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Insights into Kv1.2 activation and deactivation using voltage clamp fluorimetry Horne, Andrew James 2010

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Insights into Kv1.2 activation and deactivation gating using voltage clamp fluorimetry by Andrew James Horne B.Sc. (Hons), The University of British Columbia, 2004  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in The Faculty of Graduate Studies (Physiology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) July 2010  © Andrew James Horne, 2010  Abstract Voltage-gated potassium (Kv) channels are essential membrane proteins in modulating membrane excitability and related cellular processes. Many details associated with the voltage response are unclear, particularly the complete role of the voltage sensing domain, and not just the densely charged S4 helix. Based on its crystal structure, the Kv1.2 channel represents an ideal model in which to study these questions. This thesis investigates Kv1.2 activation and deactivation gating, using the voltage clamp fluorimetry technique. This technique utilizes an environmentally sensitive fluorophore introduced at locations of interest in order to visualize conformational changes in protein structure. Labelling of Kv1.2 channels at the extracellular end of S4 reports a fast quenching of fluorescence emission upon depolarization that correlates extremely well with gating current measurements, suggesting it is a report of voltage-dependent S4 translocation. In addition, a slow quenching component is observed with a very negative voltage-dependence (V1/2 = -73.9 mV ± 1.4 mV), not seen in any other Kv channels studied to date, that involves regions of the voltage sensing domain in S1 and S2. This slow quenching is selectively removed from the fluorescence report with transfer of extracellular S1-S2 or S3-S4 linkers from the homologous Shaker potassium channel, suggesting that it arises from channelspecific interactions between the Kv1.2 linker segments. However, transfer of Kv1.2 linker segments into Shaker fail to recapitulate this quenching component, suggesting that these linker interactions likely underlie further differences in voltage sensor domain gating and/or structure. This slow quenching component correlates with deactivation of ionic current, and is prolonged with co-expression of the N-type inactivation-conferring Kvβ1.2 subunit. In the presence of the beta subunit, this likely reflects unbinding of the inactivation moiety from the pore domain, allowing deactivation and S4 return, but in the α-subunit alone we suggest that this may be a report of a voltage-dependent rearrangement in the voltage sensor domain that stabilizes the S4 ii  in an activated conformation. Such interactions have been reported in other voltage-gated proteins, and provides further evidence that we must consider more than just S4 translocation when it comes to understanding the complete potassium channel voltage response, Kv1.2 or otherwise.  iii  Table of Contents Abstract ........................................................................................................................................... ii Table of Contents........................................................................................................................... iv List of Tables ................................................................................................................................ vii List of Figures .............................................................................................................................. viii List of Abbreviations ..................................................................................................................... ix Acknowledgements....................................................................................................................... xii Dedication .................................................................................................................................... xiii Co-Authorship Statement............................................................................................................. xiv Chapter 1: Introduction ................................................................................................................... 1 Overview..................................................................................................................................... 2 Principles of fluorescence emission............................................................................................ 4 The physics of fluorophore emission...................................................................................... 4 Mechanism of voltage-dependent quenching in ion channels ................................................ 5 Voltage clamp fluorimetry in Shaker.......................................................................................... 8 Gating currents........................................................................................................................ 8 Subunit cooperativity ............................................................................................................ 10 Inactivation ........................................................................................................................... 12 Applying fluorescence data to the crystal structures to model activation gating ................. 15 Fluorimetry in other voltage-gated channels and proteins........................................................ 19 Kv1.5..................................................................................................................................... 19 Ci-VSP .................................................................................................................................. 20 Scope of the thesis .................................................................................................................... 21 References................................................................................................................................. 24 Chapter 2: Fast and slow voltage-sensor rearrangements during activation gating in Kv1.2 channels detected using tetramethylrhodamine fluorescence ....................................................... 29 Introduction............................................................................................................................... 30 Materials and methods .............................................................................................................. 33 Molecular biology and RNA preparation ............................................................................. 33 Oocyte preparation and injection .......................................................................................... 34 Two-electrode voltage clamp................................................................................................ 34 Voltage clamp fluorimetry.................................................................................................... 35 Gating currents...................................................................................................................... 36 Data analysis ......................................................................................................................... 36 Supplemental material .......................................................................................................... 37 Results....................................................................................................................................... 39 Kv1.2 WT channels do not show voltage-dependent changes in fluorescence emission..... 39 Kv1.2 fluorescence at A291C exhibits differences from the Shaker A359C homologue .... 39 The slow Kv1.2 fluorescence is recapitulated at other labelled sites in the S3-S4 linker .... 42 Increases in Kv1.2 fluorescence upon hyperpolarization reflect mobility of the voltage sensor .................................................................................................................................... 46 The fast and slow fluorescence phases in Kv1.2 have different voltage-dependencies ....... 49 The fast phase of fluorescence correlates with S4 movement and the Q-V relationship ..... 52 Slow changes in fluorescence recovery match the rates of Kv1.2 channel deactivation and reactivation............................................................................................................................ 54 Slow changes in fluorescence emission reflect internal rearrangement within the voltage sensor domain ....................................................................................................................... 56 iv  The S1-S2 linker and S4 region are both intricately involved in Kv1.2 activation .............. 59 Discussion ................................................................................................................................. 62 Kv1.2 A291C rapid fluorescence quenching reports fast S4 displacement during channel activation............................................................................................................................... 62 Kv1.2 A291C fluorescence also reports slower voltage-dependent rearrangements of the voltage sensor domains ......................................................................................................... 65 Slow voltage-dependent rearrangement of the voltage sensor domains is confirmed by fluorophores placed in S1-S2................................................................................................ 67 Conclusion ............................................................................................................................ 69 Acknowledgments..................................................................................................................... 70 References................................................................................................................................. 71 Chapter 3: Voltage-dependent changes in Shaker and Kv1.2 fluorescence after transfer of Kv1.2 or Kv1.5 extracellular linkers ....................................................................................................... 75 Introduction............................................................................................................................... 76 Materials and methods .............................................................................................................. 78 Molecular biology and RNA preparation ............................................................................. 78 Oocyte preparation and injection .......................................................................................... 79 Two-electrode voltage clamp................................................................................................ 80 Voltage clamp fluorimetry.................................................................................................... 80 Data analysis ......................................................................................................................... 81 Results....................................................................................................................................... 83 A homologous Shaker/Kv1.2 S4 residue produces different fluorescence emission ........... 83 Shaker and Kv1.2 extracellular linkers differ in length and sequence ................................. 85 Transfer of Kv1.2 or Kv1.5 secondary quenching events are not conferred with the S3-S4 linker alone............................................................................................................................ 85 Shaker S1-S2 linker transfer into the Kv1.2 channel prevents slow fluorescence quenching of the A291C microenvironment .......................................................................................... 90 The Kv1.2 double (S1-S2 and S3-S4) linker swap lacks a slow quenching component ...... 92 Discussion ................................................................................................................................. 94 Extracellular linkers alone cannot confer secondary fluorescence changes ......................... 94 Chimera effects on activation are likely not responsible for shifts in fluorescence ............. 94 Extracellular linker alterations and glycosylation can have significant effects on channel gating..................................................................................................................................... 95 Fluorophore quenching groups are altered by changes in linker length and structure ......... 96 The Shaker S3-S4 linker may insulate the TMRM probe from secondary changes............. 96 Kv1.2 S1-S2 linker length and/or glycosylation may confer secondary fluorescence ......... 97 S4 translocation is unaffected by linker length or structure ................................................. 98 Summary ............................................................................................................................... 98 Acknowledgements................................................................................................................... 99 References............................................................................................................................... 100 Chapter 4: The molecular basis for the actions of Kvβ1.2 on the opening and closing of the Kv1.2 delayed rectifier channel .................................................................................................. 102 Introduction............................................................................................................................. 103 Materials and methods ............................................................................................................ 106 Molecular biology............................................................................................................... 106 Oocyte preparation.............................................................................................................. 106 Two-electrode voltage clamp electrophysiology and fluorimetry ...................................... 107 Data analysis ....................................................................................................................... 107 Results..................................................................................................................................... 109 v  The Kvβ1.2 N-terminus induces fast inactivation, a negative shift of activation and slowed deactivation in Kv1.2 .......................................................................................................... 109 The voltage dependences of fluorescence deflections upon depolarization from TMRMlabelled Kv1.2 S4 mutants are unaffected by Kvβ1.2 ........................................................ 112 Slowing of both current deactivation, and fluorescence return to baseline levels upon channel closing, are induced by the Kvβ1.2 N-terminus and correlated to the proportion of channels undergoing fast inactivation................................................................................. 116 Discussion ............................................................................................................................... 120 Acknowledgements................................................................................................................. 124 References............................................................................................................................... 125 Chapter 5: Discussion ................................................................................................................. 129 Kv1.2 is expressed in excitable cells throughout the body..................................................... 130 Scientific rationale for investigating Kv1.2 gating ................................................................. 130 Kv1.2 gating currents correlate with rapid fluorescence quenching....................................... 132 Slowed fluorescence upon repolarization is consistent with delayed S4 return ..................... 133 Kv1.2 inactivation is limited, but the channel shows fluorescence consistent with a relaxed state of S4................................................................................................................................ 135 Role of the extracellular linkers in slow fluorescence-associated S4 stabilization ................ 136 Further studies needed to understand the nature of this reorganization.................................. 137 Reconciling VCF data with the Kv1.2 crystal structure ......................................................... 139 Kv1.2 activation: more than just S4?...................................................................................... 140 Summary ................................................................................................................................. 140 References............................................................................................................................... 142 Appendix A: Supplemental Material for Chapter 2.................................................................... 147 Supplemental Results.............................................................................................................. 148 Kv1.2 WT channels do not show voltage-dependent changes in fluorescence emission... 148 Substitution of the Kv1.5 S2 transmembrane segment does not affect Kv1.2 fluorescence ............................................................................................................................................. 