LUMINESCENCE RESONANCE ENERGY TRANSFER STUDIES OF THE SHAKER K + VOLTAGE-GATED ION CHANNEL DAVID JOHN POSSON DISSERTATION

Transcription

1 LUMINESCENCE RESONANCE ENERGY TRANSFER STUDIES OF THE SHAKER K + VOLTAGE-GATED ION CHANNEL BY DAVID JOHN POSSON B.S., University of Cincinnati, 1997 DISSERTATION Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Physics in the Graduate College of the University of Illinois at Urbana-Champaign, 2005 Urbana, Illinois

2 LUMINESCENCE RESONANCE ENERGY TRANSFER STUDIES OF THE SHAKER K + VOLTAGE-GATED ION CHANNEL David John Posson, Ph.D. Department of Physics University of Illinois at Urbana-Champaign, 2005 Paul R. Selvin, Advisor Members of the superfamily of voltage-gated ion channels are the molecular components underlying electrical excitability in nerves and muscle. Voltage-gated channels allow the selective flow of ions across the hydrophobic lipid bilayer of a cell, opening and closing in response to changes in the voltage across the membrane. The Shaker K + channel is a standard model system for studying the structure function relationships in this important class of ion channels. Voltage sensing is known to involve a highly charged segment of the channel called S4. When a channel opens it moves some of these S4 charges across the membrane electric field. The movement of four S4s, one from each identical subunit of Shaker, is coupled to the gate which opens and closes the pore. Therefore, a central question for understanding the functionality of these proteins is; how exactly does S4 move? Recently, the first crystal structure for a voltage-gated K + channel was solved. This structure, of the KvAP channel, led the authors to propose a new model of S4 movement. This new model, called the paddle model, hypothesized a large translational motion of S4 across most of the lipid bilayer thickness, a view that has been very controversial. In this study, we examine the conformational movements associated with the S4 segment during voltage sensing. We use a technique called Lanthanide Resonance Energy Transfer (LRET), which gives an accurate distance measurement between two positions on the ion channel. We use two different configurations for LRET on Shaker. The first measures distances between the four identical S4 amino-acid sites on the homotetrameric K + channel. The second measures distances between S4 sites and a scorpion toxin bound to the symmetric axis of the channel just above the pore. These LRET studies argue strongly against the paddle model of voltage sensing, and demonstrate that the physical movements of the S4 segments of Shaker K + channels are quite small. iii

3 Acknowledgements So many people helped me out at every stage of grad school and in so many different ways. I thank them all for their generosity. My parents and sisters have been my base of support for 30 years and counting. Paul Selvin taught me everything I know about LRET and how to do fluorescence experiments. I thank him for passing on his techniques and skills and most of all for believing in me during times when I was not the most ideal of grad students and when I was just down right flakey. Francisco (Pancho) Bezanilla and Dorine Starace taught me so much about the Shaker channel and about electrophysiology in general. Pancho s knowledge is inspirational and Dorine provided support both professional and personal. Without them I could not have succeeded. Chris Miller was instrumental with his scorpion toxin preparations. Without his help my Ph.D. would have turned out a bit more scrappy than it did. I thank him most for his heady encouragement that buoyed me along through the final stages of this work. I thank Pinghua Ge for his skill at preparing lanthanide chelates. Without his synthetic abilities, none of my experiments would have been possible. I would also like to thank Ming Xiao for determining the chelate s quantum yield and Jeff Reifenberger for measuring the anisotropy. Greg Snyder provided initial instruction in voltage clamping and oocyte work. Both Greg and his wife, Tania, were wonderful colleagues in the Selvin lab and I miss them. iv

4 Anne Gershenson taught me basic mutagenesis and provided friendship with a needed dose of mentorship. For her crucial help, I am deeply grateful. Michel Bellini passed on his knowledge of Xenopus, including the surgical procedure. Bob Gump and his staff at the Morrill hall animal facility took great care of our frogs. Sara Chalifoux and Lisa Klodnicki performed numerous frog surgeries and oocyte preps. Tatyana Lawrecki helped with surgeries and oocyte preps through the most productive period these two years past. For her loyalty and the unwavering quality of her efforts I am very grateful. I thank the Aldrich lab for providing the ILT-Shaker plasmid. Benoit Roux kindly combined the coordinates for two of his models, the Shaker open state and the agitoxin-shaker complex. I thank him for his general enthusiasm for my experiments. Dane Sievers provided silicon wafers and greatly assisted with the oxide growth for the silicon quenching project. Other members of the Selvin lab have provided friendship and support - Evan Graves, Ahmet Yildiz, Comert Kural, Hyokeun Park, Hamza Balci, Sheyum Syed, & Erdal Toprak. Jeff Reifenberger has been a good roommate, lab colleague, and a great friend through some tough times. v

5 I have gratefully received personal support from Scott Stewart, Paul Melby, Patrick Hentges, Trevor Vickey, & Mary Upton. Patrick showed me what personal courage really is all about. I acknowledge the financial support for this work with NSF MCB , Carver Foundation, and Cottrell funds of the research corp. CS0706 vi

6 Table of Contents Chapters 1 Introduction Ion channels the cartoon picture Classical membrane topology of the Shaker K + channel How does an ion channel handle its charge? K + channels the atomic picture The KscA structure a closed K + channel The MthK structure an open K + channel The MscS structure a slightly voltage sensitive, non-selective mechanosensitive channel The KvAP structure a full-fledged, highly voltage-dependent channel MacKinnon goes to Stockholm Electrophysiology of the Shaker K + Channel The Xenopus laevis oocyte expression system The two-electrode voltage clamp Ionic currents Gating currents The ILT-Shaker phenotype Fluorescence Spectroscopy Methods Introduction to fluorescence Resonance Energy Transfer (RET) theory Lanthanide Resonance Energy Transfer (LRET) LRET meets the voltage clamp Instrumentation...65 vii

7 4 LRET Part I: The ILT-Shaker Channel What can measurements on the ILT channel tell us? Initial LRET results on the ILT channel LRET Part II: Shaker with Scorpion Toxin Ion channels are toxin receptors LRET configuration using acceptor labeled toxin putting the paddle model to the test LRET results S3b, S3-S4 linker, and S Comparison to a model for Shaker Conclusions Future experiments...89 Appendices A Molecular Biology A.1 Shaker constructs...92 A.2 Primers...96 A.3 Mutagenesis A.4 mrna synthesis A.5 Toxin biochemistry B Animal Use Protocol C Xenopus Oocyte Preparation D Electrophysiology Solutions E Silicon Quenching Unbinding Bioassays References Vita viii

8 Chapter 1 Introduction 1.1 Ion channels the cartoon picture Ion channels are protein pores that allow ions to flow across the otherwise impermeant cell membrane. Most channels are highly selective for a single ion such as Na +, K +, Ca 2+, or Cl -. Well-conserved pore structures, called selectivity filters, confer this specificity. Another essential structural element, called the gate, opens to permit the flow of ions or closes to block the pore. Ion channels usually couple their gates to some external influence such as ligand binding ( ligand-gated channels), membrane voltage ( voltage-gated channels), or mechanical force ( mechanosensitive channels). Figure 1.1 shows cartoon representations of these ion channel types. All animal, plant, and bacterial cells have membranes in order to keep inside stuff separate from outside stuff. Ionic concentration gradients are maintained across their plasma membranes and a negative voltage is present across the membrane when the cell is at rest [1]. It is customary to express membrane voltage as V intracellular V extracellular, and let V extracellular = 0. Animal cells typically have membrane voltages between 60 mv and 100 mv at rest. Therefore, ionic currents across cell membranes depend on the electrochemical gradients as well as the presence or absence of open ion channels. Table 1.1 gives the measured concentrations of the biologically relevant ions in mammalian skeletal muscle. The Equilibrium Potential, the voltage at which the concentration gradient exactly balances the electrical gradient, is given by equation 1.1. Z n is the valence, [N] out is the extracellular concentration, and [N] in is the intracellular concentration of ion n. Equation 1.1 is known as the Nernst equation. E n RT [ N] out = ln (1.1) ZnF [ N] in 1

9 Closed Open a L L L L Neurotransmitter gated b L L L L Second messenger gated c Voltagegated V = -100 mv V = 0 mv d Mechanical Force gated Figure 1.1 All ion channels have gates (red doors) that open and close the channel. Different types of channels open in response to different stimuli. a. Neurotransmitter-gated channels bind diffusional ligands from the external solution in order to transfer electric excitation from one neuron to another. b. Some ligand-gated channels open in response to intracellular second messengers such as Ca 2+. c. Voltage-gated channels have charged protein segments that move in response to membrane voltage changes thereby opening the gate. d. Mechanosensitive channels open when mechanical membrane stress or bending is applied. Table 1.1 Free Ion Concentrations and Equilibrium Potentials for Mammalian Skeletal Muscle at 37ºC. Adapted from Hille, Ion Channels of Excitable Membranes Third Edition, pg. 17, Sinauer Associates, 2001 [1]. Ion Extracellular Concentration (mm) Intracellular Concentration (mm) Equilibrium Potential (mv) Na K Ca Cl

10 Examining table 1.1 we note that the equilibrium potentials for K + and Cl - are close to the resting potential of 90 mv for the muscle cell. Na + and Ca 2+ on the other hand have equilibrium potentials far from the resting potential. K + and Cl - set and stabilize the resting potential while Na + and Ca 2+ tend to drive or excite the membrane towards positive potentials. For this reason K + and Cl - channels are fundamental to all cells, with the K + family displaying extraordinary diversity. Na + channels are much less diverse and are found more specifically in excitable cells. Hille [1] broadly defines an excitable cell as any cell expressing voltage-gated Na + and Ca 2+ channels. For the most part, we consider nerves and muscle as the prototypical excitable cells. In these cells, we can easily appreciate the importance of the large superfamily of voltage-gated ion channels. Although these channels have other functions as well, it is perhaps clearest to introduce them as the molecular components underlying the propagating action potentials of neurons. The action potential is the basic unit of electrical signaling in nerves and muscle. Figure 1.2 shows an early recording of an action potential recorded from a giant squid axon [2]. During the first half of the 20 th century, Hodgkin, Huxley, and others set out to understand the ionic basis of membrane excitation. For a detailed description of the experimental history, see [1] and [3]. Armed with a newly invented instrument, the voltage clamp, Hodgkin and Huxley performed detailed studies of ionic conductance changes in the giant squid axon [4-6]. These experiments made possible a quantitative description of the action potential using an empirical kinetic description of the observed membrane conductance changes [7]. At the time of the HH model, the molecular basis of ionic flow across membranes was unknown. Today, we understand that ion permeation occurs through ion channel proteins and describe the action potential with this language, rather than referring merely to ionic conductance. The action potential is simply a local, transient membrane voltage change towards positive potentials. As a matter of vocabulary, we say the membrane is depolarized whenever the membrane potential is more positive than the resting state. Therefore, an action potential is a transient depolarization, even though the membrane voltage may become positive. We say the membrane is hyperpolarized whenever the membrane potential is more negative than the resting state. In the nerve axon, the upstroke of the 3

11 action potential is generated by a sudden influx of Na + ions through voltage-gated Na + channels, which drives the membrane potential towards the Nernst equilibrium potential for Na + (Table 1.1, Fig. 1.2). The Na + channels are designed to close after being open for a short time, a process common to all voltage-gated channels, called inactivation. At the same time, the membrane depolarization caused by sodium entry opens up voltagegated K + channels allowing K + to flow out of the cell, which drives the membrane potential back to the resting state. Figure 1.2, right plots these ionic permeability changes as a function of time. The action potential propagates along the nerve axon because the local influx of depolarizing Na + ions spread via electrodiffusion to neighboring membrane regions and initiate an above-threshold depolarization to activate downstream voltage-gated Na + channels. In this manner, the process repeats down the length of the nerve cell. The ability to propagate action potentials is hard-wired or programmed into the nerve axon by the voltage-gated channels present in the membrane. Na+ rushes into the cell causing swift depolarization. mv K+ channels open re-polarizing the membrane with outward K+ current. 1ms Figure 1.2 Nerve and muscle cells pass transient electrical pulses along their membranes as information. Left. An action potential recorded by Hodgkin and Huxley from a giant squid axon. Right. A plot showing the time course of membrane permeability changes that produce the action potential. Na + rushes into the cell causing membrane depolarization followed by quick inactivation of Na + channels. The membrane depolarization then causes K + channels to open after a short delay, resulting in restoration of the resting potential. Hodgkin and Huxley (H & H, Fig. 1.3) had no way of knowing what actually constituted the ionic pores they were studying, however they logically hypothesized the 4

