was postulated that this slow component may be a contaminating signal from extraneous donors that had not participated in energy transfer. This idea was further supported by the fact that there may be a small but recognizable amount of emission from Tb3+ starting from 580 nm, and the upper limit for the bandpass emission filter used in the current set-up is 585 nm. A test to confirm this would be to insert a narrower bandpass filter at 565 nm or an additional short pass filter set at 575 nm. Upon assuming that the slowest component is a donor bleed through due to inadequate spectral segregation, the next slowest time constant was treated as the one reflecting the distance across the pore. As noted in section 3.3, this distance closely aligned with the reported distance from the same site and arrangement. Taken together, complexity that is clearly introduced when interpreting sensitized emission data from homotetramers, lends further support for the dimer-of-dimer approach used here in hERG channels.

4.2.2. Comparison of LRET distance measurements to crystal structures

In a number of LRET studies involving VG ion channels, verification for the calculated distances was obtained by taking distance measurements from homologous residues in the available VG ion channel crystal structures (Cha et al., 1999; Faure et al.,

2012). For hERG channels, there is no crystal structure that has been resolved for a direct comparison. Instead, measurements from an analogous position in a resolved mammalian VG channel (Kv1.2) crystal structure (S289 in 3LUT) resulted in an 83 Å distance between opposing voltage sensors across the pore (X. Chen, Wang, Ni, & Ma,

2010). This was in contrast to 56 Å reported here from hERG L520 and G516, but such an outcome was not much of a surprise due to a similar observation previously made (98

Å in that case) in the crystal structure of KvAP channels (Bezanilla, 2007; Jiang et al.,

2003). Furthermore, the low sequence homology between hERG and Kv1.2 channels limits the strength with which conclusions can be made regarding the hERG channel structure.

Those who fervently dismiss the S4 radial distance measurements taken from the crystal structures argue on the basis that such a gross geographical separation of the voltage sensor from the rest of the channel is unrepresentative of the channel’s native form. This is due to: i) the unrealistically high energy cost that is implied from such a structural conformation requiring drastic movement of charges through the lipid bilayer, and ii) non-physiological condition that is imposed during the protein crystallization process (Bezanilla, 2007). More convincingly, distances derived from crystal structures can be questioned based on experimental evidence from techniques that are regarded as “spectroscopic rulers.” For example, studies using LRET, FRET, or tethered TEA

derivatives all report a unanimous distance of 50 Å from voltage sensors across the pore

(Blaustein, Cole, Williams, & Miller, 2000; Cha et al., 1999; Glauner, Mannuzzu, Gandhi,

& Isacoff, 1999; Posson et al., 2005; Posson & Selvin, 2008). All of these results, including the one reported here, collectively suggest that the S4 segment is not extended into the bilayer, but is against the bulk of the channel protein.

4.2.3. Considerations for measured distances

Although LRET is considered to produce more reliable distance measurements than FRET (refer to advantages of LRET in section 1.3.2), there are several caveats regarding the determined distances.

Movement of the donor and acceptor probe

The resolution of LRET distances can be limited by the movement of the donor and acceptor, as it is plausible for probes to wobble about their linker to a position that can introduce error into the distance measurement (Cha et al., 1999). This is dependent on the length and flexibility of the linker, as well as the environment that the protein is in.

As for the maleimide linker used in this report, the arm length is known to be about 5.9 Å.

Previous studies have shown that the flexibility of maleimide linkers inserts minimal error

(< 2Å) (Cha et al., 1999). Nonetheless, it should still be noted that the position of both the donor and acceptor with respect to the labeled residue is unknown at any given point.

Temporal resolution

Temporal resolution may also be limited, due to the long lifetime decay

(approximately 5 ms) that is characteristic of LRET. Protein conformation changes that

occur on a similar millisecond range would complicate lifetime decay measurement, and the temporal resolution would be lost (Blunck, 2015). To avoid this, it is of best interest to take all LRET measurements at steady state of the conformation of interest, as it has been done in this report as well as in previous work (Cha et al., 1999; Posson et al.,

2005; Posson & Selvin, 2008).

Because of the long lifetime decay of the sensitized emission in LRET, it is implied that the technique is most appropriate for studying slow conformation changes.

This should be noted when considering application of LRET on hERG channels, where activation and deactivation take place over several hundred milliseconds and may be accompanied by the slow movement of the voltage sensor (Piper et al., 2003; Smith &

Yellen, 2002). The converse situation is illustrated with a recent study that used LRET to explore the existence of an alpha helix to a 310 helix transition of the S4 segment in response to a membrane voltage change (Kubota, Lacroix, Bezanilla, & Correa, 2014).