152 Fluorescence scanning of the Kv1.2 voltage sensor reveals information about the slow fluorescence component...................................................................................................... 153 References............................................................................................................................... 155  vi  List of Tables Table 2.1. Boltzmann fits to Kv1.2 and Shaker G-V and F-V relationships from Figure 2.3..... 45  vii  List of Figures Figure 1.1. Schematic cartoon of voltage clamp fluorimetry theory. ............................................ 3 Figure 2.1. An alignment of the S3-S4 linkers of Shaker and Kv1.2 highlights similarities in the region. ........................................................................................................................................... 32 Figure 2.2. Voltage-dependent Shaker and Kv1.2 conductance and fluorescence deflections. .. 41 Figure 2.3. A scan of the Shaker and Kv1.2 S3-S4 linkers reveals differences in fluorescence phenotypes. ................................................................................................................................... 44 Figure 2.4. Holding potential affects the directionality of Kv1.2 fluorescence deflections, but does not affect the overall F-V relationship.................................................................................. 48 Figure 2.5. Kv1.2 A291C voltage-dependent fluorescence is well characterized by a double exponential function...................................................................................................................... 51 Figure 2.6. Correlation of the fast component of Kv1.2 fluorescence quenching with gating charge movement. ......................................................................................................................... 53 Figure 2.7. Slow fluorescence return upon hyperpolarization correlates with deactivation of ionic current. ................................................................................................................................. 55 Figure 2.8. Kv1.2-Kv1.5 chimera channels lack the slow fluorescence quenching at negative potentials. ...................................................................................................................................... 58 Figure 2.9. Effect of Shaker S1-S2 linker replacement on Kv1.2 fluorescence. ......................... 59 Figure 2.10. TMRM attached to residues in the S1-S2 linker of Kv1.2 report only slow changes in fluorescence emission in response to voltage. .......................................................................... 61 Figure 2.11. Proposed model of Kv1.2 activation based on TMRM fluorescence...................... 63 Figure 3.1. Shaker and Kv1.2 fluorescence from a homologous residue in the S3-S4 linker exhibit unique fluorescence effects upon depolarization.............................................................. 84 Figure 3.2. Kv1.2 slow fluorescence quenching is absent in S3-S4 linker chimera channels, and not conferred to Shaker with linker transfer. ................................................................................ 87 Figure 3.3. Secondary Kv1.5 fluorescence emission is not conferred by S3-S4 linker replacement in Shaker................................................................................................................... 89 Figure 3.4. Kv1.2 slow fluorescence quenching is lost in S1-S2 linker chimera channels, and not conferred to Shaker with linker transfer. ...................................................................................... 91 Figure 3.5. Interactions between the S1-S2 and S3-S4 linkers do not alone confer unique channel fluorescence phenotypes. .............................................................................................................. 93 Figure 4.1. Kvβ1.2 confers a spike-and-decay “fast inactivation” to Kv1.2 currents, a hyperpolarizing shift of the activation curve, and a slowing of current deactivation................. 111 Figure 4.2. Kv1.2 current and/or fluorescence kinetics are not modified by substitutions required to record fluorescence. ................................................................................................................ 113 Figure 4.3. Voltage-dependence of S4 movement upon depolarization in Kv1.2 is unaffected by Kvβ1.2. ....................................................................................................................................... 115 Figure 4.4. The voltage dependence of the fraction of the fluorescence that is slowed upon repolarization is correlated with the Kv1.2 G-V curve. .............................................................. 117 Figure 4.5. Current deactivation and the return of fluorescence to baseline are coupled and slowed by the Kvβ1.2 N-terminus. ............................................................................................. 119  viii  List of Abbreviations Amino Acid Alanine Arginine Asparagine Aspartate Cysteine Glutamate Glutamine Glycine Histidine Isoleucine Leucine Lysine Methionine Phenylalanine Proline Serine Threonine Tryptophan Tyrosine Valine  3 Letter code Ala Arg Asn Asp Cys Glu Gln Gly His Ile Leu Lys Met Phe Pro Ser Thr Trp Tyr Val  1 letter code A R N D C E Q G H I L K M F P S T W Y V  4-AP = 4-aminopyridine ΔFmax = maximum change in fluorescence τ = time constant μΑ = microampere μm = micrometer Å = Angstrom cDNA = complementary deoxyribonucleic acid CO2 = carbon dioxide cRNA = complementary ribonucleic acid Ci-VSP = Ciona intestinalis voltage sensor-containing phosphatase CNG = cyclic nucleotide-gated (channel) C-terminus = carboxy-terminus (of protein)  ix  DMSO = dimethyl sulfoxide F-V = fluorescence-voltage (relationship) Gmax = maximum conductance G-V = conductance-voltage (relationship) GFP = green fluorescent protein KCa = calcium- and voltage-activated potassium (channel) hERG = human ether a-go-go related gene (channel) H20 = water k = slope factor kHz = kilohertz LQTS = long QT-interval syndrome MΩ = megaohms MEM = minimal essential medium mg = milligram mL = millilitre mM = millimoles per liter ms = millisecond MTS = methane thiosulfate mV = millivolt Nav = voltage-gated sodium (channel) nL = nanoliter nm = nanometer N-terminus = amino-terminus (of protein) pA = picoampere PCMBS = para-chloromercuribenzene sulfonate x  PCR = polymerase chain reaction PIP2 = phosphatidylinositol 4,5-bisphosphate PMT = photomultiplier tube Q-V = gating charge-voltage (relationship) s = second S4 = fourth transmembrane segment (of voltage-gated channel) SEM = standard error of the mean TEA = tetraethylammonium TMRM = tetramethylrhodamine-5-maleimide WT = wild-type V1/2 = half-activation voltage VCF = voltage clamp fluorimetry  xi  Acknowledgements Firstly, I would like to thank my supervisor, Dr. David Fedida, for his support and assistance throughout my graduate studies. He continually pushed me beyond my comfort range, and I am all the better as a result. I would also like to thank Dr. Thomas Claydon for his inspiring enthusiasm for science, pivotal support, encouragement and friendship extending well beyond their tenure in the lab. Furthermore, I owe a large amount of gratitude to Dr. Claydon for the work he put into establishing the fluorescence program within our lab, and teaching me the basics of the technique. I am grateful to Dr. David Steele for his assistance with countless molecular biology conundrums, and to Dr. Zhuren Wang for teaching me the basics of patch clamping and for his assistance with my studies. Special thanks to Fifi Chiu and Kyung Hee Park for their cell culture expertise and general lab assistance. My graduate supervisory committee, Dr. Steven Kehl, Dr. Eric Accili and Dr. Chris Ahern has been extremely supportive and I thank them for that. Lastly, I would like to thank my friends and family for all their encouragement throughout the last six years, in good times and bad. You kept me going, and gave me a level of support you cannot begin to imagine. I would like to extend a special thank you to May Cheng for our discussions of science, her shoulder to whine on, and for sharing the adventure with me. To each and every one of you, I am truly in your debt.  xii  Dedication  To Bob, Ray and Pat  xiii  Co-Authorship Statement Chapter 2: Fast and slow voltage-sensor rearrangements during activation gating in Kv1.2 channels detected using tetramethylrhodamine fluorescence Andrew J. Horne was responsible for designing and performing most of the experiments and mutagenesis, the computer modelling (Figure 2.11) and all of the analysis presented in this chapter, except for some of the constructs presented in Figure 2.3, which were performed by Thomas W. Claydon. Christian J. Peters and Thomas W. Claydon performed some of the experiments (~10% and 5% respectively), and assisted with manuscript preparation. Andrew J. Horne and David Fedida prepared the manuscript. David Fedida, Christian J. Peters and Thomas W. Claydon edited the manuscript, and assisted with experimental design and data interpretation. A version of this chapter has been published. Andrew J. Horne, Christian J. Peters, Thomas W. Claydon, and David Fedida. (2010) Fast and slow voltage sensor rearrangements during activation gating in Kv1.2 channels detected using tetramethylrhodamine fluorescence. The Journal of General Physiology 136(1), 83-99. Chapter 3: Voltage-dependent changes in Shaker and Kv1.2 fluorescence after transfer of Kv1.2 or Kv1.5 extracellular linkers Andrew J. Horne was responsible for performing most of the experiments (~80% contribution), with the remainder performed by Christian J. Peters. Andrew J. Horne performed the analysis and prepared the manuscript. David Fedida edited the manuscript, and assisted with experimental design and data interpretation. Chapter 4: The molecular basis for the actions of Kvβ1.2 on the opening and closing of the Kv1.2 delayed rectifier channel Andrew J. Horne contributed to data collection and analysis in this study (~10% contribution). Christian J. Peters and Moni Vaid performed the rest of the experiments and analysis. The manuscript was prepared by Christian J. Peters and edited by Andrew J. Horne, Moni Vaid, David Fedida, and Eric A. Accili. A version of this chapter has been published. Christian J. Peters, Moni Vaid, Andrew J. Horne, David Fedida, and Eric A. Accili. (2009) The molecular basis for the actions of Kvβ1.2 on the opening and closing of the Kv1.2 delayed rectifier channel. Channels 3(5), 1-9. Appendix A: Supplemental material to Chapter 2 Andrew J. Horne designed and performed all of the experiments, and conducted the analysis and figure preparation. The text was written by Andrew J. Horne, and edited by David Fedida. A version of this chapter has been published online as supplemental material to the study described in Chapter 2. Andrew J. Horne, Christian J. Peters, Thomas W. Claydon, and David Fedida.  xiv  Chapter 1: Introduction  1  Overview The ability of voltage-gated ion channels to sense and modulate cellular membrane potentials, and in doing so influence essential cellular processes, underscores their importance in all physiological systems (Hille, 2001). However, many of the details of channel function remain unanswered, and are the subject of continuous study in the field.  Voltage clamp  fluorimetry (VCF) is a powerful and relatively new technique in ion channel biophysics, complementary to other electrophysiological techniques, used to understand protein conformational changes during gating. The technique utilizes a class of molecules that absorb light of a certain wavelength and emit at another, collectively referred to as fluorophores or dyes. Emitted light (fluorescence) can be influenced by any interactions with the probe resulting in a loss of energy; in other words, measured emission is highly sensitive to changes in fluorophore environment. VCF takes advantage of this by using fluorophores tethered to sulfhydryl-linker moieties and covalently attached to cysteine residues introduced at specific positions within the channel. Changes in the level of emitted light in response to depolarization provide a report on real-time changes in channel structure (Figure 1.1). In addition, changes in protein structure that occur in the absence of measurable current, and which would therefore elude electrophysiological detection, may also be measured using this technique; such changes can be elicited through drug-binding, altering pH, and many other interactions. VCF is exquisitely sensitive to local microenvironments, as even adjacent mutated and labelled amino acid residues can exhibit significantly different emission profiles during depolarization (Gandhi et al., 2000; Loots and Isacoff, 2000; Pathak et al., 2007).  2  Figure 1.1. Schematic cartoon of voltage clamp fluorimetry theory. A voltage-gated potassium channel subunit, transmembrane helices labelled S1-S6, with a fluorophore attached to an introduced cysteine residue at the external end of S4 (C), is shown in the top image. The fluorophore absorbs light of a certain wavelength (green arrows) and emits at a second, longer wavelength (red area), subject to interactions with the local environment. In response to a depolarization (ΔV), location of the fluorescence probe at a dynamic region of the protein may result in a change in the level of fluorophore emission (as seen in the bottom image) due to a change in the local microenvironment, the time- and voltage-dependence of which can be correlated to the underlying change(s) in protein conformation; see text for details. 3  Fluorescence data collected from ion channels has shed some light on the fundamental gating processes associated with channel opening and inactivation; in particular, fluorophores located in or near the pore and external end of S4 have been used to further understand how these regions move to sense voltage and modulate activation and inactivation. To date, the majority of these data have been collected in the archetypal Shaker potassium channel, although recent studies have utilized this technique in the mammalian Shaker homologue Kv1.5 channel (Vaid et al., 2008), KCa channels (Savalli et al., 2006; Savalli et al., 2007; Pantazis et al., 2010), ether-àgo-go (eag) channels (Schonherr et al., 2002; Bannister et al., 2005), human ether-à-go-gorelated gene (hERG) channels (Smith and Yellen, 2002), Nav1.4 (Cha et al., 1999; Chanda and Bezanilla, 2002; Chanda et al., 2004; Campos et al., 2007; Campos et al., 2008; Muroi and Chanda, 2009; Muroi et al., 2010), cyclic nucleotide-gated (CNG) channels (using patch clamp fluorimetry, rather than VCF) (Zheng and Zagotta, 2000), and voltage-dependent phosphatase proteins (Villalba-Galea et al., 2008). The purpose of this chapter is to review the fundamental physical principles of VCF and our current understanding of ion channel activation and inactivation from application of this technique in Shaker and other voltage-gated proteins as a background for the series of studies outlined in subsequent chapters.  Principles of fluorescence emission The physics of fluorophore emission VCF requires the use of fluorescent probes or dyes: molecules characterized by their ability to enter a transient high electron energy state upon absorption of a photon of light. The instability of this high-energy state results in the loss of some energy due to non-radiative sources of electron relaxation, followed by emission of a photon as the fluorophore sheds its remaining energy upon return to its relaxed, “ground” state. Alternatively, these final quanta of energy can be lost or stabilized through stochastic collisions with protein, other fluorophores, 4  compounds in solution, or even the solvent molecules themselves. The probability of relaxation through photon emission compared to all other sources of energy loss is referred to as the quantum yield of a fluorescent molecule, and varies as a function of environment. Voltagedependent changes in an ion channel can alter the microenvironment of a conjugated fluorophore, modifying interactions with the protein and solvent and ultimately affecting the quantum yield in a manner consistent with the time course and extent of this change. Importantly, absorption and emission occur in the femtosecond (10-15 s) and nanosecond (10-9 s) time scale, respectively, many orders of magnitude faster than the fastest known gating processes.  Thus, changes in the protein environment near the fluorophore can be accurately  tracked in real time as alterations in the quantum yield of the probe, unlimited by the kinetics of the fluorescence process. The ideal wavelengths at which absorption and emission occur depend on the unique properties of the fluorophore. For example, for tetramethylrhodamine maleimide (TMRM), a common VCF dye, maximal absorption and emission occur at 542 nm and 574 nm respectively in H2O. The rightward shift in emission wavelength relative to absorption, termed the Stoke’s shift, is observed in most fluorescent molecules and results from the lost energy from internal relaxation at excited states. In addition, emission wavelength is strongly dependent on solvent polarity, as dipole interactions with solvent molecules can further stabilize and absorb energy from the excited state. As a result, the wavelength of photon emission varies with solvent polarity, with more polar solvents shifting maximal emission to longer wavelengths (Cha and Bezanilla, 1997).  Mechanism of voltage-dependent quenching in ion channels It was shown in the early VCF papers that TMRM, or other fluorophores, attached to residues at the external end of S4 in Shaker potassium channels exhibited a voltage-dependent 5  quenching of fluorescence emission (Mannuzzu et al., 1996; Cha and Bezanilla, 1997). While it was initially tempting to believe these decreases in fluorescence emission were due to outward S4 translocation and subsequent displacement of TMRM from the hydrophobic lipid bilayer into a polar extracellular solution (Mannuzzu et al., 1996), aqueous accessibility to TMRM attached in S4 is observed at both resting and activated potentials (Cha and Bezanilla, 1997; Cha and Bezanilla, 1998) with only a small degree of voltage-dependence.  