12 existence of channels that were gated with voltage. Furthermore, they suggested the kinetic data implied the channels were controlled by several independent membranebound particles (they used four independent particles for their model [7]). These particles should carry electrical charge in order to make their movements sensitive to voltage. H & H pointed out that the motion of these charged gating-particles should also produce a small detectable electrical current preceding ionic current. Twenty years later, their prediction was validated with the first recording of gating currents [8-10] (see Chapter 2.4). In modern terms, the gating-particles are now called the voltage-sensors. The charges on these sensors are called the gating charges. Alan Lloyd Hodgkin Andrew Fielding Huxley Nobel Prize in Physiology or Medicine, 1963 "for their discoveries concerning the ionic mechanisms involved in excitation and inhibition in the peripheral and central portions of the nerve cell membrane" Figure 1.3 Hodgkin and Huxley shared the Nobel Prize in physiology or medicine with Sir John Carew Eccles in Their groundbreaking electrophysiological work epitomized the peak of classical biophysics. 1.2 Classical membrane topology of the Shaker K + channel In this work we study the conformational changes underlying voltage sensing in a voltage-gated K + channel from drosophila melanogaster (fruit fly) called Shaker. Shaker was the first K + channel to be cloned. The channel was identified and named because mutations in this gene caused flies to shake their legs under anesthesia [11]. Similar structural principles are assumed to underlie all voltage-gated channels because broad sequence homology exists across the entire superfamily of channels. Na V and Ca V 5

13 channels (modern nomenclature for voltage-gated Na + and Ca 2+ channels) have four repeat domains I, II, III, and IV (Fig. 1.4a). K V channels (modern nomenclature for voltage-gated K + channels) have analogous sequence structures made from four identical subunits they are homotetramers (Fig. 1.4b) [12]. The proteins in Fig. 1.4 are called channel α-subunits because they are the principal channel-forming subunits. Other proteins called auxiliary or regulatory subunits can interact with the channel and modulate various aspects of channel properties. Here we concern ourselves with K V channels in the absence of auxiliary subunits. The K V α-subunit is made of 6 transmembrane helices, denoted S1-S6, and a pore-forming loop (P) that contains the K + specific selectivity filter. Four α-subunits associate (tetramerization) in the membrane to form a rotationally symmetric protein with a central ion conduction pore (Fig 1.5). a Na V or Ca V α-subunit b ¼ of a K V channel (1 α-subunit) S1 S2 S3 S4 S5 P S6 Figure 1.4 Membrane topology for all voltage-gated channel α-subunits (principal channelforming subunits). a. The α-subunit for Na V and Ca V channels consist of four repeat domains that form the channel. b. The α-subunit for K V channels make up ¼ of a channel and tetramerization 6

14 of four K V α-subunits forms the channel. Each primary subunit of the K V channel has 6 transmembrane segments, S1-S6, and a pore-forming loop (P). S4 is called the principle voltage-sensor because it has intrinsic charge that constitutes most of the gating charge. K V Channel tetramer Four identical subunits surround the K + conduction pathway. K+ K+ S5 S3 S6 S2 S4 S1 K+ Figure 1.5 Four K V α-subunits form a rotationally symmetric homotetramer with the K + conduction pathway along the central axis. Cartoon channel viewed from above (left) with hypothetical S1- S6 positions indicated for one subunit. The channel topology of Fig. 1.4 is further divided into two major functional parts. S5-S6 is called the pore-domain because these segments make up both the pore and the gate(s). Every K + channel contains a pore domain homologous to S5-S6. S1-S4 7

15 is called the voltage-sensor domain because it contains the structural elements responsible for voltage-sensitive gating. S4 is called the primary voltage-sensor because it is highly charged and interacts with the membrane electric field in order to couple the gate (open probability) to voltage changes (cartoon Fig. 1.1c). The S4 segments are the charged gating particles hypothesized by Hodgkin and Huxley. The conformational changes of the voltage-sensor domain, particularly of the S4 segment, are of great interest. In this study we use a spectroscopic technique called lanthanide resonance energy transfer (LRET, see Chapters 3-5) to determine the physical movements responsible for voltage sensing. 1.3 How does an ion channel handle its charge? The fundamental task of an ion channel is to provide a pathway for strongly hydrophilic ions to cross the greasy, hydrophobic membrane barrier. Putting an ion into a membrane (dielectric constant ε = 2) from water (ε = 80) costs about half the total hydration energy of the ion, typically about kcal/mol. It is expected that the channel s conduction pathway should be polar, so that the passageway will be energetically favorable for charged ions. In fact, as early as the 1970 s, scientists (including Clay Armstrong) had deduced the general topology of the interior conduction pathway for K + and Na + channels. The principal gate was at the cytoplasmic side of the pathway followed by an interior aqueous vestibule that can bind channel blockers depending on whether the gate is open or closed [13]. Between the central vestibule and the outside solution, the protein pathway narrows into a polar selectivity filter. These features are shown in a cartoon from Hille, 1977 (Fig. 1.6) [14]. These early predictions were elegantly validated by K + channel X-ray crystallography (see Section 1.4). 8

16 Early cartoon representation of the ion conduction pathway. Hille Figure 1.6 The ion conduction pathway through an ion channel is hydrophilic. In between the gate (inside face) and the selectivity filter (outside face) resides a wide aqueous cavity that can contain fully hydrated ions. The selectivity filter is assumed to be very narrow and polar to support high fidelity conduction of a particular ion. We have established that a channel is voltage sensitive when its gate is coupled to a charged voltage-sensor domain that moves across the membrane electric field. Since the electric field falls across the membrane, it follows that the charged voltage-sensor must be located in the membrane. Therefore, K V channels have to solve the problem of moving charges across the membrane barrier twice; 1) the central ionic conduction pathway that we have described above and 2) a gating-charge conduction pathway. X- ray crystallography has provided exquisite molecular detail of the ionic conduction pathway (see Section 1.4) however the structure and mechanism of the voltage-sensor domain and gating charge motion are still under very active investigation. The S4 segment has a very particular arrangement of positively charged residues that is conserved in all voltage-sensitive channels. Arginines (R) and lysines (K) occur every three residues along the S4 (Fig. 1.7a), which if folded into an α-helix would produce a stripe of charge that slowly wraps around the helix (Fig. 1.7b). The positively charged S4 segments are in their closed position when the membrane voltage is negative. Membrane depolarization causes these voltage-sensor segments to move outward causing a net motion of gating charge from the inside solution to the external solution. 9

17 For the Shaker K V channel, the total gating charge per channel is about q e moving across the membrane electric field [15-17] during channel opening. Through mutational analysis, most of this gating charge for Shaker has been attributed to the first four arginines on the S4 (R362, R365, R368, and R371), although one acidic site (E293) on S2 also contributes to the charge movement [16,17]. a b Figure 1.7 a. Amino acid sequences of S4 segments from various voltage-gated K + channels with positively charged residues highlighted. The amino acid numbers for Shaker charges are shown on top. b. Positive charges every three amino acids results in a stripe of charge that slowly wraps itself around an alpha helical secondary structure. Figures adapted from Isacoff, 2002 [18]. It is generally believed that the intrinsic charges on S4 must be kept isolated from the low dielectric environment of the lipid membrane for energetic reasons. To accomplish this, the structure of the protein was expected to bury the charged S4 face against other protein segments, such as S2, that contain counter charges (acidic residues and partial charges). This expectation turned out to be only partially correct, as many of the basic residues have been shown to be in direct contact with either internal or external water. Numerous studies on Shaker have established the aqueous accessibility of cysteine substitutions along the S4 to thiol-reactive reagents (MTS reagents) applied from either the inside or the outside solution [19-22]. Only a small fraction (~10 amino acids) of S4 is not in contact with water and moving the channel from closed to open shifts which residues are buried. These changes are shown in a topological cartoon (Fig. 1.8) taken from Larsson et al., 1996 [19]. 10

18 S4 S4 Figure 1.8 Topological changes in aqueous accessibility along the charged S4 segment moving from closed (left) to open (right) states. Figure from Larsson et al., 1996 [19]. Accessibility results demonstrating a watery-environment for S4 motivated a revised generation of voltage-sensor models which include watery invaginations that penetrate the protein and put much of S4 in contact with the inside and outside solutions. These watery crevices are imagined to form a gating canal or gating-pore through which the S4 moves its gating charges. In general, three types of protein movement have been commonly used to model voltage-gating: (1) S4 translates in the up direction, perpendicular to the membrane, towards the external solution [19,23,24]. This type of motion is implied in the topological diagram Fig. 1.8 and the first cartoon Fig. 1.1c. (2) S4 rotates about its axis, moving charges from one aqueous crevice to another [25,26]. (3) Aqueous crevice reshaping [27-29]. These three types of protein rearrangements can exist in any number of combinations. For instance, a model that includes both vertical translation and rotation describes the motion of S4 as a helical screw (Fig. 1.9). The motivation for adding a twisting movement to a vertical translation of S4 is that the counter-charges located on surrounding protein segments can then be stationary while 11

19 successive S4 charges pass by. Figure 1.10 shows a cartoon version of the crevice reshaping model. Figure 1.11 shows the a purely rotational model for S4 proposed in Cha et al., 1999 [25] (discussed at length in Chapter 3.5) Voltage- Driven Helical Screw Figure 1.9 Helical Screw Model for S4 movement through an aqueous gating canal. Only the gating canal is pictured. Upon channel opening, the S4 segment rotates while translating outward into the external solution, carrying the gating charges outward. This cartoon is a realization of the model presented in Gandhi and Isacoff, 2002 [18]. 12

20 No Translation No Rotation Voltage- Driven Crevice Shaping Figure 1.10 Crevice reshaping model. Only the gating canal is pictured. S4 movement is not required to move gating charge across the membrane field if the gating canal changes shape during voltage-driven opening. 13

21 Figure 1.11 Pure rotation model for S4 movement. Top channel closed state. Two subunits of the entire channel are shown, with the other subunits removed for clarity. The S4 gating charges are exposed to an internal crevice, outlined in blue. Bottom open channel state. 180 degree rotation of S4 moves the gating charges from the internal crevice to the external crevice, outlined in magenta. No transmembrane displacement of S4 is predicted in this model. Figure taken from Cha et al., 1999 discussed in Chapter 3.5 [25]. In 2003, Roderick MacKinnon s lab published a new model of voltage-sensing based on their crystal structure of an archaebacterial voltage-gated channel, KvAP [30]. For a complete discussion of their data see below, Section The new model was 14

22 called the paddle model because the principal voltage-sensor S4 was observed to form a helix-turn-helix motif with the S3 (actually just part of S3, called S3b, Section 1.4), and the structure looked paddle-like. The paddle model consists of two very striking features that fly in the face of the generally accepted views shared among the conventional models described above: (1) The S4 voltage-sensor was placed at the periphery of the protein and in particular, the charges were allowed to make contact with the lipid membrane, despite the energetic cost such exposure implies. (2) The S3b-S4 paddle structure was hypothesized to undergo a large transmembrane displacement, generally from the bottom of the membrane to the top (Fig. 1.12, taken from Jiang et al [30]). As the paddle was hypothesized to undergo such a large displacement (15-20 Å [31]) through the membrane environment, it was described as a highly mobile hydrophobic cation. Closed Channel Paddle Down Open Channel Paddle Up Figure 1.12 MacKinnon s paddle model in cartoon form from Jiang et al [30]. Left. In the closed channel state, the paddles (S3b-S4 segments) are near the intracellular solution at the periphery of the protein. Right. Membrane depolarization causes the paddles to move upward towards the outside solution, transporting their charge across the membrane and pulling on the K + pathway gate. 15

23 In this dissertation, we examine the motion of the primary voltage-sensor S4 and other segments. Experiments have tested both motions parallel to the membrane (Section 3.5 and Chapter 4) and perpendicular to the membrane (Chapter 5). We have established that voltage-sensing segments undergo very small movements, and in particular the vertical movement of sites on S4 is ~ 2 Å. Our work greatly constrains the type of models that can be used to describe gating charge movement (Chapter 5.5, conclusions), and in particular, we conclude the paddle model does not describe the true mechanism of voltage sensing. 1.4 K + channels the atomic picture X-ray crystallography has greatly advanced the study of biological macromolecules by resolving three-dimensional structures with atomic detail (2-3 Å resolution). However, getting proteins to form an ordered crystal is not always straightforward and is notoriously difficult for integral-membrane proteins such as ion channels. The other stumbling block has been the need for milligrams of protein for crystallization trials and so a basic molecular-biological problem requires resolution. Great progress has been made in overcoming these problems, although much progress is surely yet to come. Firstly, the genomic era has uncovered a startling fact: bacterial organisms have ion channels of every important type, even those thought to be highly specialized such as voltage-gated channels. Some of these bacterial channels can be expressed at high levels (especially K + channels) and purified using standard biochemical procedures. Therefore, the protein quantity problem for crystallization trials is not intractable. Secondly, the field has been cracked open by the heroic efforts of Roderick MacKinnon, ion channel biophysicist-turned-crystallographer. MacKinnon has built his laboratory on the sound principle of combining traditional structural-functional methods mutagenesis, electrophysiology, and the like with structure determination. This section summarizes in detail the relevant ion channel structures, paying special attention to K + channels. 16