Donor and acceptor probes were covalently linked to the N-terminal and C-terminal end of the S4 segment of Ci-VSP (a voltage-dependent phosphatase), respectively. For a 19 residue long S4 segment, the expected length with a full alpha helical conformation was

28.5 Å (19 residues x 1.5 Å rise per residue) and the expected length with a full 310 helical conformation was 38 Å (19 residues x 2 Å rise per residue). Therefore, a change in vertical distance detected in that range upon depolarization can infer transition in the

S4 secondary structure. However, no such change was found, most likely due to the sterically unstable nature and thereby transient existence (nanoseconds) of the 310 helical structure (Kubota et al., 2014; Vieira-Pires & Morais-Cabral, 2010).

Measurements from multiple sites

Finally, there have been observations of inconsistency in the degree of distance change measured from various labeling sites, despite the fact that they are all located on the same part of the protein. This was the case for measurements taken from S3-S4 linker of the Shaker potassium channel, where a range of voltage-dependent distance changes was perceived from a number of sites along the linker (Posson et al., 2005;

Posson & Selvin, 2008). A simple explanation would be that the protein structure is undergoing a complex reconfiguration process, and is depicted as variability in the probe movement. Another possibility is that certain sites may be better than others at tracking structural reconfiguration. Select residues that move into a constrained environment as a result of protein conformational change may experience forced arrangement of the attached probe relative to the protein (Posson & Selvin, 2008). To be able to determine which explanation is more applicable for a given scenario, it is recommended to take measurements from multiple sites from the structure of interest.

In the context of examining radial movement of hERG channel voltage sensor, additional residues in the S3-S4 linker (S517, E518, and E519) should be considered to establish whether the minimal radial distance change observed in response to voltage is due to a residue specific effect. Since the hERG channel S3-S4 linker is alpha helical, these additional measurements can also reveal if there is a rotation of the structure, as was the case for three consecutive sites in the Shaker channel S3-S4 linker (Cha et al.,

1999).

4.3. Future directions

4.3.1. Other donor-acceptor pairs

In addition to the recommendations above, further verification of distance measurements should be obtained through other donor-acceptor pairs. This has been considered in the current study by using TMRM, MTSR and Alexa 546; however, all the acceptors had similar spectral properties and thus similar R0 values. To truly assess that the derived distances are independent of the specific donor-acceptor pair, they should be tested with donor-acceptor combinations that give different R0 values (Blunck, 2015;

Faure et al., 2012). This is because the transfer efficiency is a steep function of distance, and only distances within the range 0.5R0< R < 1.5R0 can be reliably measured

(Sandtner et al., 2007). As such, data from diverse donor-acceptor pairs (e.g. Tb3+-

DTPA-cs124-BMPH and fluorescein R0= 42.7 Å) is necessary for confidence in the measured distance (Faure et al., 2012).

4.3.2. Detecting vertical distance

The present experimental approach sets the stage for making initial measurements from the channel with LRET. However, this arrangement can only offer information on radial displacement, which has been shown in numerous studies to be minimal (Bezanilla, 2002; Cha et al., 1999; Posson et al., 2005; Posson & Selvin, 2008;

Starace & Bezanilla, 2004). Upon confirming that this is true via measurements from more sites in the S3-S4 linker as well as with other acceptors, investigation of other types of displacement of the voltage sensor can be considered. Namely, this was achieved in Shaker potassium channels by labeling the top of the S4 with the donor and applying acceptor-labeled charybdotoxin or agitoxin-2 that blocked at the pore of the

channel, which revealed voltage-dependent vertical displacement (Posson et al., 2005;

Posson & Selvin, 2008). A similar approach could be taken with hERG channels using hERG toxins (Jimenez-Vargas, Restano-Cassulini, & Possani, 2012).

4.4. Significance of the study

Despite the technical difficulties and limitations associated with LRET, the technique offers an unparalleled opportunity to examine structural reconfigurations at an atomic level. This report offers a step-by-step tested methodology on applying LRET to examine hERG channel voltage sensor dynamics. The culmination of this process is the first distance measurement from voltage sensors across the hERG channel pore.

Further application of the outlined method has the potential to shed light on the molecular basis of physiologically significant slow activation and deactivation in hERG channels. Recently, there has been a finding that the extracellular S3-S4 linker in Kv channels may be involved in modulating the affinity for gambierol, a type of polyether toxin (Kopljar et al., 2016). As hERG channels are notorious for their interactions with drugs, better understanding of conformation changes in the S3-S4 linker through LRET may also contribute to the pharmaceutical field.

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Appendix A.

Side-products from DTPA-cs124-BMPH synthesis

Appendix A. Structure and molecular weight of potential side-products formed from DTPA-cs124-BMPH. Schematics were generated using ChemDraw (courtesy of Hong Yee Tan).

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Appendix B.

Mass spectrum of DTPA-cs124-BMPH synthesis sample

Appendix B. Mass spectrum of a sample from DTPA-cs124-BMPH synthesis. The major competing products identified were cs124-DTPA-cs124, DTPA- cs124-BMPH, and BMPH-DTPA-BMPH.

102