Furthermore, spectral  measurements of TMRM fluorescence when attached to M356C or A359C at the outer end of S4 had a peak emission wavelength approximating that of TMRM in water alone, and was also voltage-independent (Cha and Bezanilla, 1997). Thus, fluorophore environment appears to be largely aqueous at all potentials with some moderate interactions with polar molecules aside from water, perhaps peripheral amino acid residues.  Importantly, changes in aqueous  accessibility cannot completely explain the voltage-dependent quenching observed upon depolarization. Agitoxin and external TEA effects on the M356C-TMRM fluorescence profile (Cha and Bezanilla, 1998) at first glance appeared to indicate that ion flux may affect fluorescence emission, and perhaps contribute to the mechanism of voltage-dependence. However, observed changes did not correlate on the same time scale to changes in ionic current or reflect the reversal of current following rapid hyperpolarization; rather than a directional switch in emission, a slow decay to baseline was observed. In addition, the voltage-dependence of fluorescence at most labelled sites in S4 or the pore regions does not correlate with conductance. Thus, it seems highly unlikely that ionic current itself represents a significant component of the voltagedependent quenching observed with fluorophores in the external milieu. Originally, it was suggested that changes in the fluorescence emission during depolarization might also be influenced by the inherent chemical-specific properties of the fluorescence dye itself, or by interactions between the fluorophores themselves, in the absence of 6  interactions with environmental constituents.  However, fluctuations in dipole orientation  (anisotropy) during depolarizing pulses are not responsible for observed fluorescence, based on identical voltage-dependence in the presence of multiple polarizing filter orientations (Cha and Bezanilla, 1998). Furthermore, striking differences in the directional change in fluorescence emission of different probes attached to the same residue (i.e. increased emission with some probes, and decreased emission with others) argue against any mechanism of fluorescence emission that would be expected to be identical in all probes, such as voltage-dependent dipole changes and dimer formation, or transfer of photon energy to a nearby fluorophore (Cha and Bezanilla, 1997). In addition, fluorescence emissions in response to voltage changes have been recorded with channels in which only one subunit has an externally available cysteine, and found to be qualitatively identical to channels in which all four subunits are labelled (Loots and Isacoff, 1998). Thus, experimental data appears to suggest that the primary changes in fluorescence emission arise from some degree of fluorophore interaction with the environment, rather than being limited to changes within the probe itself. From these fundamental experiments described above, voltage-dependent fluorescence observed at external Shaker residues cannot be completely explained by dye-specific properties, changes from a hydrophobic to polar environment, and do not correlate with the movement of ions through the pore. Rather, voltage-dependent changes in fluorescence emission appear to be explained best by differential collisional quenching from molecules in solution and the solvent itself, as well as through interactions with amino acids within the probe’s microenvironment. This conclusion is strengthened by studies in which fluorescence changes are recorded from residues in the voltage-sensor of Shaker channels in which the majority of the S3-S4 linker has been removed. With the Shaker linker reduced from ~31 amino acids to 5, a fluorophore at residue 356 showed a 10-fold decrease in quenching of the fluorescent signal, and lost its pHdependence (Sorensen et al., 2000). This experiment suggests that such protein-fluorophore 7  interactions are a key component of the voltage-dependent emission decrease, and that the fundamental mechanism of voltage-dependent changes in fluorophore emission is related to the change in interactions between the fluorophore and these nearby quenching groups as they approach or diverge from each other in a gating-dependent manner.  Voltage clamp fluorimetry in Shaker Gating currents Since it was first cloned, the Drosophila Shaker potassium channel (Papazian et al., 1987), has been regularly used as the archetype of voltage-gated K+ (Kv) channels. Based on the large existing body of knowledge for all aspects of gating in this channel, pioneering studies in VCF relied heavily on the use of Shaker in correlating the kinetic behaviour of emission changes with gating. The first published VCF studies used TMRM fluorophores attached to multiple residues at the external end of S4 in fast inactivation removed (due to the deletion of the Nterminal inactivation moiety, residues 6-46) Shaker-IR channels to investigate local changes in the S4 environment during activation (Mannuzzu et al., 1996). Labelling of residues in the S3S4 linker, as a measure of solvent accessibility, was accomplished efficiently at both hyperpolarized and depolarized potentials. Residues closer to, and within, S4 were less likely to be labelled at hyperpolarized potentials depending on their position within the α-helix, in similar agreement with studies using methanethiosulfonate (MTS) (Larsson et al., 1996; Baker et al., 1998) or parachloromercuribenzene sulfonate (PCMBS) (Yusaf et al., 1996; Wang et al., 1999) reagents. Once labelled, deflections in fluorescence emission from TMRM attached to A359C, M356C and other residues in the region exhibited voltage-dependencies which overlaid with the charge-voltage relationships, suggesting that an outward translocation of the S4 helix could be measured by both changes in fluorescence emission and channel gating currents. Modulation of gating charge movement with the L382V background mutation (Lopez et al., 1991) resulted in 8  parallel changes in the fluorescence-voltage relationship, further supporting the authors’ conclusion. Similar studies conducted by Bezanilla’s group differed slightly in that Shaker-IR M356C and A359C showed peak fluorescence-voltage (F-V) relationships left-shifted by 15-20 mV compared with the charge-voltage (Q-V) relationship (Cha and Bezanilla, 1997). However, there was a clear temporal correlation between the fluorescence and gating currents, as the time constant of the slow component of gating charge movement for M356C overlaid the major component of fluorescence decay, and A359C fluorescence was less shifted from the Q-V relationship with longer depolarizing pulses, which suggested that the fluorescence provides a valid report of S4 movement (Cha and Bezanilla, 1997). Temporal correlation of fluorescence decay and gating charge can also be shown by superimposing the fluorescence record for a given depolarizing stimulus with the integral of gating current for that pulse, which represents total gating charge. While these records do not always precisely overlap, they present a strong case for fluorescence reporting on aspects of gating charge movement across the membrane; indeed, other reports have shown a closer relationship between these two measurements (Gandhi et al., 2000). In another study, mutation of R365 to a cysteine or serine resulted in a separation of gating charge movement into two components Q1 and Q2. Channels labelled with TMRM at S352C with the R365S mutation reflected this separation in the biphasic fluorescence report (Baker et al., 1998). However, the F-V relationship still did not overlap the Q-V. Thus, the evidence appears to support a close correspondence between gating charge movement and the time course and voltage-dependence of fluorescence emission from the top of S4, although it is not always perfect as initially proposed (Mannuzzu et al., 1996). Given that the fluorescence report is specific to the microenvironment of the fluorophore, while gating current represents the summation of all charged particle movement within the channel, it is not unrealistic to expect that these relationships may deviate from one another. The S2 and S3 9  segments of the voltage-sensor have also been implicated in contributing to gating charge movement (Papazian et al., 1995; Planells-Cases et al., 1995; Seoh et al., 1996), which may not be sensed by a fluorophore in S4. TMRM attached at residue T276C in the Shaker S1-S2 linker showed a voltage-dependent decrease in fluorescence that preceded the Q-V relationship, and which decayed with kinetics faster than the increase in integrated gating charge (Cha and Bezanilla, 1997). The response was maximal at -40 mV, leading the authors to suggest that properties of the fluorescence profile of T276C match the electrophysiological characterization of the Q1 component of gating charge, and suggest that early movement of the S2 helix may be responsible for this component, followed by S4 movement that correlates well with Q2 movement (Cha and Bezanilla, 1997). This is in contrast to the conclusions of Baker et al. (1998), who determined that Q1 and Q2 both described aspects of S4 movement. However, these data do not necessarily have to be in disagreement; voltage-dependent changes in fluorescence arise from changes in fluorophore environment, which may mean movement within the protein moiety linked to the probe itself, or the surrounding region relative to the fluorophore. It may be possible that the TMRM attached to T276C exclusively reports early S4 movement associated with the early stages of activation, while later activating steps (i.e. Q2 charge movement) are outside the local microenvironment of the probe.  In contrast, a  fluorophore attached to S4 may detect all aspects of movement associated with activation and report on multiple phases of the conformational change.  Subunit cooperativity VCF can also be used to investigate the cooperative gating movements associated with channel opening, distinct from the movements of the voltage sensor. By co-injecting oocytes with carefully calculated ratios of Shaker subunits containing an engineered cysteine near S4 (S352C) in the background of the gating-altering R365S mutation (Wang et al., 2000a), and 10  either WT or R365S mutant subunits without labellable cysteine residues, Mannuzzu et al. were able to investigate the effects of subunit-subunit interactions on movement of the labelled voltage sensor (Mannuzzu and Isacoff, 2000). The subunit ratios were selected so that the majority of functional channels would have only one subunit with the externally available cysteine (S352C), tracking the movement of only that voltage sensor and its environment. If S4 movement occurred independently of the influence of other subunits, the fluorescence would be expected to be identical regardless of the identity of the associated subunits, as the labelled subunit is the same in both channels. In fact, differences were clearly observed in S352C fluorescence-voltage relationships when co-expressed with WT or R365S subunits and in a second set of experiments with another gating altering mutation (L382V), particularly in the second component of gating charge movement, despite the invariant nature of the labelled subunit (Mannuzzu and Isacoff, 2000). Based on these data, the authors were able to confirm independent and cooperative phases of S4 movement in a single subunit. Cooperative S4 movement and channel opening was explored further by looking at the effects of an ILT triple mutation on Shaker fluorescence (Pathak et al., 2005). The ILT construct comprises a group of three semi-conservative hydrophobic mutations in S4 that have been shown to disrupt interactions between S4 and the pore, destabilizing S4 and shifting the Q-V relationship to more negative potentials, while shifting the final steps of activation that lead to pore opening to more positive potentials (Ledwell and Aldrich, 1999). The F-V relationship of ILT channels measured from TMRM at A359C showed a double-Boltzmann relationship with components correlating to these different gating charge components, indicating that the fluorophore was reporting on two very distinct phases associated with channel opening (Pathak et al., 2005). Hetero-tetrameric channels with only one labelled WT or ILT subunit were assayed for evidence of cooperativity in the form of mutant influences on WT subunits, or vice versa. Labelled ILT subunits in association with three unlabelled WT subunits exhibited an F-V 11  relationship, fit with a single Boltzmann equation, which overlaid well with only the first phase of the ILT homotetramer. This suggested that cooperative interactions of the ILT subunit with the WT subunits had facilitated the secondary phase of ILT charge movement and prevented the dissociation of the two voltage-dependent conformational changes (Pathak et al., 2005). The opposite effect was seen when WT subunits were labelled and co-assembled with ILT subunits. These data show how the VCF technique can be used to investigate not only the individual aspects of S4 movement as it relates to gating charge displacement, but also to shed light on S4 interactions with neighboring subunits that underlie the cooperative gating steps that lead to channel opening.  Inactivation As discussed, VCF has been used to label residues in and around S4 to understand gating movements associated with activation. However, prolonged depolarizations result in slower further changes in fluorescence quenching of fluorophores attached to the voltage sensor domain. These changes have been better shown with fluorophores attached in the external pore domain of Shaker channels. For example, TMRM-labelled S424C mutant channels show predominantly slow changes in fluorescence emission with long depolarizing pulses (Loots and Isacoff, 1998), with a time course correlating extremely well with the time constant for ionic current inactivation. Interventions affecting the progression of inactivation including alteration of the external potassium concentration (Lopez-Barneo et al., 1993; Baukrowitz and Yellen, 1995; Kiss and Korn, 1998) and pH (Kehl et al., 2002; Starkus et al., 2003) affected ionic current and fluorescence equally (Loots and Isacoff, 1998), providing strong evidence that this slow change in fluorescence emission reported the induction of slow inactivation (Loots and Isacoff, 1998). Furthermore, the voltage-dependence of fluorescence associated with this process is hyperpolarized with respect to conductance (Loots and Isacoff, 1998), which supports previous 12  ideas that inactivation may occur from both open and closed states (Kurata et al., 2005; Kwan et al., 2006). The time constant for the returning fluorescence component during repolarization (2.4 s) is also similar to that of ionic current recovery from inactivation, suggesting that a probe at this location reports well on multiple aspects of the process. More recent studies have reported a slow quenching component to S4 fluorescence with TMRM-labelled A359C residues, during depolarizations as short as 100-200 ms (Claydon et al., 2007a). Although the contribution of this slow component to the total fluorescence was small, it suggested that changes associated with slow inactivation could be initiated very quickly after activation. Longer depolarizations resulted in a more prominent slow quenching component at A359C similar to that observed at S424C (Claydon et al., 2007a), suggesting that changes related to slow inactivation are sensed by both voltage sensor and pore fluorophores, and therefore may involve both domains. Studies with 4-AP, a drug shown to bind in the internal pore cavity and prevent the opening of the intracellular gate normally coupled to S4 movement (McCormack et al., 1994; Armstrong and Loboda, 2001), prevented the slow fluorescence component in both S4 and pore reports. This agrees with studies suggesting that the fast fluorescence reports on gating charge movement, as both rapid fluorophore quenching (Claydon et al., 2007a) and the movement of early gating charge (McCormack et al., 1994) have been shown to be unaffected by 4-AP binding. Additionally, opening of the S6 activation gate (or possibly the late component of gating charge blocked by 4-AP) (McCormack et al., 1994) precipitates changes in the outer pore leading to slow activation, which are prevented by 4-AP binding (Del Camino et al., 2005; Claydon et al., 2007a). Also observed in the S424C fluorescence signal was a small, fast quenching component that represented approximately 10-15% of the total signal (Loots and Isacoff, 1998). This component increased in magnitude during paired-pulsing, and was suggested to be a report of S4 movement that was detected more efficiently from channels inactivated during the prepulse. 13  Permanently inactivated W434F channels (Perozo et al., 1993) labelled at S424C also showed only a fast fluorescence quenching upon depolarization (Loots and Isacoff, 1998), and thus it was suggested that ionic current inactivation caused alterations in the external environment of the pore with respect to the S4 segment that allowed TMRM attached at S424C to sense rapid conformational changes associated with voltage sensor movement. This reorientation of the outer pore structure is consistent with previously detailed inactivation processes in this region, and classified as P-type inactivation (DeBiasi et al., 1993). After much longer depolarizations, the off-fluorescence report showed a slower voltagedependent decay back to baseline, which was also observed in the W434F mutant. It was suggested that this process represented an additional inactivation process which shared a number of properties with the well-known C-type inactivation (Yang et al., 1997; Olcese et al., 1997; Loots and Isacoff, 1998).  