24 1.4.1 The KcsA structure a closed K + channel. KcsA was the first ion channel MacKinnon s lab successfully crystallized [32]. It is a K + channel from the bacterium Streptomyces lividans [33] and is proton-gated. This ground-breaking structure at 3.2 Å resolution showed how the K + conduction pathway works, how the channel rejects Na + but passes K + at near diffusional limits. It was now possible to understand the pore domain of potassium channels in great detail. As was shown in Fig. 1.6 above, classical studies had outlined how the pore was thought to be shaped. The physical gate was thought to reside near the intracellular side and the channel center should have an aqueous vestibule that prepares ions to enter into close physical contact with a selectivity filter. These predictions were beautifully shown by the atomic structure (Fig. 1.13a). All potassium channels have a signature sequence, minimally GYG but very often TVGYG. Mutations of these residues had resulted in decreased K + selectivity so it seemed likely that the signature sequence lined the selectivity filter [34,35]. MacKinnon s laboratory solved a higher resolution structure (2 Å) of KcsA by complexing the channel with monoclonal Fab antibody fragments [36]. At this resolution, detailed protein chemistry and ordered water could be resolved. The structure demonstrates unambiguously that K + comes into close contact with the carbonyl oxygens of the TVGYG sequence at the narrowest part along the permeation pathway (Fig. 1.13b). The coordination shows that the protein provides surrogate oxygen atoms that mimic the hydration shell for the K + ion that is queued in the aqueous cavity preceding the selectivity filter [37,38]. a b 17

25 Figure 1.13 Structure of the KcsA potassium channel. a. Two subunits are shown for clarity. The C-terminal ends of the M2 helices (homologous to S6) form a bundle that mostly closes the pore off to the inside solution. Therefore M2 (S6) helices form the gate. The central vestibule is marked with a red asterisk. The selectivity filter is circled. (a. adapted from [39]) b. The selectivity filter shown in detail (adapted from [36]). A fully hydrated ion resides in the aqueous vestibule below and dehydrated ions (green spheres) reside in the filter itself, coordinated to backbone oxygen atoms (red) The MthK structure - an open K + channel. The KcsA channel (above) crystallized in the closed state so understanding the gating transition, how the channel opens, could not be directly discerned. Since the M2 inner-helices (S6 helices for 6- transmembrane channels) seemed to cross and close off the conduction pathway, there must be a conformational change associated with these helices to open the channel. MacKinnon s laboratory cloned, characterized, and crystallized another K + channel called MthK from Methanobacterium thermoautotrophicum [40]. This channel is of the type shown in cartoon Fig. 1.1b, it binds intracellular Ca 2+ ligand to gate open. The membrane-spanning domain has high sequence homology to KcsA and many other K + channels. X-ray crystallographic analysis of this channel (including the intracellular Ca 2+ binding domain) in the presence of calcium resulted in a structure (at 3.3 Å resolution) that had a pore domain quite different from the KcsA. The M2 inner-helices of MthK were not crossed into an excluding bundle, rather they were bent 30, which appears to open the conduction pathway (Fig. 1.14a). The bend in the M2 segment occurred at a well-conserved glycine residue, called the gating hinge [39]. Although bending at the gating hinge was proposed to underlie the gating of all K + channels, functional evidence on the Shaker channel has suggested that in K V channels the pathway does not open nearly as widely as MthK. It was suggested that a conserved P-X-P motif (7 amino acids lower down from the gating hinge on M2) is responsible for opening up a narrower passageway and that differences in the gating mechanism can explain the lower conductance in these channels compared with their bacterial ancestors [41]. Furthermore, in Shaker, a mutation (V478W) just three residues down from the P-X-P motif on S6 create a non-conductive mutant through the formation of a hydrophobic seal [42]. Therefore, the MthK structure has provided powerful insight into the conformational 18

26 changes associated with K + channel gates, though variation in gate structure between different channels may exist. a b c Figure 1.14 Three comparisons between KcsA structure and MthK structure. a. Left shows the MthK crystal structure that has been modified to take on the shape of the KcsA structure, i.e. the M2 helices have been bent and twisted about the gating hinge. Right shows the unaltered MthK crystal structure, an open K + channel. Figure adapted from Jiang et al., 2002 [40]. b. The structural information from KcsA and MthK are presented as the general mechanism of gating for all K+ channels. c. Looking down the conduction pathway viewed from the inside of the cell. The bundle crossing of KcsA almost completely closes off the permeation pathway (left). The MthK structure shows a large opening (right) that allows K + and larger pore-blockers access to the aqueous vestibule. Figure taken from Webster et al., 2004 [41]. 19

27 1.4.3 The MscS structure a slightly voltage sensitive, non-selective mechanosensitive channel. Doug Rees and colleagues crystallized the first mechanosensitive ion channel (of large conductance), MscL from Mycobacterium tuberculosis [43]. These channels are responsible for maintaining osmotic balance across the bacterial cell membranes. In the event of a sudden drop in external osmolarity, water begins to flow into the cell (down its concentration gradient) and the cell swells. The flow of water can be reduced if the cell instead dumps ions into the external solution. Therefore, the opening of mechanosensitive ion channels can keep the cell from exploding in such situations. Mechanosensitivity is not what explicitly interests us here. A channel called MscS is a mechosensitive channel (of small conductance) from Escherichia coli that also displays a slight voltage-dependent gating [44]. The structure of MscS was solved by Doug Rees and colleagues at 3.9 Å resolution [45]. This channel structure, though structurally distant from both MscL and voltage-gated channels, offers a chance to understand mechanisms of gating. So how does the voltage sensitivity work for MscS? Both voltage and tension are coupled to the gate such that less depolarization is required as tension is increased and less tension is required as depolarization is increased. The voltage dependence requires a voltagesensor that can create net movement of charge across the membrane field. The gating charge for MscS has been estimated to be ~1.7 charges/channel (significantly lower than K V channels, with ~13 charges/channel), possibly arising from the movement of two arginine residues, R46 and R74, though this has not been determined conclusively. Since MscS is constructed from 7 identical subunits, these proposed gating charge residues actually represent 14 charges, therefore Bass et al. [45] note that a movement of ~2.4 angstroms/charge can account for the total gating charge (assuming a 20 Å hydrophobic bilayer). The crystal structure surprisingly suggested these two arginines are likely to be exposed to the bilayer, despite the energetic cost of placing charges in a membrane. Molecular dynamics simulations have suggested that the charged residues interact directly with the lipid polar head-groups and such interaction could explain the gating mechanism s voltage sensitivity [46]. Furthermore, new results using electron paramagnetic resonance (EPR) probes have indicated the charged residues have low lipid 20

28 accessibility (Vásquez, Cortez, Perozo; Poster 1421, Biophys. Soc. Meeting 2005 [47]) for MscS reconstituted in a lipid bilayer. The mechanism for MscS gating originally proposed by Bass et al. is shown in Fig. 1.15, however many more experiments will be needed before this channel is fully understood. Figure 1.15 Proposed mechanism for gating the MscS channel. Both tension and membrane depolarization increase the open probability of this homoheptameric, nonselective ion channel. Highlighted arginines are proposed to be the voltage-sensing residues. Figure taken from Bass et al., 2002 [45]. It has been hypothesized [48] that the MscS structure suggests a shared voltagesensing mechanism with the paddle model for K V channels (described in cartoon form, section 1.3 above, see section below). Both models have voltage-sensing structures at the protein-lipid interface, allowing gating charges to contact the hydrophobic membrane interior. In both cases the actual voltage sensor is a paddle-shaped helix-turnhelix structure (for MscS, the paddle is pointed down, opposite to the K V paddle) with arginine residues for gating charge. However, these channels have very little else in common structurally. The K V paddle model proposes a large (15-20 Å) vertical displacement of the voltage-sensor because K V channels have an extremely large total 21

29 gating charge that results in a very steep voltage dependence of channel opening [49]. MscS on the other hand is very weakly voltage-dependent and so a very small total gating charge is required. Bass et al. [45] suggest a vertical gating charge displacement of only 2.5 Å may be required to account for the total gating charge movement. Furthermore, evidence exists for both MscS and K V channels that the gating charges are not directly exposed to the hydrophobic membrane bilayer [47,50]. Therefore, it seems there are serious problems accepting paddle behavior as the mechanism for the straightforward translocation of gating charges across a hydrophobic barrier The KvAP structure a full-fledged, highly voltage-dependent channel. The KcsA and MthK structures (above) were highly successful at detailing the permeation pathway of K + channels. The pore domain validated expectations and also provided elegant insights into the energetics of highly selective, highly conductive channels. However, how do channels operate in a voltage-dependent manner? MscS provided clues perhaps, or at least teased us with clues, but the channel is very different from the superfamily of voltage-gated channels. What we really need are structures of full-fledged K V channels. In a scientific tour de force, the MacKinnon lab diffracted X-rays off a crystal of voltage-gated ion channels called KvAP [30]. Characterization of the electrophysiological properties of this K V channel from the archaebacterial hyperthermophile Aeropyrum pernix demonstrated that it was functionally similar to eukaryotic channels like Shaker [51]. The KvAP voltage-dependence of opening occurs at somewhat more negative voltages compared to Shaker and the sequence of KvAP has very little linker regions between transmembrane segments (S1-S2 and S3-S4 linkers are very large in Shaker). Otherwise, it is expected that sequence homology and functional homology make these channels very similar in structure. MacKinnon and coworkers used a channel from a hyperthermophile because they reasoned that the protein may be exceptionally stable and favor crystallization. However, after many failed trials they decided to raise antibodies against the voltage-sensor so that they could hold it in place for crystal growth. Thus, they interpreted their difficulty in crystallization as evidence that the voltage sensor is a highly mobile domain. The resulting monoclonal Fab 22

30 fragment-kvap crystal structure (at 3.2 Å resolution) was in a very unexpected conformation (Fig. 1.16a). The helices of both S3 and S4 were broken near the middle and formed two separate helical segments. The parts of S3 were labeled S3a and S3b. The two separate pieces of the traditional S4 segment were labeled S4 and S4-S5 linker [30], though another study has defined the N-terminal piece S4a and the C-terminal piece S4b [50]. It is unclear at this time whether these structural details extend exactly to other K V channels like Shaker, there are likely differences since the prolines and glycines that tend to break up α-helices occur at different places in the sequences. The Fab antibody fragments bound to an epitope at the extracellular end of the S3b-S4 helices, and apparently pulled it down towards the intracellular side (Fig. 1.16a). This led to a strange artifact: the S4 and gating charges were all the way down near the inside, yet the pore appeared to be in an open conformation. (This is contrary to the irrefutable electrophysiological fact that channel opening must correspond to outward movement of gating charge.) Furthermore, the S1-S4 helices were not packed into a tight structure but appeared pulled apart (Fig. 1.16a), with S1, S2, S3b and S4 parallel to the membrane rather than transmembrane as expected. The S3b-S4 structure was called the voltagesensing paddle and is detailed in Fig. 1.16b. a b S4-S5 linker may be S4b The S3b-S4 paddle. Figure 1.16 a. The KvAP crystal structure with two subunits removed for clarity. The pore domain is white and S1-S4 are indicated in color. Part of S4 forms a continuous helix with the S5 of the pore domain and this was labeled by Jiang et al. as the S4-S5 linker [30], though if S4 is in fact transmembrane as other models propose, then S4b may be more appropriate. Figure 23

31 adapted from Cuello et al., 2004 [50]. b. Detail of the S3b-S4 voltage-sensing paddle from the full length crystal structure. This structure is hypothesized to act as a highly mobile hydrophobic cation (the paddle), moving vertically through the lipid membrane. Figure adapted from Jiang et al [30]. Jiang et al. [30] published the crystallographic data shown in Fig. 1.16, aware that distortions existed in this structure. Thus, directly interpreting the structure in terms of the mechanism of voltage sensing was difficult if not impossible. More crystallographic data and more experiments were needed. First, they crystallized an isolated voltagesensor domain S1-S4 (Figure 1.17a), and found that S3b and S4 were associated in the same helix-turn-helix, paddle motif. (Any model of K V channels is likely to have S3 and S4 next to each other in the membrane as S3-S4 linkers vary widely, from non-existent, as in the KvAP channel, to very long, as in the Shaker channel. However, Shaker functions normally even with the S3-S4 linker almost completely removed [52].) Similarities in structure between the isolated voltage-sensor and the full-length channel were interpreted as an indication that S1-S4 was not distorted very much in the fulllength structure. Therefore, they reasoned that the Fab fragments distorted the channel by pulling segments down towards the intracellular space, but that the distortions were not so severe that they could not make adjustments to the structure and then suggest a mechanism for voltage sensing. After docking the isolated voltage-sensor structure to the pore domain using the S2 segment as a docking reference guide, an altered voltagesensing domain structure was proposed (Fig. 1.17b). S3b-S4 was proposed to lie at the channel periphery, in contact with the hydrophobic membrane, and voltage sensing is based on a large charge-carrying traversal across the span of the membrane (the proposed paddle model, Fig. 1.12). 24

32 a The voltage-sensing paddle, S3b-S4 b S4-S5 linker may be S4b Isolated voltagesensor. Docked. Figure 1.17 a. The isolated voltage-sensor structure. Paddle segments are circled. The C- terminal end of S4, called the S4-S5 linker in the full channel structure, is now part of a continuous helix with the N-terminal side of S4. b. (Above) One subunit of the full-length channel structure (S1 not shown for clarity). The isolated voltage-sensor is docked to the pore domain (only 1 subunit shown) using the position of the S2 segment as a docking reference. Figures adapted from Jiang et al., 2003 [30]. MacKinnon and coworkers sought to test experimentally the paddle model. A bacterial toxin-channel called colicin forms a voltage-dependent channel with a charged segment that traverses the membrane bilayer. Finkelstein and coworkers used biotinlinkers attached to the translocating region to show that a biotin-binding protein called avidin could attach to the linker from the internal solution or the external solution, depending on the state of the channel. This type of experiment could be described as an accessibility experiment using a fishing line that can determine how deep in the membrane a protein site is located. MacKinnon applied this technique to the KvAP paddle in order to test that the paddle is near the bottom of the membrane when the channel is closed and translocates to the top during opening (Fig. 1.18) [31]. 25