From these VCF data, a model was proposed whereby Shaker  inactivation occurred in two steps. An initial occlusion of the outer pore occurs that alters the relationship between the pore and voltage sensor, and a second phase involves the stabilization of the voltage sensor in an outward position; the two phases are referred to as P-type and C-type inactivation respectively (Loots and Isacoff, 1998). From these data, it is clear that interactions between S4 and the pore modulate channel gating, potentially linking activation to inactivation through interactions in the outer pore. This coupling between voltage sensor movement and slow inactivation appears to involve a movement of the S4 helix, as fluorescence reports from 4 contiguous residues in the S4 helix all show some degree of a slow fluorescence decay (Loots and Isacoff, 2000). If the conformational changes were occurring near the helix, only a certain side would likely report on these conformational changes. Residues in the peripheral region of the external pore (E418C and A419C) also show some fast fluorescence reports, indicative of proximity to the S4 segment even in the closed state. This region appears to be at least partly responsible for coupling S4 14  movement to activation, as the E418Q mutation removes the slow fluorescence component of emission from probes located in both the pore (S424C) and S4 helix (A359C). As a model of Ptype inactivation, Loots and Isacoff (2000) propose that hydrogen bonds between E418 and residues in the outer P-loop are destabilized by S4 approach during activation (Elinder et al., 2001), which propagates to the selectivity filter through disruption of W434 interactions with the selectivity filter backbone. The resultant loss of the supporting framework conferring pore rigidity produces a non-conducting channel. In addition, such a mechanism serves to explain the key role of W434 in maintaining pore stability, as evidenced by the lack of conduction in mutant channels (Perozo et al., 1993; Yan et al., 1996). Studies of pH-dependent modulation of Shaker gating have also provided insight into inactivation processes in potassium channels (Claydon et al., 2007b).  A reduction of  extracellular pH from 7.5 to 4.0 drastically altered the fluorescence profile, in concert with a depolarizing shift in conductance- and charge-voltage relationships. High external potassium prevented changes in A359C and S424C fluorescence during depolarization, and changes in the level of baseline fluorescence emission at -80 mV were shown to be strongly dependent on pH, suggesting that inactivation-dependent rearrangements of external Shaker structure can occur from both open and closed states (Claydon et al., 2007b). It appears that low extracellular pH can enhance inactivation both at rest and during depolarization, through localized changes in or around the S4 helix.  Applying fluorescence data to the crystal structures to model activation gating With crystallization of the internal pH-activated KcsA potassium channel (Doyle et al., 1998) and more recently the mammalian Shaker-homologue, Kv1.2 channel (Long et al., 2005a; Long et al., 2005b), VCF data can be interpreted using these structural models to give a dynamic picture of ion channel gating during both activation and inactivation. Voltage-dependent probes 15  attached throughout both the Shaker voltage sensor and pore domains appear to report on conformational changes associated with not only movement in their own region, but that of their complementary domain as well. This is not unexpected, as the two domains are in close proximity to one another; crystallographic data shows that the voltage sensor of one subunit associates with the pore domain of another subunit (Long et al., 2005a; Long et al., 2005b; Long et al., 2007), a fact that may facilitate the cooperative interactions previously investigated with single-subunit fluorescence (Mannuzzu and Isacoff, 2000; Pathak et al., 2005). To further our understanding of channel gating during activation, Gandhi et al. (Gandhi et al., 2000) conducted a TMRM fluorescence scan of the S3-S4 linker and top half of S4 in Shaker-IR W434F, as well as several non-contiguous residues in the peripheral pore region, after consideration of the KcsA crystal structure (Doyle et al., 1998) which was the best available model at the time. Measuring fluorescence movement in response to depolarizations on the order of seconds, they took fast deflections to indicate proximity to S4 activation, and slow movements as a record of pore changes during inactivation (Gandhi et al., 2000). Labelled residues in the pore region all showed some degree of slow movement consistent with a report of slow inactivation (Loots and Isacoff, 1998; Loots and Isacoff, 2000). In addition, a few amino acids reported fast changes in fluorescence on the same time scale as activation; because these were not globally sensed throughout the pore, they likely represent S4 movement (Gandhi et al., 2000). When these residues were mapped onto the KCsA crystal structure, they aligned as a narrow band of residues in the peripheral pore region, suggesting a path for S4 movement parallel to these pore residues (Gandhi et al., 2000). Similarly, residues in S4 were classified as to whether fluorescence reports exhibited primarily fast changes, primarily slow changes, or an approximately equal combination of both. These changes in emission could not be correlated to the KcsA structure, as there is no voltage sensing domain in this protein.  Rather, the authors used an α-helical model of the S4 16  transmembrane segment, and found that residues reporting on fast fluorescence changes were located in the outer third of the tested segment (N-terminal from S352 in the S3-S4 linker), suggesting that activation and the rapid outward movement of S4 displaces these residues and prevents them from sensing slow changes associated with inactivation. However, hysteresis in return fluorescence upon repolarization suggested that these fluorophores instead experience the altered channel environment upon return to the resting state. The middle third of the scanned segment (between N353 and A359) consists of residues displaying both fast and slow components, indicating that while they too are subjected to outward displacement (as reported by the small fast components), they are still close enough to the pore and can sense conformational changes associated with inactivation. The innermost third of the residues, C-terminal to I360 in S4, exhibit predominantly slow reports on movement. This may indicate that the outward displacement does not significantly alter the environment of these probes, but changes in the pore structure are sensed. While there are exceptions to these generalizations (for example, L358 reports only on only fast changes in emission, and both R362 and R365 in S4 show a combination of fast and slow fluorescence changes), the results allowed the authors to infer some structural aspects of S4 during gating (Gandhi et al., 2000). For example, the approximate boundary between the mainly slow fluorophores and the mixed (fast and slow fluorescence) reporters, residues R362 to R365, delimits the range of the aqueous cavity during channel activation as determined from aqueous accessibility studies (Larsson et al., 1996; Yusaf et al., 1996), suggesting that the innermost residues of this scan remain buried during depolarization. This could of course explain the lack of environmental change during S4 translocation, in contrast to the rapid quenching observed with more external, aqueous accessible residues. Based on these data, a model was constructed where depolarization induces rapid outward S4 movement in a direction towards the central axis of the pore, relative to a band of residues within the peripheral pore region (Gandhi et al., 2000). 17  This movement brings the extracellular end of S4 to within 20 Å of the central axis, a movement of approximately 10 Å from its resting position. Maintained depolarization, which fluorescence clearly shows to involve global rearrangements of both the pore and voltage-sensing domains, leads to channel alteration, which affects the kinetics of S4 return fluorescence. When the published crystal structure of Kv1.2 (Long et al., 2005a; Long et al., 2005b) placed the S4 helix near the pore domain, as suggested by the previous fluorescence scan, a second study by the same lab re-interpreted their fluorescence data within the new model (Pathak et al., 2007). This study included a more extensive fluorescent scan, including residues in the extracellular regions of all transmembrane helices (56 residues total). Among other things, this addressed the shortcoming of using the 2-transmembrane KCsA channel to model 6transmembrane Kv channel voltage-sensor interactions with the pore. Fluorescence movements were detected from residues in the extracellular regions of all 6 helices, but as a general rule a greater success rate was obtained for S3-S6, indicating that these moieties undergo more movement than S1 and S2 (Pathak et al., 2007). Radial distribution of fluorescent reports around S3 suggests that, as opposed to remaining stationary during gating and reporting on S4 movement, it likely moves alongside S4. Rather than predicting a paddle model of voltagesensing (Jiang et al., 2003a), the authors instead suggest some level of cooperativity between the S3 and S4 helix, as fluorescence reports of residues along the S3-S4 interfacing regions indicate some relative movement between the helices in contrast to the predicted fixed paddle structure (Jiang et al., 2003b). This coincides with previous VCF studies also refuting the paddle model of movement, where residues in the S3-S4 linker were labelled by membrane-impermeant fluorophores at both positive and negative potentials (Gandhi et al., 2003), arguing against a large transmembrane movement. In order to address the primary question of how far the voltage sensor may move during activation, Pathak et al. chose to focus on the fast components of all recorded fluorescence 18  traces, which they correlated with activation. Normalizing the absolute magnitude of the fast change in fluorescence intensity to the baseline level of emission prior to any voltage-dependent changes was used as a measure of the extent to which the fluorophore (and, consequently, the residue to which it is attached) moved, assuming that the entire range of movement during this period was reported by changing fluorescence emission. From these data, they suggested that depolarization of Shaker produces a 6-8 Å outward displacement of the S4 helix, along with a 180° rotation (Pathak et al., 2007). Greater deflections in S4 than S3, combined with minimal movement of the helical portions of S1 and S2, led to a model of S4 movement relative to the outer pore that accounts for fluorescence data as well as what is already known about channel gating. However, it is worth noting that although this prediction is derived from computer modelling onto the Kv1.2 crystal structure model, the fluorescence data obtained from Shaker forms the fundamental basis of these conclusions, and thus an underlying assumption of this work is that the two channels are similar in both structure and gating.  Fluorimetry in other voltage-gated channels and proteins Kv1.5 The Kv1 family of voltage-gated potassium channels was so classified based on its sequence similarity to the archetypal Shaker potassium channel. Given the familial resemblance in this respect, it would be reasonable to expect similar characteristics of the voltage-dependent fluorescence emissions. Fluorescence has been recorded from S4 residues homologous between several members of the Kv1 family, including the first published report of mammalian Kv1 fluorescence (Vaid et al., 2008). While in each channel, S4 fluorescence shows fast fluorescence movements immediately following depolarization, subsequent changes in fluorophore quenching differ between the family members. Our understanding of the mechanisms underlying this phenomenological difference is limited, but work on Kv1.5 fluorescence suggests that the de19  quenching (increase) of emission following the peak represents a secondary report on conformations associated with channel opening (Vaid et al., 2008), as interventions that alter pore structure or coupling to the voltage sensor domain (including 4-AP, the ILT triplet mutation in S4, and increasing external potassium) all prevent this component. Inconsistencies with typical fluorescence quenching seen during inactivation, as well as the persistence of this rapid component in the background of R487V, the analogous mutation to the slow inactivationinhibiting T449V mutation in Shaker (Lopez-Barneo et al., 1993), that in Kv1.5 has been shown to abolish inactivation in Na+-conducting channels (Wang et al., 2000b), argue against a report on slow inactivation. Nonetheless, it does appear to be a report of some voltage-dependent conformational change in the pore domain, based on the aforementioned interventions, as well as its presence in multiple residues along a single face of the S4 helix (Vaid et al., 2008).  Ci-VSP In addition to confirming that the VSD can exist in a membrane without support from an ancillary pore domain, the discovery of a voltage sensor-linked phosphatase enzyme isolated from Ciona intestinalis (Ci-VSP) (Murata et al., 2005) provided a novel opportunity to investigate voltage sensor function in a homologous structure to Kv voltage sensing domains, in the absence of processes associated with pore conductance such as activation and inactivation. Utilizing VCF, it has been discovered that the voltage sensor undergoes two separate, independent changes in conformation during depolarization (Villalba-Galea et al., 2008). A fast change in emission correlates well with the time integral of gating charge movement, as seen in Shaker channels. In addition, a second component to fluorescence, slow in both onset and recovery, reports on secondary changes in conformation that reflect changes in the Q-V relationship, akin to those observed in Shaker in response to positive holding potentials (Olcese et al., 1997). The time course of this slow fluorescence change correlates extremely well with 20  the voltage-dependent shift in the V1/2 of the Q-V relationship as the holding potential is made more positive, which the authors suggested reflects the stabilization of gating charge in an activated position, or so-called relaxed state (Villalba-Galea et al., 2008). This state results from activation of gating charge and a short-lived occupancy of this activated state before transitioning to this more stable relaxed conformation, which may involve conversion of S4 from a resting 310 helical conformation to an α-helical relaxed state. Such a conversion between S4 helical conformations has also been hypothesized to occur in Kv channels, based on the crystal structure of the Kv1.2/2.1 chimera channel (Long et al., 2007).  Scope of the thesis The main aim of this thesis is to investigate Kv1.2 activation and deactivation gating, and compare our findings to the gating of the Kv1 archetypal Shaker channel, using VCF. Attempts to distill fluorescence data collected in Shaker channels into meaningful interpretations of channel gating often rely heavily on the crystal structure of Kv1.2 (Long et al., 2005a; Long et al., 2007) as a reference point, given their similar homology and perceived similar mechanisms of function. While it is generally assumed that data collected in Shaker can be applied to our understanding of related voltage-gated potassium channels, there are important differences in the gating, structure and function of Kv1.2 (Ishii et al., 2001; Gutman et al., 2003; Rezazadeh et al., 2007). Regions of the protein thought to be important for conferring voltage-dependent changes in fluorescence emission, for example the extracellular linker segments, are quite different in both structure and length. In addition, Kv1.2 does not appear to undergo significant inactivation of ionic current, even with long depolarizations (Paulmichl et al., 1991). Thus, it is possible that real-time fluorescence data collected in Shaker may be poorly matched with the Kv1.2 crystal structure as a consequence of different processes within the two channels.  A detailed  understanding of Kv1.2 fluorescence is important in the hopes that some correlation with the 21  crystal structure can further unlock the mechanics of ion channel function, and to compare with Shaker fluorescence.  In the first set of experiments we determined that Kv1.2 fluorescence  differs from Shaker at select locations in S4, underlying potentially unique voltage-dependent changes in the environment of this region during activation.  Specifically, slow rearrangements  of the protein not seen in any other Kv channel, initiated at negative potentials, appear to modulate voltage sensor mobility, as the reversal of these movements correlates with the deactivation of ionic current.  