33 Figure 1.18 Experimental cartoon for testing KvAP paddle movement using avidin-biotin binding. When the paddle is down, biotin linked sites are expected to bind internal avidin. When the paddle is up, biotin linked sites are expected to bind external avidin. Binding from either side should cause a reduction in K + current. Figure taken from Jiang et al., 2003 [31]. Sites along S3b and S4 were labeled with the biotin linker fishing line, and labeled KvAP was reconstituted into lipid bilayers so that the channels could be voltageclamped. A control current trace was recorded before the addition of avidin protein. Next, avidin was added to either the inside or the outside solution and the resulting affect on the current was recorded. For all sites on S3b and sites at the top of S4, current inhibition occurred upon addition of avidin to the external solution. Membrane depolarization significantly speeded up this inhibition. For two sites on S4, inhibition occurred upon addition of avidin to both the internal solution (when the channel was closed) and the external solution (when the paddle was open.) This behavior is attributed to the biotin linker being dragged from the inside solution to the external solution upon channel opening. For two sites lower down on S4, inhibition occurred upon addition of avidin from the internal solution and never from the external solution. The complete data set is shown in Fig This pattern of behavior was used to constrain how deeply 26

34 residues on the paddle could be located in the membrane, as a function of voltage. Such constraints model the paddle s down configuration (Fig. 1.20a) and its up configuration (Fig. 1.20b) and voltage sensing involves the paddle moving between down and up (Fig. 1.20c and 1.20d). This is the logic and experimental evidence put forth by MacKinnon and colleagues in support of their paddle model. Figure 1.19 Complete data set showing the pattern of current modification after addition of avidin to the external solution (red traces) or internal solution (blue traces). Control currents are shown in black, recorded before the addition of avidin to either side. Red sites required membrane depolarization for fast inhibition from external avidin. Blue sites were only influenced by internal avidin. Yellow sites showed inhibition when avidin was added to either the internal or external side. Figure from Jiang et al., 2003 [31]. 27

35 Figure 1.20 The derivation of a paddle model cartoon from avidin accessibility data. a. The biotin tether length puts accessible sites within 10 Å from the internal solution in the down position. b. Accessibility data defines which sites are within 10 Å from the external solution in the up position. c. Imagined closed state for the paddle model this is a cartoon not a crystal structure. d. Imagined open state for the paddle model again this is not a structure. Figure taken from Jiang et al., 2003 [31]. So is the paddle model a true representation of how voltage sensing works for K V channels? Or is it just one big artifact? Or is KvAP just a very unusual channel, for which the normal rules of structural homology sweetly flowing out of sequence and functional homology just do not apply? The paddle model has been extremely 28

36 controversial, and many traditional data (usually obtained on the Shaker channel) have not been easily understood in terms of the crystal structure. Contradictions between the structure of Fig and traditional biophysical and biochemical data on functional channels are numerous. The concept of the paddle model for voltage-sensing has also been difficult to reconcile with many experimental results. Many of the issues have been discussed in reviews [53-55]. For example, the channel N-terminus is known to be intracellular (forming the T1 tetramerization domain in Shaker) [56] but in the structure appears to be buried in the membrane. Cysteine-scanning mutagenesis studies examining voltage-dependent accessibility to thiol-reactive reagents have shown that much of S3b and the linker between S3b and S4 are accessible to external reagents independent of voltage [19,21,57]. In the structure, these sites appear to be near the intracellular side of the membrane. Recently, the S4 segment was shown to be very close to the pore domain in the open state, as disulphide crosslinks are formed between pairs of cysteines introduced in S4 and the pore domain, S5/S6 [57-59]. This indicates that the structure of the open state is very different from Fig Interestingly, the residue on S4 that was shown to crosslink with the top of the pore domain is just about the farthest residue away from this crosslinking site in the structure of Fig. 2, and significantly, S1, S2, and S3b would have to get out of the way for this crosslink to occur. A low-resolution KvAP structure (10.5 Å) from electron microscopic (EM) single particle analysis (also from MacKinnon and coworkers) has also placed the voltagesensor paddle up against the pore domain (Fig. 1.21) [60]. This EM structure was not subject to the crystal packing forces of the X-ray sample and suggests a very different conformation. The location of the voltage-sensing paddle was determined because the EM data included the Fab antibody fragments (easily identified in the low-resolution structure) that bind to the paddle s tip. Two possible docking orientations were possible for S3b-S4 (both shown in Fig. 1.21). The second possible docking orientation (Fig. 1.21b and 1.21d) seems to agree best with the above-referenced S4 to pore domain crosslinking data. Unfortunately, both X-ray structure determination and single-particle EM analysis study channels outside of their native membrane environment. All of these issues suggest that the full length KvAP crystal structure is very distorted, unlike the conclusions of Jiang et al. [30]. 29

37 Figure 1.21 Models of the KvAP channel using single particle electron microscopy analysis. Crystallographic data for the pore-domain and the isolated paddle structure (S3b-S4) are docked to the low 10.5 Å resolution density map (blue mesh c and d). Two docking orientations for the paddle are possible. Orientation 1 is shown (a.) and from the top (c.). Orientation 2 is shown (b.) and from the top (d.). Red ovals indicate density that is unaccounted for and is likely taken up by segments S1, S2 and S3a. I have covered the paddle model proposal in detail because our measurements (Chapter 5) directly address the validity of this unconventional and surprising model. Furthermore, the paddle model has been presented as a cartoon and is necessarily ambiguous. Many of the data that run contrary to the paddle model are somewhat indirect, and there is a problem with attributing predictions that are too specific to an ambiguous model. We have tested the central hypothesis of the paddle model: a large vertical translation of the voltage-sensing segments. We find that no such translation occurs, and therefore conclude that the paddle model is not the general mechanism of gating charge movement in voltage-gated ion channels. 30

38 1.4.5 MacKinnon goes to Stockholm. In 2003, Roderick MacKinnon shared the Nobel Prize in Chemistry with Peter Agre. The structures discussed above, especially KcsA, have pushed our understanding of ion channels forward at a tremendous rate. As well as the structures discussed in detail above, MacKinnon crystallized a chloride channel, demonstrating how anionic channels are structured [61]. Even though contentions exist over the KvAP paddle model, research into K V channels has intensified and progressed as a result. For a highly readable perspective on MacKinnon s place in the history of ion channel research, see Chris Miller s essay, Ion Channels go to Stockholm this time as proteins [62]. Nobel Prize in Chemistry, 2003 "for structural and mechanistic studies of ion channels Roderick MacKinnon Figure 1.22 Roderick MacKinnon shared the Nobel Prize in Chemistry in 2003 with Peter Agre. Agre pioneered the work on Aquaporins water channels. MacKinnon propelled ion channel research forward with structure after structure. 31

39 Chapter 2 Electrophysiology of the Shaker K + Channel 2.1 The Xenopus laevis oocyte expression system With the advent and development of cloning techniques in the 1970 s and 1980 s it became possible to manipulate proteins such as Shaker and other ion channels in the lab. In order to study the function of the gene products of these DNA clones, an expression system was needed. By this time, the frog species Xenopus Laevis (Fig. 2.1, left) had already been established as a model system for reproductive biology and development, and the frogs were easy to handle and the extraction of egg cells straightforward. Furthermore, Xenopus oocytes (unfertilized egg cells, Fig. 2.1, right) were quite large (~1 mm diameter) and simple injection of mrna encoding membrane proteins resulted in protein expression [63,64]. The oocyte translates the mrna, incorporating the protein in a membrane vesicle, and then traffics it to the plasma membrane (outer membrane of the cell). Thus, shortly after the first K V clone (Shaker) was created [11], mrna was injected into oocytes and a voltage dependent K + current, familiar from classical neuronal biophysics, appeared under voltage clamp. The study of ion channels had entered the age of molecular biology. The further development of molecular biology, namely mutagenesis and recombinant DNA manipulation, has become the heart of basic ion channel biophysics. To this day, the standard procedure for studying an ion channel follows the mutate and measure strategy. Xenopus oocytes continue to be a convenient system for the expression and study of channels of many types. Voltage-gated channels are no exception, as these cells express huge amounts of protein and are easy to record using voltage clamping instrumentation (see below). We use the Xenopus system in our experiments - expressing ~10 10 Shaker channels/cell! 32

40 Xenopus laevis 1 mm Figure 2.1 Left. The African-clawed frog, Xenopus laevis laevis. Mature female frogs are about cm long. Right. Isolated oocytes at two different magnifications. The dark side (the animal pole) becomes the top of the frog and the light side (the vegetable pole) becomes the bottom. 2.2 The two-electrode voltage clamp In Chapter 1 we mentioned that Hodgkin and Huxley used a newly invented instrument, called the voltage clamp, to propel forward their studies of the giant squid axon. We also casually referred to channel currents, to K + flowing through ion channels. We referred to the phenomenon of gating currents, predicted by Hodgkin and Huxley [65] and eventually measured by Armstrong and Bezanilla [9]. In this chapter we discuss measurement of these currents using the Shaker K + channel expressed in oocytes. First, we describe the instrument used for taking these electrophysiological recordings. The term voltage-clamping refers to the control of membrane voltage. In particular, voltage clamps are used to hold the membrane voltage constant. This innovation allows for the direct measurement of ionic currents across membranes without contamination from capacitive current responses of the membrane (C m *dv m /dt = 0, Fig. 2.2a). When an experiment requires a change in voltage, the instrument applies a fast voltage step such that membrane capacitive currents occur as transients (Fig. 2.2b). To 33

41 demonstrate that the Xenopus oocyte membrane acts as a linear capacitor, we apply voltage steps with a voltage clamp, integrate the measured current transient to obtain the charge moved, and plot Q vs. V. The slope shows the constant linear capacitance of the oocyte membrane (Fig. 2.2c). a V C m The membrane is a linear capacitor that is charged, Q = C m V b c V I A voltage step command (V) initiates a transient charging current (I) of the membrane capacity. C = 0.31µF Q = *V Figure 2.2 a. A biological membrane is like a capacitor. The lipid bilayer has a low dielectric constant (ε=2) and the aqueous media on the external side and the internal side are conductors 34

42 (salt water). b. A sudden voltage step results in a transient charging/decharging current due to the membrane capacitance. c. Xenopus oocytes used in our experiment have linear membrane capacitance. Transients are generated with voltage clamp steps to various voltages from a starting voltage of -100 mv. Integration of these transients results in capacitance charge vs. voltage that is linear. The current recordings shown in Fig. 2.2c are significant because they demonstrate experimentally that depolarizing voltage steps elicit no currents other than the membrane capacitance response. This is an important property of the Xenopus oocytes: they do not exhibit endogenous expression of ion channels that interfere with the expression and study of exogenous channels like Shaker. However, Chapter 1 mentioned that K + and Cl - channels were fundamental to all cells, and therefore it is likely that some native channels are there. These endogenous channels are not likely to be voltage sensitive channels. For example, an endogenous Ca 2+ activated Cl - conductance has been described and studied [66]. Also, the over-expression of ion channels has been shown to induce currents in the oocyte that are unrelated to the exogenous channel expressed [67]. Voltage clamps come in several configurations, but the two principal techniques are whole-cell clamping and patch clamping. Patch clamping is perhaps the central modern technique that allows for the recording of small cells or a tiny excised patch of cell membrane. The resolution of patch clamping extends down to the single ion channel level [68]. Sakmann and Neher shared the Nobel Prize in Physiology or Medicine in 1991 for their discoveries concerning the function of single ion channels in cells. For our studies (Chapter 4 and 5), we record a very large ensemble of ion channels, using a standard whole-cell two electrode voltage clamp instrument. The important elements of the two-electrode voltage clamp are shown in Fig Two electrodes impale the oocyte. The first is a voltage measuring electrode (V 1 ) that simply determines the voltage inside the oocyte. The second electrode is the current injection electrode (V i ) that is used to change the voltage inside the oocyte. The clamp measures the current that crosses the membrane with an ammeter connected to ground. Therefore, the instrument includes a measure of V m, the membrane voltage, and a means to change it (current injection). The last piece is a feedback amplifier, which compares the measured V m with the voltage command input by the experimenter and delivers the 35

43 appropriate current to the current injection electrode in order to null its inputs. In other words, the amplifier clamps V m to the value requested. Command Voltage Pulse Feedback Amp V Voltage Measuring Electrode V 1 Outside Cell K + A Current Injection Electrode V i K + Inside Cell Figure 2.3 The two-electrode voltage clamp. Currents crossing the membrane such as K+ through ion channels and outward movement of voltage-sensing segments are recorded with an ammeter (A) connected to ground. The feedback amplifier is the essential element that compares the command voltage pulse to the measured voltage (V 1 ) and supplies a current to V i (current injection electrode) in order to null the amplifier inputs. The instrument we use (CA-1b high-performance oocyte clamp, Dagan) is in reality more complicated than the schematic cartoon of Fig. 2.3, however the idea is the 36