Through fluorophore placement within the S1-S2 linker, and  chimera channels in which this region was replaced with either Kv1.5 or Shaker, we implicate this region as being important for interacting with S4 and modulating this process. We further investigate the origin of this slow secondary movement from a structural viewpoint, investigating the role of extracellular linker structure in Kv1.2, Kv1.5 and Shaker fluorescence. In this series of experiments, we analyze the role of the S1-S2 and S3-S4 linkers on these important differences in fluorophore environment through the use of chimera channels. From the fluorescence reports of the different chimeras, we determine that the unique secondary changes in Kv1.2 and Kv1.5 emission are complex and not transferred with individual linker segments. Furthermore, insertion of the Shaker S3-S4 linker into Kv1.2 precludes any report of a slow fluorescence change. However, S4 translocation is well reported in all chimera channels regardless of S1-S2 and S3-S4 linker length or structure. We suggest that the linkers may strongly influence the environment of a probe in S4, but that the ultimate underlying changes in protein structure may depend heavily on the identity of the channel itself. The final set of experiments investigates the role of beta subunit modulation on Kv1.2 gating. We found no change in fast or slow fluorescence emission, or the fluorescence-voltage relationship, from fluorophores attached to S4 in the presence of the Kvβ1.2 subunit despite a leftward shift of conductance. This suggests that voltage sensor activation is unaffected by beta subunit modulation, and that the shift in conductance must instead be attributed instead to 22  voltage-dependent block of the pore. S4 return, as measured by VCF, was delayed in the presence of Kvβ1.2 with a time course of recovery similar to ionic current deactivation, providing evidence that pore occlusion by the N-terminal region of the auxiliary protein, by preventing activation gate closure, inhibits the return of gating charge similar to the effect(s) observed in Shaker with the endogenous α-subunit N-terminal inactivation moiety (Bezanilla et al., 1991).  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Phosphoinositide phosphatase activity coupled to an intrinsic voltage sensor. Nature 435:1239-1243. Muroi, Y., M. Arcisio-Miranda, S. Chowdhury, and B. Chanda. 2010. Molecular determinants of coupling between the domain III voltage sensor and pore of a sodium channel. Nat Struct Mol Biol 17:230-237. Muroi, Y. and B. Chanda. 2009. Local anesthetics disrupt energetic coupling between the voltage-sensing segments of a sodium channel. J Gen Physiol 133:1-15. 26  Olcese, R., R. Latorre, L. Toro, F. Bezanilla, and E. Stefani. 1997. Correlation between charge movement and ionic current during slow inactivation in Shaker K+ channels. J Gen Physiol 110:579-589. Pantazis, A., V. Gudzenko, N. Savalli, D. Sigg, and R. Olcese. 2010. Operation of the voltage sensor of a human voltage- and Ca2+-activated K+ channel. Proc Natl Acad Sci U S A 107:4459-4464. Papazian, D. M., T. L. Schwarz, B. L. Tempel, Y. N. Jan, and L. Y. Jan. 1987. Cloning of genomic and complementary DNA from Shaker, a putative potassium channel gene from Drosophila. Science 237:749-753. Papazian, D. M., X. M. Shao, S. A. Seoh, A. F. Mock, Y. Huang, and D. H. Wainstock. 1995. Electrostatic interactions of S4 voltage sensor in Shaker K+ channel. Neuron 14:12931301. Pathak, M., L. Kurtz, F. Tombola, and E. Isacoff. 2005. The cooperative voltage sensor motion that gates a potassium channel. J Gen Physiol 125:57-69. Pathak, M. M., V. Yarov-Yarovoy, G. Agarwal, B. Roux, P. Barth, S. Kohout, F. Tombola, and E. Y. Isacoff. 2007. Closing in on the resting state of the Shaker K+ channel. Neuron 56:124-140. Paulmichl, M., P. Nasmith, R. Hellmiss, K. Reed, W. A. Boyle, J. M. Nerbonne, E. G. Peralta, and D. E. Clapham. 1991. Cloning and expression of a rat cardiac delayed rectifier potassium channel. Proc Natl Acad Sci USA 88:7892-7895. Perozo, E., R. MacKinnon, F. Bezanilla, and E. Stefani. 1993. Gating currents from a nonconducting mutant reveal open-closed conformation in Shaker K+ channels. Neuron 11:353-358. Planells-Cases, R., A. V. Ferrer-Montiel, C. D. Patten, and M. Montal. 1995. Mutation of conserved negatively charged residues in the S2 and S3 transmembrane segments of a mammalian K+ channel selectively modulates channel gating. Proc Natl Acad Sci U S A 92:9422-9426. Rezazadeh, S., H. T. Kurata, T. W. Claydon, S. J. Kehl, and D. Fedida. 2007. An activation gating switch in Kv1.2 is localized to a threonine residue in the S2-S3 linker. Biophys J 93:4173-4186. Savalli, N., A. Kondratiev, S. B. de Quintana, L. Toro, and R. Olcese. 2007. Modes of operation of the BKCa channel β2 subunit. J Gen Physiol 130:117-131. Savalli, N., A. Kondratiev, L. Toro, and R. Olcese. 2006. Voltage-dependent conformational changes in human Ca2+- and voltage-activated K+ channel, revealed by voltage-clamp fluorometry. Proc Natl Acad Sci U S A 103:12619-12624. Schonherr, R., L. M. 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Voltage clamp fluorimetry reveals a novel outer pore instability in a mammalian voltage-gated potassium channel. J Gen Physiol 132:209-222. Villalba-Galea, C. A., W. Sandtner, D. M. Starace, and F. Bezanilla. 2008. S4-based voltage sensors have three major conformations. Proc Natl Acad Sci U S A 105:17600-17607. Wang, M. H., U. Oh, and H. I. Rhee. 2000a. Amino acid substitution within the S2 and S4 transmembrane segments in Shaker potassium channel modulates channel gating. Biochemical and Biophysical Research Communications 275:720-724. Wang, M. H., S. P. Yusaf, D. J. S. Elliott, D. Wray, and A. Sivaprasadarao. 1999. Effect of cysteine substitutions on the topology of the S4 segment of the Shaker potassium channel: implications for molecular models of gating. J Physiol 521:315-326. Wang, Z. R., X. Zhang, and D. Fedida. 2000b. Regulation of transient Na+ conductance by intraand extracellular K+ in the human delayed rectifier K+ channel Kv1.5. J Physiol 523:575-591. Yan,Y., Y.Yang, and F.J.Sigworth. 1996. How does W434F block Shaker channel current? Biophys J 71:A190 (Abstr.). Yang, Y. S., Y. Y. Yan, and F. J. Sigworth. 1997. How does the W434F mutation block current in Shaker potassium channels? J Gen Physiol 109:779-789. Yusaf, S. P., D. Wray, and A. Sivaprasadarao. 1996. Measurement of the movement of the S4 segment during the activation of a voltage-gated potassium channel. Pflugers Arch 433:91-97. Zheng, J. and W. N. Zagotta. 2000. Gating rearrangements in cyclic nucleotide-gated channels revealed by patch-clamp fluorometry. Neuron 28:369-374.  28  Chapter 2: Fast and slow voltage-sensor rearrangements during activation gating in Kv1.2 channels detected using tetramethylrhodamine fluorescence1  1  A version of this chapter has been published. Andrew J. Horne, Christian J. Peters, Thomas W. Claydon, and David Fedida. (2010) Fast and slow voltage sensor rearrangements during activation gating in Kv1.2 channels detected using tetramethylrhodamine fluorescence. The Journal of General Physiology 136(1), 83-99.  29  Introduction Voltage-gated potassium channels, or Kv channels, comprise a large class of membranespanning proteins involved in mediating the recovery of excitable cells from depolarizing stimuli. Because of recent success in crystallizing various forms of potassium channels, they have been used as structural models for many other channels. Crystal structures from Kv1.2 (Long et al., 2005a; Long et al., 2005b) and a Kv1.2/Kv2.1 chimera channel (Long et al., 2007) show the voltage sensor and pore domains as being separate from one another, coupled mechanically via the S4-S5 linker, and structurally by associations between the extracellular region of S1 and residues in the P-loop helix (Lee et al., 2009). Crystallographic modeling of protein conformation has gone a long way towards helping us understand coupling between the voltage sensor and pore domains, and provided ideas of how less structurally-defined components of the channel, particularly the transmembrane linker segments, may be oriented. Yet, it is still often difficult to understand the real-time changes in channel structure that occur during opening and closing (gating) from these structural models. Fluorescence studies, on the other hand, can provide time-resolved measurements of transmembrane domain movement, and these can be interpreted along with ionic and gating current recordings made simultaneously. When a fluorescent probe is linked to a cysteine introduced at the extracellular end of S4, in both Shaker (A359C) and Kv1.5 (A397C) the time course and voltage-dependence of gating charge movement during depolarization matches the fluorescence quenching and fluorescence-voltage relationships (Mannuzzu et al., 1996; Cha and Bezanilla, 1997; Claydon et al., 2007b; Vaid et al., 2008), suggesting that rapid changes in fluorescence reflect conformational changes at the outer end of S4 that underlie Kv activation gating. Fluorophores in S4 in both channels also consistently report secondary changes in fluorescence emission that correlate with events occurring in the pore domain, like inactivation (Loots and Isacoff, 1998; Claydon et al., 2007a; Vaid et al., 2008). Although the transmembrane 30  domains of potassium channels are well conserved among all families in terms of primary and secondary sequence (Doyle et al., 1998; Jiang et al., 2003; Long et al., 2005a), the nature of the secondary fluorescence differs, suggesting that there may be dynamic differences between the pore and/or voltage sensor domain interactions of Kv1 channels arising from differences in channel structure/gating. Voltage clamp fluorimetry has recently been applied to the study of other Kv channels, such as KCa (Savalli et al., 2006), hERG (Smith and Yellen, 2002), Kv1.5 (Vaid et al., 2008) and Kv1.2 (Peters et al., 2009). The high resolution structure of Kv1.2 provides an ideal model to further explore differences in Kv1 gating. In this paper, we have undertaken the first detailed examination of gating current and fluorescence measurements in the Kv1.2 voltage sensor.  Tetramethyl-  rhodamine-5-maleimide (TMRM) labelled at the externally accessible A291C residue in S4 (Figure 2.1) exhibits a fast quenching component that correlates with the time course and voltage-dependence of gating charge movement. In addition, Kv1.2 A291C-TMRM and other residues in the S3-S4 linker, unlike all other voltage-gated channels studied, also detects a second and much slower quenching that represents up to 60% of the total fluorescence change. Through the use of chimera channels and fluorophores placed within S1 and S2, we suggest that this slow phase may report rearrangements of voltage sensor moieties other than S4. The reversal of these slow rearrangements may be required for the channel to deactivate, suggesting that they play an important role in determining the stability of the open channel. Our findings are interpreted in the context of the channel structures in the activated state (Long et al., 2005a; Long et al., 2007), and computer modelling of the closed state (Yarov-Yarovoy et al., 2006; Pathak et al., 2007).  31  Figure 2.1. An alignment of the S3-S4 linkers of Shaker and Kv1.2 highlights similarities in the region. Grey boxes denote the S3 and S4 regions of the protein. Residues assayed for voltage-dependent fluorescence (Figure 2.3) are underlined, and the residue denoted by the filled circle corresponds to Kv1.2 A291C (Shaker A359C).  32  Materials and methods Molecular biology and RNA preparation Shaker and Kv1.2 constructs were expressed in Xenopus laevis oocytes using a modified pBluescript SKII expression vector (pEXO) (a gift from A. Sivaprasadarao, University of Leeds, UK). An N-terminal-deleted, fast-inactivation-removed mutant Δ6-46 (Shaker-IR) (Hoshi et al., 1991), with the lone externally accessible endogenous cysteine removed (C245V), was utilized as the base construct for all subsequent Shaker mutations. Cysteine residues were introduced through site-directed mutagenesis. Oligonucleotide primers were synthesized by either SigmaGenosys (Oakville, ON, Canada) or Integrated DNA Technologies (Coralville, IA), and mutations were generated using the Stratagene QuikChange kit (Stratagene, La Jolla, CA). Successful mutations were confirmed by sequencing the constructs using the core facility unit at the University of British Columbia.  cRNA was synthesized from linear cDNA using the  mMessage mMachine T7 Ultra cRNA transcription kit (Ambion, Austin, TX). Kv1.5-Kv1.2 chimera channels were made as previously described (Rezazadeh et al., 2007). Briefly, PCR amplification of the desired segment of Kv1.2 was used to introduce either a PmlI (Kv1.5-S12LKv1.2) or ClaI (Kv1.5-S23L-Kv1.2) restriction enzyme site. Kv1.5 cDNA was then subcloned into the Kv1.2 channel using the specified enzyme and EcoRI.  Constructs were further  subcloned into a pBluescript SK+ vector using HindIII and EcoRI restriction sites. The A291C mutation was made and sequenced as described above. For construction of the Shaker-Kv1.2 chimera, both channels were first subcloned into a pBluescript SK- vector (EcoRI) in which the XbaI polylinker site had been removed (Klenow). Silent mutations were made in Kv1.2 for the purposes of addition of NsiI and XbaI sites into the equivalent positions to Shaker, flanking the S1-S2 linker. After removal of unwanted occurrences of these restriction sites at other locations in the channels (again through silent mutagenesis), the Shaker S1-S2 linker was subcloned into Kv1.2, and the A291C mutation was made as described above. 33  Oocyte preparation and injection Xenopus laevis frogs were terminally anesthetized by immersion in 2 mg/mL tricaine methanesulphonate (Sigma-Aldrich, Mississauga, ON, Canada); unless otherwise stated, all chemicals were purchased from Sigma-Aldrich.  Stage V-VI oocytes were isolated and  defolliculation was performed through a combination of collagenase treatment, involving mild agitation in 1 mg/mL collagenase Type 1a for approximately 1 hour, and manual defolliculation. Between defolliculation and injection, oocytes were incubated for 1-18 hours in Barth’s solution, which contained (in mM), 88 NaCl, 1 KCl, 2.4 NaHCO3, 0.82 MgSO4, 0.33 Ca(NO3)2, 0.41 CaCl2, 20 HEPES, titrated to pH 7.4 using NaOH. Injection of 50 nL of cRNA encoding the construct of interest was performed using a Drummond digital microdispensor (Fisher Scientific, Ottawa, ON, Canada), followed by incubation at 19°C in Barth’s solution. Currents were recorded 1-4 days after injection. All animal protocols were performed in accordance with University of British Columbia animal care guidelines.  Two-electrode voltage clamp Oocytes were placed in a bath chamber that was perfused with control ND96 bath solution containing (in mM), 96 NaCl, 3 KCl, 1 MgCl2, 2 CaCl2, and 5 HEPES, titrated to pH 7.4 with NaOH. Microelectrodes were filled with 3 M KCl and had resistances of 0.2 to 2.0 MΩ. Unless otherwise stated, data were recorded from a holding potential of -80 mV. Voltage control and data acquisition were achieved with a Warner Instruments OC-725C amplifier (Hamden, CT), and Axon Digidata 1322 A/D converter (Axon Instruments, Foster City, CA), connected to a personal computer running pClamp9 software (Molecular Devices Corp.).  34  Voltage clamp fluorimetry Fluorimetry was performed simultaneously with two-electrode voltage clamp. Labelling of the oocytes with 5 µM tetramethylrhodamine-5-maleimide (TMRM; Invitrogen, Carlsbad, CA) dye was performed for 30 minutes at 10°C in a depolarizing solution containing (in mM), 98 KCl, 1 MgCl2, 2 CaCl2, and 5 HEPES, titrated to pH 7.4 using KOH. Signals were recorded via a Nikon TE300 inverted microscope with Epi-Fluorescence attachment and a 9124b Electron Tubes photomultiplier tube (PMT) module (Cairn Research, Kent, UK). TMRM dye, exhibiting maximal light absorption at 542 nm, was excited by light from a 100 W mercury lamp filtered through a 525 nm band pass filter, reflected by a 560 nm dichroic mirror through a 20x objective lens focussed on the oocyte. Emitted light, with a maximal emission at 567 nm, was collected by the objective lens and passed through the dichroic mirror to the PMT module. Voltage signals from the PMT were digitized using an Axon Digidata 1322 A/D converter and passed to a computer running pClamp9 software to record fluorescence intensity. To minimize fluorophore bleaching, a Uniblitz computer-controlled shutter (Vincent Associates, Ottawa, ON, Canada) was used, and opened shortly prior to application of voltage-clamp pulses. For 100 ms pulses, fluorescence signal sampling frequency was 20 kHz; traces were averaged, with each signal representing the average of five sweeps, and were filtered offline at 1000 Hz. To account for any photobleaching of fluorophores that may occur during shutter opening, control fluorescence data were recorded in the absence of any change in voltage, and subtracted from the voltagedependent signal. Fluorescence records were normalized to baseline emission in the absence of a change in voltage, to control for cell-to-cell variability, and fluorescence-voltage (F-V) relationships are shown normalized to maximum and minimum levels of emission over the voltage ranges tested.  35  Gating currents Mammalian tsA201 cells were grown and maintained in Minimal Essential Medium (MEM) at 37°C in an air/5% CO2 incubator. Media contained 10% bovine serum and 0.5 mg/ml geneticin.  On the day before transfection, cells were washed with MEM, treated with  trypsin/EGTA for one minute and plated on 25 mm2 glass coverslips. WT Kv1.2-pGW1 and GFP-pcDNA3 cDNA were then transiently co-transfected using Lipofectamine 2000 (Invitrogen, Mississauga, ON, Canada). Twenty-four to forty-eight hours after transfection, coverslips with adherent cells were placed in a superfusion chamber (volume 300 µl) containing the control bath solution at room temperature (approximately 25°C). Whole-cell current recording and analysis were carried out using an Axopatch 200A amplifier and pClamp10 software (Axon Instruments). Patch electrodes were pulled from thin-walled borosilicate glass (World Precision Instruments, Sarasota, FL) on a horizontal micropipette puller (Sutter Instruments, Novato, CA). Electrodes had resistances of 1.0-1.5 MΩ when filled with control filling solution. Membrane potentials were not corrected for junction potentials that arise between the pipette and bath solution. Extracellular bath solution contained (in mM), TEA (140), HEPES (10), Dextrose (10) and MgCl2 (1), pH adjusted to 7.4 (HCl). Intracellular pipette solution contained (in mM), NMDG (140), HEPES (10), MgCl2 (1) and EGTA (10), pH adjusted to 7.2 (HCl).  Cells were  maintained at a holding potential of -100 mV, and currents were recorded during 12-ms voltageclamp pulses from -80 mV to +80 mV in 10-mV increments. Leak subtraction was performed using a -P/6 protocol from holding potential. Data were filtered at 10 kHz and sampled at 100 kHz.  Data analysis Conductance-voltage (G-V) relationships were derived using normalized chord conductance, which was calculated by dividing peak current by the driving force based on the 36  potassium equilibrium potential (internal potassium concentration was assumed to be 99 mM). F-V relationships were calculated based on the peak change in emission of the total trace or, in some stated cases, the individual components of the signal. All G-V, gating charge-voltage (QV), and most fluorescence-voltage (F-V) relationships (see below) were fit with a single Boltzmann function,  y=  1 ⎡V − V ⎤ 1 + exp ⎢ 1 / 2 ⎥ ⎣ k ⎦  where y is the conductance normalized with respect to the maximal conductance, V1/2 is the halfactivation potential, V is the test voltage, and k is the slope factor. For analysis of the slow phase of Kv1.2 fluorescence and S289C fluorescence, data were best fit with a double Boltzmann function,  y=  A1  ⎡ (V ) − V ⎤ 1 + exp ⎢ 1 / 2 1 ⎥ k1 ⎣ ⎦  +  A2  ⎡ (V ) − V ⎤ 1 + exp ⎢ 1 / 2 2 ⎥ k2 ⎣ ⎦  where symbols are as described above, A refers to the amplitude of the fit component, and 1 and 2 refer to the separate components of the fit. Unless otherwise indicated, data throughout the text and figures are reported as mean ± SEM.  Supplemental material Accompanying supplemental figures and text are included in Appendix A. Figure A.1 explores the effect of TMRM on WT Kv1.2 activation and conductance. Figure A.2 compares TMRM fluorescence at A291C in the presence and absence of the removal of the lone potentially  37  reactive endogenous cysteine residue (C181). Figure A.3 shows representative fluorescence traces obtained from a Kv1.5-Kv1.2 chimera channel in which the voltage sensor domain of Kv1.2 was replaced up to and including the S2-S3 linker segment. Figure A.4 shows a structural diagram of the voltage sensor domain, indicating residues in the S1, S2 and S4 regions labelled with TMRM in this study.  38  Results  Kv1.2 WT channels do not show voltage-dependent changes in fluorescence emission Removal of the lone potentially-labellable cysteine residue (C181) in the outer S1 greatly reduced channel expression levels, possibly due to a role of this residue in stabilizing interactions between the voltage sensing and pore domains (Lee et al., 2009). Supplemental Figure A.1 shows no effects of TMRM on WT Kv1.2 gating, and no voltage-dependent changes in emission. Furthermore, fluorescence from a probe attached to A291C shows identical voltage-dependent characteristics with and without the WT cysteine at position C181 (Supplemental Figure A.2). Based on these results, all studies presented here were carried out in the background of Kv1.2 C181 (WT).  Kv1.2 fluorescence at A291C exhibits differences from the Shaker A359C homologue As shown previously (Claydon et al., 2007a), along with activation of ionic currents, TMRM fluorescence from Shaker C245V A359C shows rapid fluorescence quenching during short depolarizations (Figure 2.2A,B). During large depolarizations a rapid quenching accounts for 80-90% of the total fluorescence change and correlates with S4 translocation (Mannuzzu et al., 1996; Cha and Bezanilla, 1997; Cha and Bezanilla, 1998).  At potentials negative to  activation of ionic current (e.g. -60 mV), fluorescence quenching was still observed, suggesting that some S4 movement precedes pore opening; this is also shown in the comparison of peak GV and F-V relationships (Figure 2.2C). In Shaker C245V A359C, the V1/2 of the F-V relationship is left-shifted by ~27 mV compared to the G-V. Kv1.2 A291C ionic current (Figure 2.2D) had similar voltage-dependent and kinetic characteristics to both Shaker C245V A359C (Figure 2.2A) and Kv1.2 WT (Figure A.1) channels. In contrast, the fluorophore emissions (Figure 2.2B, 2.2E) were strikingly different. Depolarization produced an initial fast fluorescence decay similar to that observed in Shaker, that 39  was complete within the first 2-5 ms of depolarization. This was followed by a large secondary slow component that comprised the majority of the total signal and continued for the duration of the depolarization (grey line in Figure 2.2E). Fluorescence quenching was observed at potentials quite negative to channel opening, more so than in Shaker (compare Figures 2.2B and 2.2E between -80 mV and -120 mV). Consequently the total F-V relationship was found to be ~51 mV left-shifted compared to the G-V (Figure 2.2F), much larger than the left shift in Shaker A359C channels (Figure 2.2C).  40  Figure 2.2. Voltage-dependent Shaker and Kv1.2 conductance and fluorescence deflections. Typical current traces (A and D) and fluorescence signals (B and E) are shown for Shaker C245V A359C and Kv1.2 C181 A291C for 100 ms pulses between -120 mV and +60 mV, from -80 mV. The dashed line in panel (E) marks the approximate division of the two observed components of A291C fluorescence for depolarizations positive to 0 mV. (C and F) Normalized conductance-voltage (G-V, filled symbols) and fluorescence-voltage (F-V, hollow symbols) relationships were calculated from data obtained at the end of each 100 ms pulse (mean ± SEM; n = 7-8), and data were well fit with Boltzmann equations. The V1/2 values for Shaker C245V A359C were -12.9 ± 1.6 mV and -39.5 ± 1.5 mV for the G-V and F-V, and slope factors (k) were 17.8 ± 1.1 mV and 22.3 ± 1.4 mV respectively. The Kv1.2 A291C G-V had a V1/2 and slope factor of -11.2 ± 1.6 mV and 22.4 ± 0.9 mV respectively, while the F-V relationship had a V1/2 and k of -62.9 ± 1.2 mV and 23.2 ± 1.0 mV.  41  The slow Kv1.2 fluorescence is recapitulated at other labelled sites in the S3-S4 linker To investigate whether the differences between Kv1.2 A291C and Shaker A359C fluorescence were specific to this position, or indicative of a more general difference between the two channels, we performed a scan of seven residues within the S3-S4 linker and S4 in each channel. Three residues N-terminal to A291C (or A359C) and three C-terminal residues (Figure 2.1) were tested for voltage-dependent deflections, and typical traces for five of these are shown for Shaker (Figure 2.3A) and Kv1.2 (Figure 2.3B) for 100 ms pulses to +60 mV. Previous studies have extensively scanned the externally accessible residues of Shaker (Gandhi et al., 2000; Pathak et al., 2007), though these studies have been more concerned with changes occurring on the order of seconds rather than milliseconds. As a result, it is difficult to draw direct comparisons between these data and previous work, other than to say the directionality of the changes in fluorescence (quenching vs. dequenching) appear to be the same. We were able to obtain deflections for all 7 residues in Shaker and 6 of 7 residues in Kv1.2, with the R294C construct not expressing any discernible current or fluorescence in over 40 recordings. In both Shaker C245V L361C and Kv1.2 L293C, we were only able to obtain fluorescence deflections upon hyperpolarization (data not shown). Residues in the Shaker S3-S4 linker showed a predominantly rapid quenching phase, with a negligible or very small slow quenching component. In Kv1.2, M288C and S289C as well as A291C exhibited a distinct slow quenching phase in addition to the initial fast component. Conductance- and fluorescence-voltage relationships of the first four residues of the scan for both Shaker and Kv1.2 are shown in Figure 2.3C-F. All data are well fit with a single Boltzmann function (Table I), with the exception of Kv1.2 S289C fluorescence, which can be reasonably fit with a double Boltzmann function. The first component of this relationship, scaled to its maximum value at -20 mV, is shown in grey in Figure 2.3D. Overall, the data show that the negative shift in the F-V relationship between Kv1.2 and Shaker observed with 42  A291C/A359C (Figure 2.3F) is not confined to this residue, as S289C and L290C (Figure 2.3CE) also show hyperpolarized F-V relations compared with Shaker equivalent residues. The data suggest that global rearrangements of the Kv1.2 voltage sensor domain occur at more negative potentials relative to observed gating movements in the Shaker channel.  43  Figure 2.3. A scan of the Shaker and Kv1.2 S3-S4 linkers reveals differences in fluorescence phenotypes. (A and B) Representative fluorescence traces collected in a cysteine scan of 5 consecutive homologous residues in the S3-S4 linker and NH2-terminal end of S4 in Shaker (A) and Kv1.2 (B). Cells expressing these constructs were held at -80 mV and depolarized to +60 mV for 100 ms. (C to F) Normalized G-V (filled symbols) and F-V (hollow symbols) relationships for four of the five Shaker (squares) and Kv1.2 (circles) constructs expressing changes in fluorescence upon depolarization, as labelled in the top left corner of each panel. For Kv1.2 S289C (Panel D), the grey circles and accompanying fit denote the Boltzmann fit to the voltage-dependent fluorescence observed between -140 mV and -20 mV. Mean halfactivation and slope data obtained from the fits to all four mutant constructs for each channel can be found in Table I. Data are shown as mean ± SEM, and are the average of 3-8 cells collected from each mutant.  44  Table 2.1. Boltzmann fits to Kv1.2 and Shaker G-V and F-V relationships from Figure 2.3.  Kv1.2  Shaker G-V  Construct M288C S289C (1) S289C (2) L290C A291C  V1/2 (mV) -12.4 ± 1.1 9.8 ± 3.3 n/a -39.3 ± 4.7 -12.6 ± 1.9  F-V k (mV) 23.0 ± 0.4 18.1 ± 0.7 n/a 26.5 ± 4.7 26.1 ± 1.4  V1/2 (mV) -56.9 ± 3.5 -105.0 ± 2.8 41.7 ± 9.2 -91.9 ± 2.4 -62.5 ± 1.3  G-V k (mV) 26.2 ± 1.6 -9.3 ± 2.6 -33.3 ± 2.0 19.7 ± 1.7 23.3 ± 1.0  Construct M356C S357C L358C A359C  V1/2 (mV) -0.5 ± 1.5 5.6 ± 2.3 n/a -26.2 ± 4.2 -14.6 ± 1.7  F-V k (mV) 17.3 ± 0.7 18.6 ± 1.1 n/a 18.2 ± 1.8 19.8 ± 1.4  V1/2 (mV) -41.8 ± 2.6 -42.9 ± 4.4 n/a -59.5 ± 0.5 -38.8 ± 1.9  k (mV) 20.0 ± 1.7 9.7 ± 1.2 n/a 21.6 ± 1.7 22.6 ± 1.4  Mean half-activation (V1/2) and slope (k) data were calculated from single Boltzmann fits to the data, with the exception of the F-V relationship for S289C, which was best fit with a double Boltzmann function.  45  Increases in Kv1.2 fluorescence upon hyperpolarization reflect mobility of the voltage sensor As seen in Figure 2.2, there were large differences between Shaker and Kv1.2 in the extent of fluorescence change when channels were pulsed to potentials more negative than -80 mV. Depolarizations from -120 mV (Figure 2.4A) still showed the same biphasic fluorescence phenotype as that seen from -80 mV, though hyperpolarizations from this voltage no longer resulted in increased fluorescence emission. When cells were held at -50 mV (Figure 2.4B), the fluorescence increase was much greater upon hyperpolarization, and fluorescence observed during depolarization was smaller and composed almost exclusively of a fast quenching component. Ionic currents (data not shown) and G-V relationships (Figure 2.4C, filled symbols) were not any different between the three holding potentials. Furthermore, peak F-V relationships were unchanged, when total increases in fluorescence emission at hyperpolarized potentials were taken into account (hollow circles in Figure 2.4C). Peak F-V data from a holding potential of -120 mV (squares) and -50 mV (circles) had half-activation voltages of -69.5 ± 2.0 mV and -69.9 ± 5.8 mV respectively, compared to -69.1 ± 4.5 mV at -80 mV. These data suggest that the fluorescence unquenching at hyperpolarized potentials reflects the holding potential-dependent position of the voltage sensor domain relative to its environment, and that mobility of this region reaches one of its extreme positions at or close to -120 mV. The fact that Kv1.2 F-V relationships are similar from holding potentials between -50 mV and -120 mV also suggests that only readily reversible gating changes occur over this potential range. While it has been shown that Kv1.2 channels do not undergo significant inactivation of ionic current (Paulmichl et al., 1991; Russell et al., 1994) it is well-known that in Shaker and other Kv1 channels prolonged depolarizations associated with inactivation immobilize off-gating charge and shift the Q-V relationship in the hyperpolarized direction due to stabilization of the  46  S4 segment in an activated conformation, and delay its return upon hyperpolarization (Fedida et al., 1996; Chen and Fedida, 1997; Olcese et al., 1997). In Shaker, holding the membrane potential at -50 mV resulted in a ~40 mV hyperpolarizing shift of the A359C F-V relationship compared with those from -80 and -120 mV (Figure 2.4D, hollow squares), despite minimal differences between the corresponding G-V relationships. Given that fluorophore labelling at this Shaker residue has been shown to track changes in S4 environment during activation, these data suggest impaired movement of the Shaker S4 helix at -50 mV, compared to -80 mV and -120 mV, consistent with holding potential-dependent modulation of S4 gating.  47  Figure 2.4. Holding potential affects the directionality of Kv1.2 fluorescence deflections, but does not affect the overall F-V relationship. Representative deflections of Kv1.2 C181 A291C held at -120 mV (A) and -50 mV (B), in response to 100 ms changes in voltage between -150 mV and +60 mV. (C) Normalized G-V (filled symbols) and F-V (hollow symbols) relationships for Kv1.2 C181 A291C from three different holding potentials (mean ± SEM; n = 3-5), with F-V relationships adjusted to account for upward deflections. From -120 mV (triangles), the G-V had a V1/2 of -10.7 ± 2.5 mV and a slope factor of 24.8 ± 0.9 mV, and corresponding values of -69.5 ± 2.0 mV and 22.7 ± 1.1 mV respectively for the F-V. V1/2 and slope factor data from -80 mV (circles) were -2.2 ± 2.3 mV and 24.6 ± 0.7 mV for the G-V and -69.1 ± 4.5 mV and 20.8 ± 0.8 mV for the F-V. At -50 mV (squares), the V1/2 and slope factor for the G-V and F-V were 2.6 ± 0.8 mV and 24.2 ± 1.1 mV, and -69.9 ± 5.8 mV and 22.6 ± 1.5 mV respectively. (D) Shaker A359C G-V and F-V relationships from the same three holding potentials as in (C) (mean ± SEM; n = 7-8). From -120 mV (triangles), the G-V had a V1/2 of -22.2 ± 2.1 mV and a slope factor of 15.8 ± 1.6 mV, and corresponding values of -35.0 ± 4.5 mV and 27.5 ± 1.6 mV, respectively, for the F-V. V1/2 and slope factor data from -80 mV (circles) were -22.1 ± 1.9 mV and 15.9 ± 1.7 mV for the G-V and -39.8 ± 4.0 mV and 29.1 ± 2.1 mV for the F-V. At -50 mV (squares), the V1/2 and slope factor for the G-V and F-V were -12.7 ± 1.9 mV and 15.6 ± 1.7 mV, and -79.6 ± 4.8 mV and 32.4 ± 1.5 mV, respectively.  48  The fast and slow fluorescence phases in Kv1.2 have different voltage-dependencies We have shown that upon depolarization, fluorescence from the S3-S4 linker of Kv1.2 A291C shows two distinct phases of quenching: a fast quenching component, followed by a slower secondary quenching component that continues for the duration of the depolarization. Figure 2.5A shows a typical fluorescence trace during a depolarization to +60 mV, that was best fit with a bi-exponential function (grey dotted line) at this and all other potentials tested. Approximately 40% of the total deflection comprised a fast quenching component with a time constant of ~1.3 ms, while the remaining 60% of quenching occurred ~20 times slower, with a time constant of 23.9 ms in this particular example. The time constants of the rapid quenching component (τF, fast) decreased with depolarization (3.87 ± 0.6 ms at -40 mV, compared to 1.2 ± 0.3 ms at +80 mV) and were faster than the ionic current activation time constants at all potentials (2.47 ± 0.9 ms at +80 mV) while the slow quenching component (τF, slow) was clearly slower than activation at depolarized potentials, and showed little voltage dependence (Figure 2.5B). These data suggest that the fast fluorescence may report on a process required for activation, while the event that the slow fluorescence change reflects is not required per se for channel opening. As a function of potential it is clear that each component of the total fluorescence signal is distinct (Figure 2.5C). The slow phase of fluorescence (hollow squares) was fitted with a double Boltzmann function with widely separated V½s of -74 mV and +44 mV. Most (89%) of the slow fluorescence corresponded to a change in voltage sensor environment that occurred at more negative potentials, based on the hyperpolarized position of the F-V compared to either the fast quenching movement (open circles) or ionic conductance (filled circles).  The second  component of this slow fluorescence quenching, present only at very positive potentials,  49  constitutes a relatively minor component (~10%) of the total slow fluorescence signal, similar to that observed presently (Figure 2.2B) and previously in Shaker A359C (Claydon et al., 2007a). The separation of the fast and slow fluorescence components is well illustrated by comparing fluorescence records from different holding potentials (Figure 2.5D). The fast phase of fluorescence differs only slightly (~20%) during a pulse to +60 mV from either -80 or -50 mV, whereas the slow component is reduced by more than two-thirds. A similar result can be seen during a depolarization to -30 mV from the same two potentials. These data clearly support the idea that the fast and slow components of fluorescence can be modulated separately from one another, and thus likely represent different conformational changes within the channel.  