44 same. Our clamp has the two electrodes, as shown, but the bath (external solution) is also under active voltage clamp for faster, more accurate voltage control of V m. Another advanced element is a transient generator that allows the instrument to null out the capacity transients shown in Fig. 2.2 (capacity compensation). This is required for accurately measuring current responses that overlap the fast kinetics of the membrane response and is otherwise useful for reducing this uninteresting, high magnitude current. 2.3 Ionic currents Shaker channels conduct K + ionic current from the inside of Xenopus oocytes to the outside. Thus, when channels open a positive current will result (cells have high [K + ] on the inside). Generally, ionic current is very large and so only very low channel expression is required to detect K + current across the membrane. All channels have more states than simply closed and open, they have inactivation states. Inactivation refers to the elimination of current upon extended depolarizations. It is clear that channels have developed mechanisms to help safeguard the ionic gradients that they use for signaling. Also, inactivation properties are just as important as the activation properties in defining the shape and timing of action potentials in nerves. For Shaker there are two types of inactivation. The simplest form is called fast inactivation (or N-type inactivation). This type of inactivation is associated with a peptide segment at the channel N-terminus that plugs the channel (from the inside) after the activation gate opens [69]. This view of fast inactivation was called the ball and chain model because the peptide plug is tethered to the protein with a disordered peptide that diffuses around according to polymer statistics [70-72]. This inactivation ball could be cut off (destroyed with proteinase [70] or chopped off using molecular biology [73,74]) and fast inactivation was eliminated. K + currents persisted well after inactivation usually turned the channels off. In all of our experiments we use a construct called Shaker-IR, that has the inactivation ball chopped off with molecular biology ( 6-46) [75]. The difference between Shaker K + currents with and without the fast inactivation ball is shown in Fig. 2.4a (Data taken from Hoshi et al., 1990 [73]). 37

45 The second and more complicated form of Shaker inactivation is called slow inactivation (or C-type inactivation). This type of inactivation is more fundamental to the channel and can not be eliminated by simply chopping off an otherwise inconsequential part of the protein, like the fast inactivation peptide. In fact, the C-type inactivation gate seems to be associated with the selectivity filter itself (narrowing of the filter perhaps) [76]. Therefore, our Shaker-IR clones still inactivate. The timescale of slow inactivation is on the order of seconds, roughly 100 times slower than fast inactivation (Fig. 2.4b). In all of our experiments we depolarize and record for only 50 ms, a timescale for which slow inactivation is completely negligible. Example ionic K + currents through Shaker-IR channels recorded with our setup are shown in Fig a Shaker B 6-46 channel currents persist for many milliseconds b 20 ms Addition of the fastinactivation ball causes the channels to get plugged Slow inactivation turns Shaker Off over the course of seconds Figure 2.4 a. Top traces show Shaker-IR currents that display no fast inactivation, the currents persist over the course of 10s of milliseconds. Bottom traces show currents after the addition of diffusing inactivation balls to the intracellular side of the membrane. Fast inactivation is apparent from the sudden drop in current (data taken from [73]). b. Shaker-IR channels inactivate via slow inactivation over the course of several seconds. Traces show the addition of the inactivation peptide to the intracellular side of the membrane recovers fast inactivation. A mutant peptide has little effect on inactivation (data taken from [77]). 38

46 Figure 2.5 Example K + current recordings of Shaker-IR measured on the Selvin lab electrophysiology rig. Oocytes were expressing low amounts of ion channels so that the current is less than 50 µa. This form of presenting many recordings (current family) is common in electrophysiology presentations. The test pulse was varied from -100 mv to +100 mv. The test pulses are applied for 50 ms. 2.4 Gating currents In Chapter 1 we discussed models that describe movements of the highly charged S4 voltage-sensor. This segment ultimately opens and closes the channel by moving when the membrane electric field changes. Therefore, the conformational changes associated with S4 are of particular interest in understanding the mechanism of voltage sensing. Traditionally, S4 movements and voltage sensing were studied with 39

47 electrophysiology. The currents that arise from the movement of the S4 charges are called gating currents. These microscopic currents are a direct measure of protein conformational changes associated with the movement of S4. The magnitude of gating current (approximately 100x smaller than ionic current [78]) is very small because it arises from 13 elementary charges/channel crossing the field. Therefore, in order to detect gating currents, ionic currents need to be eliminated. Ionic currents are abolished by making mutations that eliminate conduction or by blocking the channels with molecules or proteins that block the flow of K +. These blocking methods need to leave the function of the voltage-sensing domains unaltered so that S4 movement can still be detected. In this study we use several types of channel block. In chapter 4, we use a common mutation, W434F [79], which makes Shaker nonconducting, probably by closing the C-type inactivation gate of the selectivity filter [80]. In chapter 5 we block channels with scorpion toxins that bind to the external end of the pore, clogging the mouth of the selectivity filter [17]. Both of these mechanisms block K + conduction but leave gating currents unaffected. In Fig. 2.6 we repeat the experiment of Fig. 2.2c, except this time we do not voltage clamp a blank background oocyte, we clamp an oocyte expressing a huge number (> ) of Shaker channels that are non-conducting via scorpion toxin block. The current transients that result from depolarizing voltage steps are not as fast as in Fig. 2.2c and highly complex kinetics have become apparent (Fig. 2.6 left). The integral of these recorded currents give a measure of the charge change across the membrane (which is only due to membrane capacitance and the voltage-sensing domains of Shaker) as a function of voltage (Fig. 2.6 right). Compare this plot with Fig. 2.2c. It is clear that a nonlinear capacitance has appeared in addition to the normal linear capacitance of the membrane. This nonlinearity is the charge movement of Shaker voltage-sensors. 40

48 10.0 µa 2.00 ms Figure 2.6 Left. Transient currents recorded from voltage clamp steps from -100 mv to voltages ranging from -150 mv to +50 mv, recorded on the Selvin lab electrophysiology rig. These transients are a mixture of the membrane capacitance response and the movement of the membrane-associated voltage-sensors (S4 segments). Currents show complex kinetics unlike the background transients from non-expressing oocyte membranes (Fig. 2.2c). The protein in this experiment was Shaker-IR blocked with 2 µm scorpion toxin (wild-type CTX). Right. The integrated charge ( Q) vs. test voltage (V) is now nonlinear (compare to Fig. 2.2c) because the voltage sensor segments move gating charge across the membrane with very nonlinear voltage dependence. Unlike the recordings shown in Fig. 2.6 (left), normally we use membrane capacitance compensation hardware to cancel the membrane response to voltage steps. Therefore, the recordings will only contain the channel gating currents. In Fig. 2.7 (left) we show gating currents recorded with a Shaker-IR/W434F non-conducting mutant using our instrument. Gating currents recorded from Shaker-IR blocked with a scorpion toxin are also shown (Fig. 2.7, right). The currents are transient because after a sudden step in voltage, the ensemble of channels move to a new conformational equilibrium. While the average conformational shift is occurring, there is a net movement of S4 charge across the membrane that produces a current. After the new state is reached on average, no net movement of S4 occurs and the current stops. At intermediate voltages, the S4s can be 41

49 moving between their closed and activated conformations, but the ensemble average shows no net movement after the gating current transient is over. Figure 2.7 Shaker gating currents recorded on the Selvin lab electrophysiology rig. Left. Shaker-IR/W434F, series of depolarizing pulses resulted in gating currents. Right. Shaker-IR blocked with 2 µm external charybdotoxin, a pore blocking protein from scorpion venom. The voltage dependence of S4 movement is usually expressed as the percentage of total gating charge moved vs. voltage, called a normalized Q-V curve. This function is easily calculated by the integration of gating currents (Fig. 2.7). This is exactly what we have been showing in Fig. 2.2c and Fig. 2.6, however with the membrane capacity response removed from our recordings we can now characterize the charge movement of the voltage-sensors uncontaminated (Fig. 2.8). Typically, the Q-V curve is normalized because the number of ion channels present in an experiment is not of fundamental interest, however the total gating charge is a good way to quantify how many channels you are managing to express. For our experiments we clamp ~10 10 channels. 42

50 Figure 2.8 Normalized Shaker-IR Q-V curve obtained from the integration of clean gating currents like those of Fig. 2.7 with no membrane capacity contaminations. In this experiment, K + conduction was blocked using a scorpion toxin (2 µm CTX). By 0 mv all voltage sensors have moved to their outward conformations. At hyperpolarized voltages, ~ -150mV, all voltage sensors are in their resting (closed) conformations. What is the relationship between channel opening and the voltage dependence of S4 movement, the Q-V curve? This question goes back to the very beginning with the Hodgkin and Huxley model. They assumed four independent gating particles moving with identical kinetics opened a channel [65]. This model predicted the kinetics of channel opening and predicted the existence of several closed channel states (1 gating particle moved, 2 gating particles moved, etc.). However, they had no way of measuring gating currents which give a direct measure of these otherwise silent closed states. Nowadays, scientists have access to detailed descriptions of macroscopic ionic currents, single channel currents, and for voltage-gated channels, gating currents. Therefore, a complete description of channel activation kinetics is possible if all of this information can be understood in terms of a model. Several detailed models have attempted to describe all the data for K V channels [81-83]. Although differences exist among these various models, several essential features have become clear. K V channels have four voltage sensors that move independently through at least two kinetic steps. All four voltage sensors have to be in their fully outward position at the same time so that a 43

51 subsequent cooperative transition(s) can then open the channel. As a consequence, gating currents precede opening and the Q-V curve is not identical to the open probability, as there will not be significant activation of channels until the percentage of charge moved is quite high. Fig. 2.9 plots the Shaker Q-V alongside the G-V (conductance curve or open probability), data taken from Stefani et al., 1994 [84]. Although there is significant overlap between Q-V and G-V, it is clear that a significant amount of the gating charge can move in a voltage range where channels rarely open. These features of the gating and activation voltage dependence are strikingly underscored in a mutant phenotype, the ILT-Shaker, which we discuss in the next section. Figure 2.9 Comparison of the Charge-voltage curve, Q-V, with the normalized conductance curve (G-V or open probability) for Shaker. The Q-V curve measures the voltage sensor transitions from the closed, inward position to the activated, outward positions. The plot has a best-fit that shows the Q-V curve can not be described by a single boltzmann function. The G-V describes the steep voltage dependence of channel opening. Data taken from Stefani et al., 1994 [84]. 44

52 2.5 The ILT-Shaker phenotype Aldrich and coworkers sought to understand how the S4 sequence affects the voltage dependence of gating charge movement, channel opening, and activation kinetics. Different K V channels behave quite differently in all of these respects, independent of the total charge on S4. Therefore, it is expected that the uncharged residues of S4 influence the basic activation pathway of the channels, probably by influencing the cooperative transition(s) the channel undergoes immediately before opening. Aldrich s approach was to construct chimera channels, using the Shaker channel S1-S3 and S5-S6 but with an S4 spliced in from four very different donor channels. The resulting chimeras had properties that were not predictable from the properties of the donor channels [85]. One channel, the Shaker/Shaw-S4 chimera, had an activation curve (normalized conductance) that was shifted to very positive voltages [86]. Evidently, the chimera greatly disrupted the cooperative transition(s) required for opening. This change in Shaker activation was shown to be mostly influenced by the single S4 amino acid change (I372L). However, a triple Shaker-S4 mutant (V369I, I372L, S376T, called the ILT mutant) reproduced all of the properties of the Shaw-S4 chimera channel. Study of the ILT-Shaker channel demonstrated that the gating currents occur normally like wild-type Shaker, however the channel is only opened by very strong depolarizations. Therefore, the Q-V curve of voltage sensing and the G-V curve of channel opening are completely separated with respect to voltage (Fig 2.10) [87]. The independent conformational transitions of the voltage-sensors are unhindered, but the cooperative transition is isolated along the voltage axis. The ILT-Shaker phenotype therefore, allows for the study of voltage sensing transitions separately from the transition(s) of opening. In chapter 4, we use the ILT-Shaker channels to study conformational changes associated with these separate stages of the activation pathway. 45

53 Figure 2.10 The ILT-Shaker phenotype. Open symbols: The charge-voltage relation (Q-V) is similar to the wild-type Shaker channel but with an approximately -40 mv shift (and a shallower slope). Closed symbols: The normalized conductance (G-V) is very different from wild-type with an approximately +110 mv shift (and a shallower slope). The Q-V and G-V no longer overlap. There is a very slight amount of gating charge movement for the G-V voltage range (~5%, [87,88]). Data taken from Ledwell et al., 1999 [87]. 46

54 Chapter 3 Fluorescence Spectroscopy Methods 3.1 Introduction to fluorescence Luminescence refers to the emission of light from an object as a result of excited state relaxation. Fluorescence is a sub-class of luminescence that results from singlet to single transitions (spin-preferred transitions) that occur rapidly. Organic dye molecules, called fluorophores, exhibit excited state lifetimes on the order of 1-10 ns (fluorophores used in this work are shown in Fig. 3.1). Other important properties of organic fluorophores include; high extinction coefficients (efficient absorption of light), high quantum yields, and broad emission spectra that are usually independent of the excitation process [89]. Furthermore, the short excited state lifetimes plus the high quantum yields make these molecules very bright and readily detectable as single molecules. The lifetime is also much shorter than the time-scale of many biological processes and protein motions, thus they act as instantaneous reporters when attached to biological macromolecules. Fluorescein-5- Maleimide (molecular probes) Atto465- Maleimide (atto-tec) Lucifer yellow- Iodoacetamide (molecular probes) Bodipy Fl- Maleimide (molecular probes) Figure 3.1 Organic dyes that are used in the experiments of this thesis. All of the molecules have organic ring structures containing electronic states with fluorescent transitions. These 47

55 probes are all green and have a maleimide or iodoacetamide for attachment to the thiol groups of protein cysteine residues. The phenomenon of fluorescence is often explained in terms of the Jabłoński diagram which illustrates the energy levels underlying fluorescence (Fig. 3.2, left). Upon absorption of a photon of energy hν, the electrons are kicked from the ground state S 0 into an excited state, either S 1 or S 2. The electron then relaxes to the lowest vibrational state of S 1, a thermal equilibration process (internal conversion) that is essentially instantaneous compared to the fluorescent excited state lifetime. The extremely high density of closely spaced vibrational energy levels associated with S1 and S2 explain the broad absorption spectra of fluorescent molecules (Fig. 3.2, right). Next, fluorescence photons are emitted when the electron falls back to various vibrational levels of the ground state. The distribution of emitted photon energies is explained by the tremendous number of closely spaced vibrational states of S 0 (see emission spectrum, Fig. 3.2, right). Another excitation relaxation pathway occurs when the excited electron undergoes a spin conversion to the triplet state T1 (intersystem crossing Fig. 3.2, left). The transition from T 1 to S 0 is forbidden and so phosphorescence lifetime is on the order of milliseconds, several orders of magnitude slower than fluorescence. Fluorophores such as those used here (Fig. 3.1) exhibit no observable phosphorescence. Figure 3.2 Left. Jabłoński diagram illustrating the transitions underlying fluorescence and phosphorescence. Fluorescence occurs when an excited electron falls from the first excited state S 1 to the ground state S 0. Phosphorescence occurs when the excited state undergoes intersystem crossing to the triple state T 1 and then falls to the ground state S 0. Right. Absorption 48

56 and emission of a commonly used fluorophore, tetramethylrhodamine. The spectra are broad because of the tremendous number of closely spaced vibrational energy levels of S 0 and S 1. Applications of fluorescence to the study of biology are numerous, varying from simple detection and localization studies to detailed structure-function studies exploiting various spectroscopic properties. Ensemble techniques as well as single-molecule techniques are now widely applied to a variety of biological systems [90-94]. The attachment of fluorescent probes to biological molecules is accomplished conveniently using thiol-reactive chemistry (amine reactive chemistry is also common). The fluorophore is synthesized with a chemical group that reacts under appropriate conditions with the cysteine thiols of proteins (maleimides and iodoacetamides in Fig. 3.1). Cysteines are somewhat rare in proteins, and when they occur, they can often be removed with mutagenesis without affecting protein function. Subsequently, cysteine mutations made on a protein with a cys-lite background define a unique labeling site for the fluorescent probe. Cysteine-scanning mutagenesis refers to the production of many protein mutants where the location of the introduced cysteine is systematically varied along a protein sequence. The reaction of maleimide and iodoacetamide with a cysteine thiol is illustrated in Fig Figure 3.3 Common reactive groups that covalently attach to protein thiols. The protein is represented by the R 2 group (green) and the fluorescent molecule is represented by R 1 (red). Figures taken from the molecular probes handbook ( 49

57 3.2 Resonance Energy Transfer (RET) theory In our work, we use fluorescence as a spectroscopic ruler [95,96] to measure intermolecular distances on the Shaker K + channel (Chapters 4 and 5). The conventional technique is called fluorescence resonance energy transfer (FRET) [97-99]. We apply a modified version called lanthanide resonance energy transfer (LRET) [ ]. Two luminescent labels of different structures and spectra are attached to the protein of interest. The shorter wavelength probe is called the donor and the longer wavelength probe is called the acceptor. The donor probe is excited by a laser or other light source, and can either emit photons or transfer its excitation energy to an acceptor. For efficient energy transfer to occur the energy of donor emission transitions must overlap the energy of acceptor excitation transitions. The energy transfer efficiency is used to calculate the distance between donor and acceptor and distance changes can be measured during protein conformational changes. Next, we discuss these techniques in detail before introducing their application to the Shaker channel. The original theoretical foundation for resonance energy transfer was worked out by Förster [103]. Here we present a discussion adapted from Selvin, 1996 [104]. Energy transfer occurs via the interaction of the electric dipole moments of the donor and acceptor. The rate at which energy transfer occurs, k et, is given by Fermi s golden rule: k et r r r r r r * µ D µ A 3( µ D R)( µ A R) * D, A D, A 3 5 R R 2 (3.1) where D and A refer to the donor and acceptor, and µ D and µ A are their electric transition dipole moments, R is the distance vector between the two dyes, and * denotes the excited state. The efficiency of energy transfer is given by: E = k et ket + k nd 1 = k 1+ k nd et (3.2) where k nd is the rate at which the donor de-excites through all non-distance dependent pathways, such as internal vibrations. Equation 3.1 is used to find a distance from 50

58 equation 3.2. Combining all of the physical constants from k nd and equation 3.1 into one quantity, R o, we are able to write the energy transfer in the common and useful form: 1 E = R. (3.3) 6 1+ ( ) The sixth-power dependence on R comes from squaring the dipole-dipole energy in Fermi s golden rule. Because E falls so steeply with increasing R, energy transfer is most sensitive to changes in distance when the distances are comparable to R o. R o is calculated from the spectral properties of donor and acceptor: R o Ro = ( JqDn κ ) (in Ǻ) (3.4) Where q D is the quantum yield of the donor, n is the refractive index of the surrounding medium, κ 2 is a factor that depends on the relative orientation of the donor and acceptor dipoles, and J is the normalized spectral overlap integral given by: J = 4 ε A( λ) f D ( λ) λ dλ. (3.5) f ( λ) dλ D Where ε A (λ) is the molar extinction coefficient of the acceptor and f D (λ) is the emission spectrum of the donor. For a FRET experiment to be quantitatively accurate, R o has to be accurately determined. This can certainly be achieved, since J is computed from easily measurable spectra, the quantum yield of the donor can be measured or approximated, and n is usually equal to 1.33 for water. κ 2 is usually the greatest source of error for energy transfer measurements. In many instances however, the donor and acceptor are attached to molecules with highly flexible attachments and the relative orientation between donor and acceptor samples all possible angles. In this ideal case, where donor and acceptor are completely unpolarized, κ 2 = 2/3. For LRET, the donor probe is intrinsically unpolarized and uncertainty in κ 2 is greatly reduced (below). 51

59 3.3 Lanthanide Resonance Energy Transfer - LRET LRET is a modification and improvement on the widely-used technique of fluorescence resonance energy transfer (FRET). These techniques utilize visible light (roughly 500 nm wavelength), yet achieve (sub-) nanometer resolution. In both techniques, a luminescent (fluorescent) probe called the donor, transfers energy via a dipole-dipole interaction to a second structurally-different probe, called the acceptor (Förster theory, above). FRET can measure distances between the probes over a range of Å, and LRET extends this range out to beyond 100 Å [102]. This high spatial resolution is possible, even with optical photons, because the amount of energy transferred (E) is a strong function of distance between the donor and acceptor fluorophores: E = 1/{(1+ (R/R o ) 6 }, where R o is the distance at which half of the energy is transferred and is generally Å. By knowing R o, which can be readily calculated or experimentally determined, and measuring E, the distance between the probes can be found. Labeling of probes to specific sites on biomolecules therefore enables the distances between these sites to be measured. Energy transfer can be measured because it reduces the donor s intensity and excited-state lifetime (E = 1- I da /I d, = 1 - τ da /τ d ), where subscript refers to donor s intensity or lifetime in the absence (I d, τ d ) and presence of acceptor (I da, τ da ), and also increases the acceptor s emitted intensity (Fig. 3.11). The donor's emission is at shorter wavelengths than the acceptor emission and hence they can be independently measured (although spectral overlap can occur). 52

60 Figure 3.11 Energy transfer between donor and acceptor falls off with the sixth power of R (top). As donor and acceptor get closer, donor fluorescence becomes dimmer, acceptor fluorescence increases, and donor lifetime is reduced (bottom). In LRET, the donor is a luminescent lanthanide atom encased in a small chelate (Fig 3.12a), and the acceptor is a conventional (organic) fluorophore (see Fig. 3.1). FRET uses conventional organic-based donors and acceptors. While relying on the same fundamental dipole-dipole mechanism, LRET has many technical advantages over FRET, including greater: distance accuracy and range; ability to resolve multiple D-A distances (donor populations); ability to isolate signal from proteins labeled with both donor and acceptor, even in the presence of proteins labeled only with donor or only with acceptor; and less sensitivity of energy transfer to orientation of dyes (which is often unknown). The fundamental advantages of LRET arise because the donor emission is longlived (Fig 3.12b; millisecond lifetime compared to nanosecond lifetime of acceptor or conventional dyes), sharply-spiked emission (Fig. 3.12c; peaks of a few nanometer width), has a high quantum yield [105], and is unpolarized [106]. (The chelate s atomic structure has also been determined [107]). 53

61 a b c Figure 3.12 a. Structure of Tb-DTPA-cs124-emph (maleimide-chelate used in our experiments, top). Crystal structure of Eu-DTPA-cs124 (dimensions 12.8 Å x 8.1 Å x 8.3 Å) [107]. b. Luminescence lifetime data for Tb-DTPA-cs124 and Eu-DTPA-cs124. c. Tb-DTPA-cs124 spectrum is sharply spiked and acceptor fluorescence (at 520nm) is measured with no donor contamination. An order of magnitude greater accuracy in distance-determination is achieved with LRET because the energy transfer process is dominated by the distance between the donor and acceptor, and their relative orientations play only a minor role in determining energy transfer efficiency. (A worst case scenario is 12% uncertainty in distance determination due to orientation effect.) This advantage results from the fact that the terbium donor emission is unpolarized [106]. This contrasts with FRET where the errors due to orientation effects can be unbounded. We have shown that angstrom changes due to protein conformational changes can readily be measured with LRET [25,108,109]. A 100-fold improvement in signal to background (S/B) is achieved with LRET. Specifically, energy transfer can be measured with essentially no contaminating background, in stark-contrast to FRET. By temporal and spectral discrimination, donor emission and acceptor emission both intensity and lifetime can independently be 54

62 measured. This leads to dramatically improved signal to background compared to FRET. Specifically, in LRET the acceptor emission due only to energy transfer called sensitized emission can be measured with no background. Contaminating background in FRET when trying to measure energy transfer via an increase in acceptor fluorescence arises from two sources: direct excitation of the acceptor by the excitation light, and donor emission at wavelengths where one looks for acceptor emission. In LRET both sources are eliminated. For example, by choosing an acceptor such as fluorescein and looking around 520 nm, donor emission is dark (Fig. 3.5c). By using pulsed excitation and collecting light at 520 nm only after a few tens of microseconds, all the direct acceptor emission (which has nanosecond lifetime) has decayed away. Samples that contain donor-only or acceptor-only can be spectrally and temporally discriminated against. Often when labeling proteins, particularly in living cells, one gets an unknown distribution of donor-donor, donor-acceptor, acceptor-acceptor mixture. In FRET this makes distance-determination difficult. In LRET, sensitized emission from acceptor arises only from donor-acceptor labeled complex (see preceding paragraph). Energy transfer of this D-A labeled complex can then be determined by comparing the lifetime of sensitized emission (τ ad ), which decays with micro- to millisecond lifetime, with the donor-only lifetime (τ d ): E = 1- τ ad /τ d. This ability to measure energy transfer even in complex labeling mixtures is essential for the LRET studies on ion channels presented below [25]. We have published a number of papers on LRET (partially reviewed in [102,104]) showing its advantage in model systems such as DNA oligomers [110,111], the ability to measure distance changes of an angstrom reliably even on large protein complexes such as actomyosin [108,109,112], and of most relevance to the studies in this dissertation, in ion channels in living cells [25]. Other workers have now successfully used the technique on DNA-protein complexes [ ], actomyosin [116,117], protein-protein interactions in cells [118], and detection of binding of many different biomolecules [ ]. 55