50  Figure 2.5. Kv1.2 A291C voltage-dependent fluorescence is well characterized by a double exponential function. (A) Fit of the Kv1.2 A291C fluorescence signal at +60 mV to a double exponential shows that approximately 40% of the signal amplitude results from a fast movement, with a time constant of 1.3 ms. The slow phase, comprising 60% of the total signal, is slower by an order of magnitude, 23.9 ms in this example. (B) Mean ± SEM time constants of the fast and slow fluorescence signal components (n = 14-20), compared to time constants of ionic current activation, fit from ~50% of maximal activation. (C) Normalized F-V relationships of the fast and slow components of Kv1.2 A291C fluorescence, normalized and plotted alongside the G-V relationship (n = 11). The normalized fast phase, fit to a Boltzmann distribution, had a V1/2 and slope factor of -39.5 ± 2.0 mV and 15.6 ± 1.0 mV. The voltage-dependence of the slow phase was best fit with a double Boltzmann function, with the first component having a V1/2 and k of -73.9 ± 1.4 mV and 12.0 ± 0.5 mV (amplitude = 88.7 ± 2.3 %), followed by a second component (11.2 ± 2.5 %) with respective V1/2 and slope factors of 44.3 ± 4.2 mV and 11.6 ± 3.0 mV. (D) Holding potential-dependent separation of the fast and slow fluorescence components. Representative fluorescence traces are shown for Kv1.2 C181 A291C channels depolarized to -30 mV (grey traces) or +60 mV (black traces), from holding potentials (HP) of either -80 mV or -50 mV as labelled. The two vertical lines to the left of the fluorescence records show the contributions of fast and slow quenching components for depolarizations to +60 mV.  51  The fast phase of fluorescence correlates with S4 movement and the Q-V relationship In Shaker A359C, rapid fluorescence quenching upon depolarization has been correlated with the translocation of S4 gating charge, based on the comparison of F-V and Q-V relationships (Mannuzzu et al., 1996; Cha and Bezanilla, 1997; Cha and Bezanilla, 1998). Given the speed and voltage dependence of the fast phase of Kv1.2 A291C fluorescence, it seemed appropriate to measure the charge-voltage (Q-V) relationship of Kv1.2. However, simultaneous measurement of gating currents with voltage clamp fluorimetry was not possible in the oocyte due to the clamp speed limitations of two-electrode voltage clamp. Therefore, we recorded Kv1.2 gating currents from tsA201 cells using whole-cell patch clamp, as described in Methods. Figure 2.6A shows representative on-gating currents during 12 ms pulses from a holding potential of -80 mV up to +60 mV. To our knowledge these are the first reported measurements of Kv1.2 gating currents and they look much the same as other Kv1 gating current recordings from mammalian cells (Hesketh and Fedida, 1999).  The Q-V relationship during depolarization (filled triangles)  matched the voltage dependence of the fast fluorescence (Figure 2.6B) and both relationships were well fit with a Boltzmann function, with half-activation voltages of -31.5 ± 2.0 mV and -39.5 ± 2.0 mV for the Q-V and fast F-V relationships, respectively. The time course of gating charge movement and fast fluorescence quenching are shown for a range of potentials in Figure 2.6C, and are reasonably well-matched, given the limited time resolution of the oocyte clamp. When the time course of the integrated gating charge movement was fit with a double exponential function, the slow component correlated well with the fast change in fluorescence emission (Figure 2.6D), as has been reported in Shaker channels (Cha and Bezanilla, 1997). These data support the idea that a fluorophore attached at the external end of S4 in Kv1.2 reports  52  fast changes in fluorescence emission that correlate with the movement of gating charge, and thus S4 movement and channel activation.  Figure 2.6. Correlation of the fast component of Kv1.2 fluorescence quenching with gating charge movement. (A) Representative gating currents for Kv1.2 WT, recorded from transiently transfected tsA201 cells. Data were recorded from 12 ms pulses from -80 mV to +60 mV in 10 mV increments; only every third voltage is shown here for clarity. (B) Overlay of the mean normalized charge-voltage (Q-V) relationship with the fast F-V relationship from Figure 2.5C. The Q-V relationship V1/2 and k values were -31.5 ± 2.0 mV and 11.5 ± 0.6 mV, respectively. (C) Superposition of fluorescence (grey) and cumulative gating charge (black) versus time in Kv1.2 channels. Data are shown for a range of depolarizations from -50 mV to +60 mV, as labelled, for the initial 8 ms of depolarization in the case of the fluorescence data, and are normalized to the respective maximum values. The fluorescence quenching at this potential is inverted in order to more closely compare with the gating charge data. (D) Mean time constants of the integrated gating charge movement compared to the fast fluorescence quenching of A291C. Gating charge data were fit to a double exponential function, and the mean ± SEM data (n = 10-15) of the fast (filled squares) and slow (hollow squares) components were plotted as a function of voltage. Time constants for the fast quenching event (hollow circles) are as plotted in Figure 2.5B.  53  Slow changes in fluorescence recovery match the rates of Kv1.2 channel deactivation and reactivation The major component of slow fluorescence quenching had a voltage dependence that was hyperpolarized from both gating charge movement and pore opening (Figure 2.5C). Given the voltage-dependent and kinetic properties of the fluorescence change, it is unlikely that the slow phase can be associated with pore opening. We did find, though, that the time course of slow fluorescence recovery after repolarization was strongly correlated with both channel deactivation and also re-availability after progressively longer interpulse intervals, as illustrated in Figure 2.7. After depolarizations to a range of potentials, fluorescence increased back to baseline with an identical time course to the deactivation of ionic current at -120 mV (Figure 2.7A) or a range of potentials (Figure 2.7B, 2.7C). For example, deactivation of ionic current to -120 mV occurred with a mean time constant of 7.6 ± 1.5 ms, similar to that observed in previous studies (Watanabe et al., 2007; Lewis et al., 2008), compared to a time constant of 9.3 ± 0.5 ms for the change of fluorescence emission. The reactivation of Kv1.2 channels during a two-pulse protocol is shown in Figure 2.7D. After short P1-P2 intervals (e.g. 6.25 ms) at -80 mV the slow fluorescence quenching during the P2 pulse to +60 mV was quite small, as expected given that little recovery of the tail had occurred in the interpulse interval. This was accompanied by faster activating ionic currents and greater instantaneous current after a 6.25 ms or 25 ms P1-P2 interval. A scaled and inverted fit of the slow off-fluorescence (grey dashed line) matched the recovery of the initial ionic current back to P1 values (grey arrow). Figure 2.7E shows mean data for the normalized instantaneous current in the P2 pulse, compared with the slow fluorescence recovery after the first P1 pulse. The current reactivation (τionic = 15.7 ± 1.3 ms) correlated extremely well with the time dependence of fluorescence recovery (τFl = 18.5 ± 0.6 ms), suggesting that the slow fluorescence  54  may track a conformational change in the protein that, upon repolarization, is rate-limiting for deactivation of ionic current.  Figure 2.7. Slow fluorescence return upon hyperpolarization correlates with deactivation of ionic current. (A) Kv1.2 A291C currents (top) and fluorescence (bottom) traces at -120 mV, -60 mV, 0 mV and +60 mV, from a holding potential of -120 mV. The right panels show an enlarged view of the tail currents and off-fluorescence emissions. (B) Overlay of ionic tail current (black lines) and slow off-fluorescence quenching (grey) for Kv1.2 A291C at the three potentials labelled, after a depolarization to +20 mV. Scale bars are as shown in the legend, left to right, for the corresponding holding potentials. (C) Deactivation and slow off-fluorescence time constants for Kv1.2 A291C at the three holding potentials shown in (B), as a function of prepulse potential (n = 3-5). Mean data are shown every 20 mV for clarity. (D) Representative current and fluorescence records from a dual pulse (P1-P2) protocol with varying interpulse recovery time. Data are shown for 100 ms pulses from -80 mV to +60 mV with interpulse intervals of 6.25, 25, 100 and 250 ms. For clarity, data are only shown up to the end of the second depolarizing pulse of intermediate records. The grey arrow shows the instantaneous level of ionic current at P1; the grey dashed line is an inverted fit of the slow component of offfluorescence from P1. (E) Overlay of the slow off-fluorescence component in (D) with the normalized initial P2 current amplitudes (black diamonds). The black line is a single exponential fit to the ionic current data. Mean time constants for the fits to individual data sets and the offfluorescence component are shown in the panel (n = 5).  55  Slow changes in fluorescence emission reflect internal rearrangement within the voltage sensor domain To investigate whether internal rearrangement of the voltage sensor could explain the slow quenching, we replaced portions of the voltage sensing domain of Kv1.2 with those from another mammalian Shaker homologue, Kv1.5.  The fluorescence of Kv1.5 has been  characterized in previous studies (Vaid et al., 2008; Vaid et al., 2009) and, importantly, does not have a Kv1.2-like slow quenching component in its voltage-dependent fluorescence. We made two chimeras in which Kv1.2 was replaced with Kv1.5 sequence up to the S1-S2 linker (Kv1.5S12L-Kv1.2) and up to the S2-S3 linker (Kv1.5-S23L-Kv1.2), in the background of A291C (Figure 2.8A). Typical fluorescence records for Kv1.5-S12L-Kv1.2 are shown in Figure 2.8B. Ionic currents during depolarization were unchanged in the chimeric channels (data not shown), but there were marked differences in quenching observed compared to Kv1.2 A291C (Figure 2.2E). Replacing the first transmembrane segment, S1, and the extracellular S1-S2 linker with those from Kv1.5 dramatically reduced the slow quenching associated with Kv1.2 A291C fluorescence (Figure 2.8C). The more extensive replacement of the Kv1.2 voltage sensor with the S2 segment and intracellular S2-S3 linker did not have any further effects on the fluorescence (Supplemental Figure A.3). The fast fluorescence quenching from these channels comprised ~80% of the total amplitude (similar to Shaker A359C over the same time course), compared to only 40% in Kv1.2 A291C. Fluorescence recorded from both chimera channels looked very similar to fluorescence recorded from Kv1.2 A291C from a holding potential of -50 mV (Figure 2.4B), a potential at which the majority of slow fluorescence change had already occurred (Figure 2.5C). A parallel loss of the slow fluorescence recovery on repolarization was observed (Figure 2.8B,C see also Supplemental Figure A.3), and, importantly, the S12L chimera showed  56  accelerated ionic current deactivation that matched the time course of fluorescence decay (Figure 2.8C, inset), further suggesting that these two processes are linked through some common or related molecular rearrangement. The lack of a slow component of fluorescence at hyperpolarized potentials in the two Kv1.5-Kv1.2 constructs resulted in a depolarizing shift of the F-V relationships (Figure 2.8D), with no change in the G-V. The fast fluorescence quenching, with time constants on the order of milliseconds, showed an identical voltage-dependence to the fast component of Kv1.2 A291C (Figure 2.8E, circles) and the Kv1.2 Q-V relationship (Figure 2.8F), suggesting that the majority of the remaining fluorophore quenching in the chimera reports conformational changes associated with S4 translocation. The residual slow component of fluorescence in the chimera evident at more depolarized potentials, had a V1/2 of +33.4 ± 2.5 mV (data not shown), very similar to the small slow component of WT Kv1.2 fluorescence.  57  Figure 2.8. Kv1.2-Kv1.5 chimera channels lack the slow fluorescence quenching at negative potentials. (A) Cartoon representation of Kv1.2 A291C (top), Kv1.5 WT (bottom), and the two chimeric channels (middle cartoons). Hollow squares (and their connecting segments) originate from Kv1.2, and slashed squares are Kv1.5 segments. A291C is located approximately with a filled circle at the N-terminal S4. (B) Representative fluorescence traces from the Kv1.5-S12L-Kv1.2 A291C chimera. Data were collected using the protocol outlined in Figure 2.2. (C) Overlay of representative fluorescence deflections for Kv1.2 A291C and Kv1.5S12L-Kv1.2 (S12L) A291C, for depolarizations to +30 mV, normalized to the fast fluorescence quenching components. The inset shows overlays of deactivating ionic tail currents and offfluorescence of Kv1.5-S1S2L-Kv1.2 A291C (above) and Kv1.2 A291C (below), with scale bars as noted. (D) Mean normalized G-V (filled symbols) and F-V (hollow symbols) relationships for the Kv1.5/Kv1.2 chimera channels, compared to Kv1.2 A291C, shown ± SEM (n = 10-13). Boltzmann fits to the data from Kv1.5-S12L-Kv1.2 A291C (S12L) gave V1/2 and k values of 2.6 ± 2.6 mV and 23.9 ± 0.9 mV and -40.3 ± 3.1 mV and 28.4 ± 2.6 mV for the G-V and F-V respectively; the fits to Kv1.5-S23L-Kv1.2 A291C (S23L) were 7.3 ± 1.9 mV and 23.9 ± 0.7 mV for the G-V relationships and -40.9 ± 2.2 mV and 23.4 ± 2.3 mV for the F-V relationships. Kv1.2 A291C values are as reported in Figure 2.4. (E) Voltage-dependence plots of the fast fluorescence component of Kv1.5-S23L-Kv1.2 A291C with that of Kv1.2 A291C from Figure 2.5C. V1/2 and k values were -40.3 ± 2.2 mV and 19.5 ± 1.7 mV for the chimera channel (n = 1517). (F) Mean normalized F-V relationship of the fast component of Kv1.5-S23L-Kv1.2 A291C emission, overlaid with the Kv1.2 Q-V relationship (filled circles) from Figure 2.6B.  58  A third chimera channel, in which only the S1-S2 linker of Kv1.2 was replaced with the equivalent segment from Shaker (Figure 2.9A), resulted in the complete abolition of slow fluorescence from A291C, similar to the Kv1.5-Kv1.2 chimeric channels, (Figure 2.9B), while ionic currents were unchanged (data not shown).  The loss of fluorescence is particularly  apparent at negative potentials between -120 mV and -50 mV where the slow phase of Kv1.2 was most prominent, and highlights the importance of the linker region in detecting slow reorganizations of the voltage sensor.  Figure 2.9. Effect of Shaker S1-S2 linker replacement on Kv1.2 fluorescence. (A) Representative fluorescence records from a Kv1.2-Shaker S1-S2 linker A291C chimera channel (Kv1.2Sh12L). Traces are shown for depolarizations to potentials as labelled, utilizing the same protocol as outlined in Figure 2.2. (B) Mean F-V relationship for Kv1.2Sh12L (triangles), compared to Kv1.2 A291C (circles) and Shaker A359C (squares) as previously shown in Figure 2.2. For Kv1.2Sh12L, V1/2 and k values for the Boltzmann fit were -56.0 ± 2.5 mV and 15.5 ± 0.8 mV (n = 7).  The S1-S2 linker and S4 region are both intricately involved in Kv1.2 activation The ability of a TMRM fluorophore, attached to residue C291, to report on movements of both S4 and the S1-S2 region suggested to us that these two regions are sufficiently close in one or both of the closed and open states to alter S4 fluorophore environment in a voltage-dependent manner. To test whether S1-S2 movements alone were responsible for slow changes in Kv1.2 fluorescence, we recorded voltage-dependent fluorescence directly from cysteines inserted in the S1-S2 linker (boxed residues in Figure 2.10A), which based on the crystal structure are 59  somewhat more distant from S4 and the pore region, compared with A291C in the S3-S4 linker (Figure 2.11). Test residues were located near the outer end of S1 (I187) or S2 (F218, T219, D220), as well as in the middle of the linker, positioned toward adjacent subunits in the crystal structure (S208, T209) (Long et al., 2005b).  Of these, only two constructs gave voltage-  dependent deflections; I187C (Figure 2.10B) and T219C (Figure 2.10C), shown in bold in the alignment, and both showed only slow quenching. Labelled I187C channels showed slow voltage-dependent emission quenching with time constants similar to the slow fluorescence emission from Kv1.2 A291C (Figure 2.5), and lacked any evidence of fast fluorescence quenching. The S2 fluorophore T219C also showed only slow decreases in fluorescence emission during depolarizations, of a similar time course; this is the homologous residue to T276C in Shaker, which has previously been shown to exhibit voltagedependent fluorescence emission left-shifted relative to the Q-V (Cha and Bezanilla, 1997). In addition, the total F-V relationship for T219C fluorescence (hollow triangles) correlated well with the voltage-dependence of the slow fluorescence component of A291C, and had a similar half-activation potential (-70.8 ± 5.9 mV, Figure 2.10D). I187C did detect movement between -140 and -90 mV, clearly apparent in Figure 2.10D, but most change of fluorescence occurred at more positive potentials.  60  Figure 2.10. TMRM attached to residues in the S1-S2 linker of Kv1.2 report only slow changes in fluorescence emission in response to voltage. (A) An alignment of best fit for the amino acid residues of the S1-S2 linker region (underlined) of Shaker, Kv1.2 and Kv1.5. Residues in grey correspond to portions of the S1 and S2 helices. Residues tested for fluorescence are marked with boxes, and those giving voltage-dependent deflections are bolded. (B and C) Representative fluorescence emissions recorded at +60 mV from I187C (B) and T219C (C) channels. (D) Mean G-V (filled symbols) and F-V (hollow symbols) relationships for I187C (diamonds) and T219C (triangles) (n = 6-15) compared to Kv1.2 A291C fast and slow fluorescence components from Figure 2.5C (hollow circles and squares, respectively). Boltzmann fits to I187C data gave V1/2 and k values of -50.6 ± 1.4 mV and 10.6 ± 0.5 mV for the F-V, and -17.7 ± 1.9 mV and 27.3 ± 1.0 mV for the G-V. T219C half-activation potential and slope factor values were -15.5 ± 2.6 mV and 22.8 ± 0.3 mV, and -70.8 ± 5.9 mV and 18.3 ± 2.0 mV for the G-V and F-V relationships.  