63 3.4 LRET meets the voltage clamp In 1999, two groups reported resonance energy transfer results on the Shaker K V channel. Isacoff and coworkers used conventional FRET [26], however nonspecific labeling and other technical limitations of FRET seemed to make the measurements inaccurate. The above technical advantages we have reviewed for LRET however, enabled Cha, Snyder, Selvin, & Bezanilla to measure accurate distances within the Shaker potassium channel in oocytes using LRET [25]. Here we review this application of LRET in detail because it is a direct blueprint for new LRET results presented in Chapter 4. Chapter 5 then presents further LRET results taken with a new donoracceptor labeling configuration in order to obtain complimentary structural information. Methodology: Site-specific labeling was obtained by substituting a cysteine for a particular residue (Fig. 3.13a) via site-directed mutagenesis and attaching cysteinereactive terbium donor (Fig. 3.12a) or acceptor (fluorescein maleimide, Fig. 3.1), to the four identical subunits of the channel (Fig. 3.13b). We refer to this arrangement as the S4-donor to S4 acceptor version of the experiment, because it measures distances from one subunit to the other, distances that are parallel to the membrane (Fig. 3.13c). Intersubunit distances were obtained by measuring the time constant of sensitized emission of the acceptor, i.e., fluorescence of the acceptor after receiving energy from the donor, and comparing this to the time constant of the donor attached to the same site in the absence of the acceptor. Results for residues S346C, S351C, S352C, N353C are reported here: they are in the S3-S4 linker, (Fig. 3.13a), accessible to labeling from the outside of the cell, and near S4, the region of primary interest. As a control, results at F425C are also reported here (see below). 56

64 a b c Donor LRET Acceptor X Figure 3.13 a. Membrane topology of Shaker with grey circles showing sites studied with LRET in Cha et al, Charged residues are indicated on S4 and S2. b. Labeling of homotetramers that have 1 cysteine per subunit results in a square geometry with two distances, R SC (subunits contiguous) and R SA (subunits across). Labeling is done with a 4:1 mixture of Tb-donor:Flacceptor such that most channels have only 1 or no acceptor. c. Cartoon of LRET experiment on the Shaker-IR channel in the S4-donor to S4-acceptor configuration. Distances are measured parallel to the membrane. Although labeling leads to a heterogeneous population of channels with different numbers of donors and acceptors, associated problems are greatly minimized for three reasons. 1) Channels that are labeled with four donors or four acceptors have no donor- 57

65 acceptor pair generating acceptor sensitized emission, i.e., they do not contribute to the signal. 2) Assuming four-fold symmetry of the channel, as demonstrated by the crystal structures of the closely related potassium channels KcsA and KvAP [30,32], there are only two possible inter-subunit distances: the distance between residues on neighboring subunits (R SC ) and the distance between residues across the pore (R SA ), which are related by the Pythagorean theorem (Fig. 3.13b). 3) The labeling is done with excess donor (4 donors for every 1 acceptor) so that there is typically only one acceptor per channel, which can readily accept energy independently from multiple donors. To avoid problems with slow-inactivation of the channel, yet ensure that the position of the channel residues were at a steady-state, the oocytes were voltage clamped at a resting potential of 90 mv, brought up to the desired test voltage for 50 msec, and then the laser fired and LRET data collected (Fig. 3.14b). The process was then repeated approximately once per second until sufficient signal to noise was achieved. a b Figure 3.14 a. Oocytes are voltage clamped on an inverted microscope so that fluorescence can be collected from below. Both voltage clamping and LRET are acquired by timed pulses (voltage pulses and laser pulses, respectively) and therefore the techniques are nicely integrated. (Note: The data from Cha et al., 1999 [25] used a cut-open oocyte voltage clamp different from the twoelectrode clamp pictured, however the concept is exactly the same. Selvin lab is equipped with a two-electrode clamp, which we use in our experiments, Chapter 4 and 5.) b. Voltage steps were applied from -90 mv to test potentials between -120 mv and 50 mv. The laser is fired 50 ms 58

66 after the initiation of voltage pulses so that channels have plenty of time to reach conformational equilibrium. Results: The intersubunit distances at several sites were measured by comparing the time constants of acceptor sensitized emission, and donor emission without acceptor: R = R o {(τ ad /(τ d -τ ad )} 1/6, where R o was determined to be 45 Å [98]. (Note: a corrected calculation shows that the R o for Terbium to Fluorescein is 43 Å, using the correct quantum yield for the Tb-donor [105].) Donor lifetime in the absence of acceptor was independent of voltage, indicating no significant change in the environment of the caged terbium (data not shown). Similarly, the intensity, polarization (which was minimal), spectra and labeling efficiency of the acceptor were unchanged with voltage, indicating that neither the acceptor nor donor are likely moving with respect to the protein as a function of voltage. Thus changes in distance between labels likely indicate changes in the underlying protein conformation. The sensitized emission for all sites displayed two time constants, reflecting two donor-acceptor distances that showed a Pythagorean relationship (Fig. 3.15a shows representative data for probes at S346C). This indicates that the technique is measuring distances between contiguous subunits and across the pore simultaneously. For example, at S346C, distances of 28 Å and 41 Å were measured, where the Pythagorean relationship predicts a distance of 40 Å for R SA given that the measured distance is 28 Å between contiguous subunits. Further verification of the technique was achieved by measuring distances at F425C, where a homologous residue is present in the crystal structure of the KcsA bacterial analog [32]. A distance of 30 Å was measured across the pore, in excellent agreement with the 29 Å distance obtained between α-carbons for the homologous residue from the crystal structure. 59

67 a b Figure 3.15 a. Representative data from S346C, a residue at the top of the S3-S4 linker on Shaker. Two time-constants were obtained from LRET sensitized emission and calculated distances followed the expected Pythagorean relationship. b. Sensitized emission at S346C displayed an obvious voltage-dependence corresponding to a voltage dependent conformational change. Next, voltage-dependent movements near the voltage-sensing regions were measured by determining inter-subunit distances as a function of voltage. Residue S346C demonstrated a robust change in the time constant of acceptor sensitized emission (Fig. 3.15b). This change in time constant reflected a voltage-dependent movement of ~3.2 Å between S346C residues on contiguous subunits, or ~4.5 Å across the pore, as the channel moves into the open state (Fig. 3.16). These results were the first measurements of actual distance changes around the voltage sensor as the channel goes from the closed to the open state. Furthermore, the voltage dependence of the physical movement for S346C correlated very well with the gating charge movement for the same channels (Fig. 3.16). In other words, the movement of this particular residue, S346C, is well correlated to the movement of all the charged residues moving in the channel, the latter creating the gating current, which is fundamental to the gating of the channel. 60

68 Figure 3.16 Open symbols: R SC vs. voltage for S346C (at the top of the S3-S4 linker in Shaker). The data shows a 3.2 Å change with the sites getting farther away from each other as the channel opens. Closed symbols: Q-V for the labeled S346C channels. Gating currents were recorded simultaneously with LRET. Charge movement and changes in LRET were strongly correlated. Voltage-dependent energy transfer changes were also detected at sites S351C, S352C, and N353C - three successive residues near the S4 segment (Fig. 3.13a). Surprisingly, the changes are different for each site (Fig. 3.17a). The donor and acceptor attached to S351C residues move ~1 Å further apart as the channel opens. In contrast, site S352C shows no significant change in distance, and site N353C moves ~1 Å closer together as the voltage is depolarized. Note that while these changes are small, they are highly reproducible (see error bars in figure). It is also possible that the energy transfer changes at these sites are caused by a reorientation of the acceptor without a corresponding translation. (Energy transfer in LRET is weakly dependent on the acceptor orientation with respect to the radius vector joining the donor and acceptor.) These changes in distance were argued to demonstrate a rotation of this protein region, although it is possible that a simple tilting or some other small rearrangement could produce these changes. If these residues were undergoing a large translation (as opposed to a rotation) across the membrane, which is the simplest and most common hypothesis for the movement of S4, we would expect all residues to change distances in the same direction, which is not found. A physical model consistent with the data is shown in Fig. 3.17b. In this scenario, this region of the protein is portrayed as an α-helix, and a

69 rotation moves one residue further from the pore (S351C; black), one residue closer (N353C; red), and one residue maintains the same distance (S352C; blue). a b closed open Figure 3.17 a. Three residues on the S3-S4 linker near S4, the primary voltage sensor, moved a small amount, with R SA for S351C becoming 1 Å greater, R SA for S352C remaining the same, and R SA for N353C becoming 1 Å less upon depolarization. b. Such a pattern in distance changes is thought to be consistent with a rotation model for the protein conformational change. S351C is represented by black dots. S352C is represented by blue dots. N353C is represented by red dots. A rotation of the S4 region with no apparent outward translation is a surprising result since it is known that the open and closed state of the channel differ in energy arising from the movement of charge through the electric field across the membrane (from inside to outside potential). Indeed, several older models have postulated a translation of S4 as much as 16 Å across the membrane [19,24] and the recent controversial KvAP paddle model predicts a vertical movement around Å [31]. However, not only does the data at suggest a rotation in the S4 (or more precisely the S3-S4 region), it was argued the sigmoidal shape of the voltage-dependent movements of S346C (Fig. 3.16), S531C, S352C, and N353C are inconsistent with a significant translation. To understand this, consider a model where S4 undergoes a significant translation (Fig. 3.18). Note that there are four S4 segments per channel, and 62

70 to go from the open to closed state all S4 segments must move. At negative potentials, the channel is closed and all S4s are down ( closed state in Fig. 3.18). At intermediate potentials ( active state), some S4s are up and some are down, although the channel remains closed. At more positive potentials, all S4s are up and the channel is open. This picture has the S4 voltage-sensors moving independently of one other, which has been demonstrating convincingly by several different lines of evidence [87,88,122]. With a transmembrane movement of 16 Å, the inter-subunit distance versus voltage would demonstrate a bell-shaped voltage dependence with a peak change in distance of approximately 2.2 Å at intermediate potentials (Fig. 3.18). Smaller translations would still be bell-shaped with simply a lower peak change. Since the actual voltage-dependent movement is sigmoidal, and not bell-shaped, it is unlikely that the S4 segment undergoes a significant transmembrane movement with voltage. This argument however, is substantially indirect and relies on the S4 helices to be slowly moving (between closed and open states at intermediate potentials) on the time scale of the LRET measurement. If LRET measurements were instantaneous then the distribution of S4s would be exactly like the central cartoon of Fig. 3.11, however since LRET measurements take longer, usually several hundred microseconds, the S4 helices will be moving during the measurement. Generally, the S4s go between closed and open on the order of milliseconds, therefore the LRET measurement is significantly faster. However this argument against vertical translation is subtle and indirect, and the experimental data could not quantify a small vertical translation if it exists. In Chapter 5 we show data that overcomes this limitation of the S4-donor to S4-acceptor version of the experiment. In order to test the MacKinnon paddle model, we use LRET to rule out a significant vertical translation of S4 conclusively. 63

71 Figure 3.18 A model for S4 movement that includes a significant vertical displacement perpendicular to the membrane. At intermediate potentials ( active ) some S4s will be displaced to their outward positions while other S4s will be in their down state. Therefore the measured distance will be greatest at intermediate potentials while a mixture of S4 states is occurring. The closed and open states show the same distance because all S4s are in the same conformational state. This bell-shaped change in distance between S4s was not observed for any site near or on S4 in Cha et al., 1999 [25]. How can a rotation of S4 carry 13 electronic charges per channel across the electric field from inside potential to outside potential? Cha et al., 1999 [25] suggested a model that is consistent with the LRET data and chemical accessibility studies (Fig. 1.11). In both closed and open states of the channel, residues in the S4 segment reside in crevices for which protons have deeper accessibility than cysteine-reactive reagents, as shown by histidine and cysteine-scanning accessibility studies [19,21, ]. As the channel goes from the closed to open state, the S4 segment rotates, moving the key charged residues R362, R365, R368, and R371 from one crevice connected to the intracellular potential to another crevice connected to the extracellular potential. The water-filled crevices focus the electric field across a relatively thin hydrophobic region, permitting a small conformational change such as a rotation to move the charge across 64

72 the field. This rotation of S4 also changes the chemical accessibility of these residues from the intracellular to the extracellular solution. No S4 translation required. In summary, these results show that the technique of LRET, combined with sitedirected fluorescent labeling, has the power to study atomic-scale structural changes in the Shaker channel in vivo. Initial results on the Shaker potassium channel suggested a rotational model of the voltage-sensing S4 region with the motion of the S3-S4 linker domain strongly correlated to gating currents arising from S4 charge movement [25]. 3.5 Instrumentation The Selvin lab voltage-clamp is the CA-1B oocyte clamp in two electrode mode (Dagan). Electrophysiology solutions are listed in Appendix D. Recordings were filtered at 20 khz and digitized with an A/D conversion card (Innovative Integration). Voltage command pulses were produced using a D/A (Innovative Integration). The electrophysiology apparatus was controlled and data was collected and analyzed using inhouse software from the Bezanilla laboratory. The optical setup consisted of an Olympus inverted IX-70 microscope with a 40x quartz objective (numerical aperture 0.8, Partec). The lanthanide was excited with a pulsed 337 nm nitrogen laser source (Oriel), reflected by a 400DCLP dichroic (Chroma). Donor and acceptor fluorescence were separated using a Q505lp beam splitter and collected simultaneously with D490/10 and HQ520/20 filters, respectively (Chroma). Fluorescence was detected with two water-cooled R photomultiplier tubes (Hamamatsu) operated at V. Prompt fluorescence was rejected using an electronic gate (Products for Research) with a dead-time of 70 µs. The detector current was converted to voltage (10 6 V/A, Hamamatsu), filtered at 50 khz (8-pole Bessel filter, Dagan), and digitized with an A/D (National Instruments). 65

73 Chapter 4 LRET Part I: The ILT-Shaker Channel 4.1 What can measurements on the ILT channel tell us? Cha et al. measured changes in intersubunit distances on Shaker using the S4- donor to S4-acceptor LRET configuration (Chapter 3.5). A summary of all the results obtained in that study are presented here in Fig Even though the change in distance measured for S346C (at the top of the S3-S4 linker) displayed a strong correlation with gating charge movement (Fig. 3.16), it is possible that the movement is in fact correlated with the opening steps of the channel. Alternatively, a part of the distance change may correlate with gating charge movement and the remaining distance change may correlate with the final opening steps of the channel. This can not be determined by studying wildtype Shaker due to the high overlap of the charge-voltage relation (Q-V) and conductance curve (G-V, see Fig. 2.10). Applying LRET to the ILT-Shaker mutant we can now determine the physical movements associated with the movement of gating charge separately from movements associated with the cooperative steps of opening. To first order, we can take the Cha et al. results (Fig. 4.1) as indicating the total movement at a particular site and we can measure in the ILT channel what voltage range displays this movement. Several sites from Cha et al. showed no movement so we naturally base our study on the sites that did show a movement; S346C, S351C, and N353C. However, it will also be important to study sites that didn t show a change in the former study because movements opposite each other along the activation pathway may become exposed whereas they simply masked each other in the wild-type channel. 66

74 Figure 4.1 Summary of all the distance measurements obtained from Cha et al., 1999 [25]. For the location of these sites on the Shaker membrane topology see Fig. 3.13a. The measurements demonstrate a movement for three sites; S346C, S351C, and N353C. The columns R SC and R SA were determined from two exponential fits and R SA was determined by taking R SC x We examine 346 and 351 with LRET using the ILT-Shaker channel to determine where these changes occur along the activation pathway. 4.2 Initial LRET results on the ILT channel The methodology used to perform LRET measurements on the ILT-Shaker were identical to those used by Cha et al. and were described in detail in Section 3.5. The constructs used are ILT-Shaker-IR/W434F. The W434F mutant prevents K + conduction and allows us to measure gating currents simultaneously with LRET. However, at the very positive potentials (0 mv to +200 mv) contamination K + currents are recorded because W434F is not strictly nonconducting [80] - strong depolarizations push ions through. Therefore, measurement of the small amount of gating charge that is moved during the isolated channel opening steps (Fig. 2.10) could not be repeated in our measurements. Slight modification was made to the voltage pulsing protocol. Test pulses from -150 mv to -25 mv (in 25 mv increments) were initiated from a prepulse potential of -100 mv in order to study the voltage range for charge movement (Q-V). For the measurements across the activation voltage range (G-V), test pulses from 0 mv to 67

75 +200 or +225 mv were initiated from a prepulse potential of 0 mv. No slow-type inactivation was detected using a prepulse to 0 mv and in fact, the ILT mutations seem to inhibit slow inactivation along with the cooperative opening transition(s). This result was tested by measuring whether gating charge was immobilized by prolonged depolarizations to 0 mv, a property of slow-type inactivation [72]. If slow inactivation occurs a hysteresis appears in the charge-voltage relation because greater hyperpolarizations are required to return the gating charges to their resting state (inward, down state). This result was qualitatively described in the initial characterization of ILT [87] and quantitatively presented in another study [122]. Fig. 4.2 plots the distance between contiguous subunits Rsc vs. voltage obtained for the ILT-Shaker-S346C mutant (at the top of the S3-S4 linker) from the shorter of two lifetime components from LRET (raw data not shown). The normalized gating charge vs. voltage curve (Q-V curve) is also plotted to show how well the LRET voltage dependence correlates with the traditional electrical measurement of voltage-sensor movement. Note how well Fig. 4.2 reproduces the distances and distance change published for the site S346C in the wild-type Shaker channel in Fig (the change is a bit greater, about 4.3 Å). The other important result of Fig. 4.2 is that the physical movement of residue 346 correlates with the charge movement, Q-V, and very little movement (< 0.5 Å) is seen for this site in the voltage range of channel opening (between 0 and 225mV, see the G-V in Fig. 2.10). Two sets of data are plotted in 4.2. The blue data were obtained from cells that were labeled with a ratio of 4 donors to every 1 acceptor (data is the average of 7 cells, error bars are the standard error of the mean). To check that the labeling conditions actually produce data representative of channels with only 1 acceptor probe and that signals from channels with 2 or more acceptors are negligible, we measured LRET on cells labeled with 9 donors to every 1 acceptor. This labeling ratio produces less overall signal, but gives even greater weight to the channels of interest (labeled with only 1 acceptor). The red data shows the distances vs. voltage for these cells and clearly the two labeling conditions give the same distance vs. voltage result (data is the average of 7 cells, error bars are the standard error of the mean). 68

76 Figure 4.2 LRET measurements for ILT-Shaker-S346C. Movement of this site correlated strongly with the Q-V and very little distance change (< 0.5 Å) is observed between 0 mv and +200 mv (channel activation). In Fig. 4.3 we present data for the ILT-Shaker-S351C mutant. The distance between contiguous S351C residues, R SC, is again calculated from the short lifetime component of sensitized acceptor emission data (blue data, average of 5 cells). Oocyte cells were labeled with 4 donors to every 1 acceptor and data were fit to 2 exponentials. This residue was previously measured to have an R SC of about 28 Å with very little movement (< 1 Å) [25]. Here we show again that the distance between contiguous S351C sites is about 28 Å and we detect an overall movement of about 1 Å (again somewhat greater than that detected by Cha et al.) that mostly correlates with the ILT- 69

77 Shaker charge movement, the Q-V curve. The movement of this site between 0 and 225mV, the voltage range of channel opening, is small, ~0.5 Å or less. Figure 4.3 LRET measurements for ILT-Shaker-S351C. Most of the movement of this site is correlated strongly with the Q-V and a small distance change (~0.5 Å) is observed between 0 mv and +200 mv (channel activation). These data for S346C and S351C demonstrate unambiguously that almost all of the physical movement of these sites is directly coupled to the movement of gating charge across the membrane. Perhaps this is expected, since the voltage-sensing arginines are near the extracellular side of the membrane (that is, in a transmembrane model, see Chapter 5) rather far from the actual activation gates (the S6 helical bundle). It could be that motion of segments near the outside of the channel will correlate with gating charge movement, which in turn allows gating conformational changes that occur 70

78 deeper in the channel (physically closer to the activation gates themselves). The ILT mutation that mostly disrupts channel opening (I372L) is 21 amino acids farther down on S4 compared to our labeling site S351C. Another site was tested on the Shaker wild type channel, L361C, which is on the S4 itself right before the first charged arginine (data not shown). No movement was found for this site, which is consistent with the finding by Cha et al. that V363C, the site right after the first charged arginine does not move. It will be necessary to test these sites with the ILT channel background to make sure that no movement occurs. Therefore, it appears that we can not detect movements of S4 itself using this S4-donor to S4-acceptor arrangement of LRET. This suggests that if S4 moves at all, it is likely a purely vertical movement. We address the issue of S4 vertical movement with direct measurements in Chapter 5. Whether other ILT channel sites, on S4 or elsewhere such as S1-S3, show movements that correlate with the actual opening of the channel is the next important question we will address. We will explore other sites and look for movements that are related distinctly to charge-movement or channel opening. This project is ongoing. We have put it aside temporarily because the paddle model inspired a new measurement that could address the issue of vertical motion for voltage-sensing segments. We now turn to this project (Chapter 5) that has been completed. 71

79 Chapter 5 LRET Part II: Shaker with Scorpion Toxin As soon as MacKinnon presented his new paddle model for gating charge movement [31] based on the first X-ray structure of a K V channel, KvAP (see Chapter 1) [30], many labs sought to disprove the paddle concept with new experiments. Various experiments included a new accessibility study [57], crosslinking studies between the pore domain and S4 [57-59], and a clever study that examined which sites on S4 are capable of carrying charge across the electric field [126]. These studies were presented as contradicting the paddle mechanism of voltage sensing. However, these studies tested predictions of the new model that were at least one step removed from the basic idea of the paddle mechanism. In our lab, we sought to directly test the central feature of the model, a large vertical translation of the S3b and S4 segments. Others have not been able to put this basic feature under scrutiny, and it is just the kind of thing that energy transfer experiments are good for. Testing the vertical translation of S3b-S4 is of central importance in evaluating the validity of the paddle mechanism, as the model s other unusual feature, the location of S4 at the lipid interface, has been shown to be experimentally plausible [50,127]. Here we show that LRET demonstrates no large perpendicular movement of the voltage sensing segments, in stark contrast to the idea of a voltage-sensing paddle. The results are consistent with S3 and S4 segments oriented in a transmembrane fashion for all voltages. The small conformational changes that we have been able to detect using LRET have demonstrated that the voltage sensor does not need to move very much in order to carry its huge gating charge across the membrane field. This fact is consistent with a model of a highly focused electric field. 72

80 5.1 Ion channels are toxin receptors Puffer fish, though a delicacy in Japan, can be lethal if not prepared properly. Japanese scientists attempting to understand the paralytic substance secreted by puffer fish discovered the first toxin - ion channel interaction [128,129]. They discovered that tetrodotoxin (TTX) blocked sodium currents in axons when only nanomolar concentrations were applied. TTX is now known to be a potent and selective blocker of the Na channel pore [130]. Homologous block of K V channels is achieved with a family of scorpion toxins called α-k-toxins [131]. These toxins are typified by charybdotoxin (CTX) and agitoxin (AgTX) from the scorpion, Leiurus quinquestriatus (Fig. 5.1a) [132,133]. These two toxins are 37 and 38 amino acid peptides, respectively, with 3 disulfide bonds stabilizing their structure (6 conserved cysteines). NMR structures have been determined (Fig 5.1b) [134,135]. Both toxins have a conserved lysine, K27, that interacts directly with the K + channel pore (Fig. 5.1c) [ ]. a b Leiurus quinquestriatus c Figure 5.1 a. Scorpion venom contains many small peptide blockers of Na +, Ca 2+, and K + channels. The first potent blockers for K V channels were isolated from the scorpion Leiurus quinquestriatus. b. NMR structures for the toxins used in our study, Agitoxin-2 (AgTX) and 73

81 Charybdotoxin (CTX). Highlighed are residues mutated to cysteine for fluorophore attachment, AgTX-D20 and CTX-R19. Also highlighted, K27, the critical lysine residue that interacts directly with the selectivity filter. c. Molecular dynamics model for AgTX bound to the Shaker channel. Figure taken from Eriksson and Roux, 2002 [138]. Therefore, these small proteins bind to the top of the channel on the central axis. Before the days of K + channel crystal structures several mutagenesis studies detailed the interaction between toxin and channel [ ]. Models have docked the toxins onto channels starting from experimental constraints and using molecular dynamics [138,143]. 5.2 LRET configuration using acceptor labeled toxin putting the paddle model to the test LRET experiments have been presented (Chapter 3.5 and Chapter 4) that were performed in an S4-donor to S4-acceptor configuration. Here we present data using an S4-donor to toxin-acceptor configuration. In this new approach, all four Shaker subunits are labeled with the lanthanide luminescent donor probe, and the acceptor dye is attached to a scorpion toxin. The toxin is either charybdotoxin (CTX-R19C [144]) or agitoxin-2 (AgTX2-D20C [145]). Procedures for labeling these toxins have been described by our collaborator Chris Miller [144]. The toxin mutations for labeling (Fig. 5.2b) are on the top side of the toxin, whereas the bottom side forms the interaction surface with Shaker. We have shown that labeling these mutant toxins with organic dyes has only a small effect on the binding affinity to the channels by qualitatively testing the toxin offrate (data not shown). We put in the mutations F425G and K427D in our Shaker construct, which greatly increases the affinity of wild-type CTX with Shaker (K i 1.5 pm [139,140]). We have shown that these mutations do not interfere with the strong binding of agitoxin-2 with Shaker (data not shown). Both of these toxins bind very strongly and specifically to the external side of the Shaker channel pore (see Fig. 5.1c), displaying a residency time of many tens of minutes [140,144]. With this geometry the acceptor is located on a stationary reference point at the very top of the channel and near the central axis. Therefore, if voltage-sensing segments are undergoing large translations 74

82 across the membrane, as the KvAP paddle model predicts, this LRET experiment will detect it. Fig. 5.2 illustrates the geometry, assuming a very conservative version of the paddle model with the paddle segments undergoing a 15 Å vertical translation, LRET between S4-donors and a toxin-acceptor show a change in distance of 10 Å. It should be noted that these values come from conservative structural estimates, and in fact the paddle model has been presented with even more extreme numbers [31]. Figure 5.2 Cartoon representation of the paddle model. LRET measures distances from donor labelled sites (blue circles) on the S3b-S4 paddle (structure taken from the isolated voltagesensor [30]) to the toxin-acceptor. The voltage-sensing arginines are shown in red. The energy transfer acceptors (green circles) are attached to the top of the channel with a scorpion toxin. The paddle model predicts a change in distance, D c - D o, of 10 Å, estimated by a conservative geometric calculation assuming a 15 Å vertical translation (red arrows). The experiments are performed as follows: Xenopus oocyte cells are injected with mrna (20 ng) for the Shaker channel (for example N353C with the F425G, K427D background) such that they over-express channels a few days post-injection. The cells are labeled with thiol-reactive Tb chelate (the donor species) such that the N353C 75