61  Discussion  Kv1.2 A291C rapid fluorescence quenching reports fast S4 displacement during channel activation In Kv1.2 it is only the fluorophores attached to cysteines placed at A291, L290, and M288 at the top of S4 and in the S3-S4 linker that detect rapid quenching as part of the overall fluorescence signal during depolarization (Figure 2.3B). From closed and open state models of Kv1.2 (Supplemental Figure A.4, Figure 2.11), the positions of these amino acids suggest that rapid quenching is best detected by residues at the top of S4 that face either S5 (of the adjacent subunit) or the voltage sensor domain in the closed state, likely due to the positioning of the fluorescent probe in an environment subject to significant voltage-dependent change.  In  contrast, residues that face away from protein, S289 and I292, and fluorophores attached to sites deeper in S4, gave only smaller slow signals, or no signal at all, which suggested more limited changes in environment during depolarization.  62  Figure 2.11. Proposed model of Kv1.2 activation based on TMRM fluorescence. (A) Closed-state structure of Kv1.2 A291C-TMRM, based on the closed-state model of Pathak et al. (2007), shown from the top. Only one voltage sensing domain (yellow) and the pore domain from an adjacent subunit (green) are shown. Residues tested are labelled and highlighted based on their ability (blue) or inability (orange) to yield voltage-dependent fluorescence; the A291C mutation is shown in red. The TMRM molecule is modeled within the external aqueous vestibule between S4 and S1-S3 as a spherical structure of carbon (green), oxygen (red) and nitrogen (blue) atoms. (B) Open-state structure of Kv1.2 A291C-TMRM. The view shown, looking down upon the channel, is similar to that in (A) for the closed-state channel, relative to the pore domain, in particular S5. (C and D) Side views of the closed- and open-state structures of Kv1.2 A291C-TMRM seen in A and B. Transmembrane helices of one subunit are as labelled in each panel.  The similarity of the fluorescence reports from probes attached to neighbouring residues, as well as the periodicity in the appearance of the rapid quenching component, support some degree of secondary structure in the S3-S4 linker, in accordance with data from an alanine-scan  63  of these residues (Li-Smerin et al., 2000; Li-Smerin and Swartz, 2001). The Kv1.2 crystal structure data is unresolved within the linker regions (Long et al., 2005a; Long et al., 2005b), but in the Kv1.2/Kv2.1 paddle chimera open-state structure (Long et al., 2007) there does appear to be some secondary structure. The rapid quenching observed in Kv1.2 could report one of two possible conformational changes. It could indicate an outward movement of S4, bringing the TMRM probe into a more hydrophilic environment or altering its orientation with respect to nearby quenching amino acids. Alternatively, it could report a rotation of the S4 that, as far as the probe and its environment is concerned, leads to a rapid quenching of TMRM fluorescence. Likely, it is some combination of both, as the open state model of Kv1.2 (Figure 2.11B, D) places the A291C residue in an extruded rotated position, away from the external cavity occupied in the resting state. Recent structural studies have questioned the extent to which this extrusion away from the pore accurately reflects the open state of Kv1.2 (Lewis et al., 2008), but even a more conservative deviation of S4 away from the pore would support these findings. In Shaker, the mainly fast fluorescence emission has been shown to have a voltagedependence similar to (Cha and Bezanilla, 1997) or identical (Mannuzzu et al., 1996) to that of the Q-V relationship. Here, in the first published recordings of Kv1.2 gating currents, we also observed a strong correlation between the voltage dependencies of fast fluorescence quenching and gating current measurements (Figure 2.6B). Small differences in the time course and voltage-dependence of gating charge movement and fluorescence quenching may reflect imperfect separation of the fast and slow components of fluorescence, or simply methodological differences between mammalian and oocyte cellular models. It should be noted that the probe is also reporting on local changes in S4, while other regions of the voltage sensor, in particular S2 and S3, contain charged amino acids which may respond to changes in membrane potential  64  (Seoh et al., 1996) and would be accounted for in a gating charge measurement, but to a lesser degree or not at all in the fluorescence signal from S4 residues. The fast quenching Kv1.2 A291C signals from both of the Kv1.5-Kv1.2 chimeric channels also resulted in F-V relationships identical to the Q-V (Figure 2.8F), and interestingly, when the Shaker S1-S2 linker (7 amino acids shorter than Kv1.2) was inserted, the resulting fluorescence (Figure 2.9A), was monophasic. From the rapid fluorescence quenching seen in all constructs by A291C-TMRM we can conclude that a rapid S4 movement occurs in Kv1.2 that is responsible for the appearance of gating charge – and which eventually leads to channel opening.  Kv1.2 A291C fluorescence also reports slower voltage-dependent rearrangements of the voltage sensor domains A major novel finding in this work is the description of a prominent voltage-dependent slow phase of fluorescence quenching that originates from movements of the S1-S3 voltage sensor domains of Kv1.2, and which has not been described in prior studies of Shaker or Kv1 channels (Mannuzzu et al., 1996; Cha and Bezanilla, 1997; Claydon et al., 2007a; Vaid et al., 2008). In Shaker, small slow changes in fluorescence at depolarized potentials were correlated to events in the pore underlying P/C-type inactivation (Loots and Isacoff, 1998; Loots and Isacoff, 2000), while such changes in Kv1.5 A397C fluorescence were linked to changes in pore structure at the level of the selectivity filter (Vaid et al., 2008). Both Kv1.2 and Kv1.5-Kv1.2 chimera channels (Figure 2.8) retain a small slow quenching component at depolarized potentials, suggesting that a similar conformational change may occur in Kv1.2, in the absence of an observable inactivation process. However, none of these channels exhibited such a large slow fluorescence as Kv1.2, nor with such a left-shifted voltage-dependence.  65  The experiments presented here suggest that reversal of these slow movements are obligatory for channel deactivation (Figures 2.7, 2.8).  They may be coupled to the same  molecular movements, or the reoriented voltage sensing domain may interact with either the activated pore or S4, preventing or altering the time course of channel closure. We cannot be sure if the slow reorientation leads to a stabilized activated/open state at positive potentials once the movement is complete, since we have not carried out experiments to investigate open-state stability directly. However, this slow reorientation process of the voltage sensor is reminiscent of S4 entry into a relaxed state, as determined by Bezanilla and colleagues in Ci-VSP (VillalbaGalea et al., 2008). In their model of activation gating, the activated state of S4 was short lived before entry to this state, though this change in state did not appear to affect the activated state of the effector portion of the protein (i.e. the coupled enzyme or pore moiety). The S1-S2 linker of Kv1.2 is required for the slow quenching observed in A291C, as substitution of this region with that of Kv1.5 or Shaker was sufficient to either abolish or prevent detection of this slow fluorescence change (Figures 2.8, 2.9). The findings point to an integrated action of the S1-S4 elements of the voltage sensor during channel function, and there is evidence in the literature to suggest that disruption of S1 or the S1-S2 linker function alters activation gating. In Kv7.1 channels, the S1 helix has been suggested to help “steer” S4 motion through interactions occurring during gating (Haitin et al., 2008), and crosslinking S1 to the pore domain through cysteine-cysteine interactions has also been shown to interfere with function of KvAP channels, suggesting that some motion of this region may occur (Lee et al., 2009). Localized interactions within the S1-S2 linker have also been shown to modulate channel activation. In chimeric Kv2.1 channels in which the S1-S2 linker was replaced with that from Kv1.2, there was a 34 mV leftward shift in the G-V relationship and a 2-3 fold decrease in the time constants for  66  activation, suggesting that the S1-S2 region was important in channel stability and capable of modulating gating (Koopmann et al., 2001). Structurally, the S1-S2 linker of Kv1.2 channels has been suggested to form a coiled loop structure near the external surface of the membrane (Zhu et al., 2003), suggesting flexibility in this region. Differences in primary sequence of this region between Kv1.2 and Shaker or Kv1.5 (Figure 2.10A), and the S3-S4 linker (Figure 2.1), could have large effects on the positioning of this flexible segment. In Kv1.2, this may contribute to the positioning of this region with respect to the A291C S4 microenvironment.  Additionally, all three channels possess extracellular  glycosylation sites in the S1-S2 linker, at a similar position in the linker, although their positioning with respect to the transmembrane helices may differ based on the number of amino acids on either side of the glycosylation site (Zhu et al., 2003) (also see Figure 2.10A). Kv1.2 glycosylation has been shown to modulate Kv1.2 activation gating (Watanabe et al., 2007), and so may interact with the voltage sensor through positioning of the S1-S2 linker, in a manner different than other Kv1 channels.  Slow voltage-dependent rearrangement of the voltage sensor domains is confirmed by fluorophores placed in S1-S2 The fluorophores attached at I187 or T219, which would be likely to occupy a different microenvironment than at A291C, do not detect rapid movement associated with S4 displacement, but the slow movement within S1-S3 seen by A291C-TMRM can be detected by TMRM attached within the S1-S2 region (Figure 2.10).  Fluorescence quenching at these  residues accurately tracks the time course of slow movement of the voltage sensor, while the voltage-dependent fluorescence change of T219-TMRM overlays exactly the slow component of A291C-TMRM (Figure 2.10D). The more faithful recapitulation of the voltage-dependence of  67  slow fluorescence from T219C compared to I187C may be in part due to the fact that fluorophores in S2 or S4 are more peripheral to the protein core, closer to lipid and solvent, and therefore experience more significant environmental changes than a fluorophore within S1. Alternatively, the slow movement could be more prevalent within the S2 helix than in S1. Interestingly, the fluorescence recorded from the homologous T276C residue in Shaker (Cha and Bezanilla, 1997) exhibits a considerably faster quenching component, and a somewhat smaller leftward shift relative to the Q-V relationship (~20 mV). Cha and Bezanilla suggested that the S2 helix may undergo voltage-dependent movement prior to S4, facilitating S4 activation but carrying only a small amount of gating charge (Cha and Bezanilla, 1997). However, in Kv1.2, the much slower time course of S1-S3 rearrangement suggests that the motion cannot precede or constrain S4 movement, and indeed it continues well beyond the time required for full channel activation (Figure 2.2). While S2 does contain negatively charged side chains that appear to help stabilize S4 in the resting and activated states (Seoh et al., 1996; Long et al., 2005b; Long et al., 2007), the minimal voltage-dependence of the S1-S3 rearrangement based on time constant data (Figure 2.5B) suggests that little net charge movement within the electric field occurs, and given the extended time course of this rearrangement it is not surprising that we were unable to resolve any associated gating charge movements. The data from TMRM attached to A291C suggests that the most extreme inward limit of voltage-dependent movement occurs at approximately -120 mV (Figure 2.4). We also conclude that the voltage sensor domain movement, measured as a change in S3-S4 linker environment and S1-S2 rearrangements, is freely reversible between -120 mV and -50 mV (Figure 2.4C), or that any holding potential-dependent effects on mobility have reversed within 100 ms of return to more negative potentials. This is in agreement with previous work that has characterized the  68  voltage sensor as a fluid portion of the protein (Larsson et al., 1996; Yusaf et al., 1996; Baker et al., 1998; Wang et al., 1999), but different from functional studies in Shaker channels in which prolonged depolarization leads to a hyperpolarization of the returning Q-V relationship (Olcese et al., 1997). This hyperpolarization is recapitulated in the Shaker A359C fluorescence report of S4 movement (Figure 2.4D), but was not seen in Kv1.2.  This may reflect the inclusion of  rearrangements involving the entire voltage sensing domain in the Kv1.2 fluorescence record, and not just the S4 translocation measured by the rapid initial quenching.  Conclusion This paper represents the first study of the Kv1.2 voltage sensor domain using fluorimetry and gating currents, and highlights at least two independent conformational changes in this region in response to depolarization. A prominent fast change in fluorescence emission, at depolarized potentials, correlates well with channel gating currents and thus likely reports on rapid changes in S4 environment (i.e. translocation during depolarization). At more negative potentials there is a large slow quenching component reflecting fluorophore environment changes relative to the rest of the voltage sensor (particularly the S1-S2 region), that does not appear to be required for channel opening to occur, and persists well after ionic current has reached its peak. Reversal of this slow change matches the voltage-dependence and time course of channel deactivation and appears to be rate limiting for channel closure. This is the first report of such a slow protein movement within the voltage-sensing domain, and highlights both potential differences in channel gating between Kv1.2 and other Kv1 channels, and the involvement/motion of the entire Kv1.2 voltage sensing domain during activation.  69  Acknowledgments  This work was supported by grants from the Heart and Stroke Foundation of British Columbia and Yukon and the Canadian Institutes of Health Research to D. Fedida. A.J. Horne was supported by a Postgraduate Scholarship from the Natural Sciences and Engineering Research Council of Canada. We wish to thank Dr. Zhuren Wang for assistance with gating current measurements, Dr. David Steele for help with designing the chimera constructs, and Ka Kee Chiu and Kyung Hee Park for assistance with cell culture.  70  References  Baker, O. S., H. P. 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S4-based voltage sensors have three major conformations. Proc Natl Acad Sci U S A 105:17600-17607. Wang, M. H., S. P. Yusaf, D. J. S. Elliott, D. Wray, and A. Sivaprasadarao. 1999. Effect of cysteine substitutions on the topology of the S4 segment of the Shaker potassium channel: implications for molecular models of gating. J Physiol 521:315-326. Watanabe, I., J. Zhu, J. J. Sutachan, A. Gottschalk, E. Recio-Pinto, and W. B. Thornhill. 2007. The glycosylation state of Kv1.2 potassium channels affects trafficking, gating, and simulated action potentials. Brain Res 1144:1-18. Yarov-Yarovoy, V., D. Baker, and W. A. Catterall. 2006. Voltage sensor conformations in the open and closed states in ROSETTA structural models of K+ channels. Proc Natl Acad Sci U S A 103:7292-7297. 73  Yusaf, S. P., D. Wray, and A. Sivaprasadarao. 1996. Measurement of the movement of the S4 segment during the activation of a voltage-gated potassium channel. Pflugers Arch 433:91-97. Zhu, J., I. Watanabe, A. Poholek, M. Koss, B. Gomez, C. Yan, E. Recio-Pinto, and W. B. Thornhill. 2003. Allowed N-glycosylation sites on the Kv1.2 potassium channel S1-S2 linker: implications for linker secondary structure and the glycosylation effect on channel function. Biochem J 375:769-775.  74  Chapter 3: Voltage-dependent changes in Shaker and Kv1.2 fluorescence after transfer of Kv1.2 or Kv1.5 extracellular linkers1  1  A version of this chapter will be submitted for publication. Horne A.J., Peters C.J., & Fedida D. (2010) Voltagedependent changes in Shaker and Kv1.2 fluorescence after transfer of Kv1.2 or Kv1.5 extracellular linkers.  75  Introduction  When a fluorophore is linked to a cysteine introduced at the extracellular end of the S4 voltage-sensing domain of voltage-gated potassium channels like Shaker (A359C), Kv1.5 (at the homologous A397C residue) and Kv1.2 (A291C) there is a rapid quenching of fluorescence emission upon depolarization. In most voltage-gated ion channels tested, the time course and voltage-dependence of gating charge movement during depolarization approximate the time course of the rapid fluorescence quenching and the fluorescence-voltage relationships (Mannuzzu et al., 1996; Cha and Bezanilla, 1997; Savalli et al., 2006; Figure 2.6). This suggests that these rapid changes in fluorescence reflect conformational changes at the outer end of S4 that underlie Kv activation gating. Although the core domains of potassium channels are well conserved among all families in terms of primary and secondary sequence (Doyle et al., 1998; Jiang et al., 2003; Long et al., 2005), the voltage-dependent fluorescence emission from Shaker, Kv1.5 and Kv1.2 channels differ quite strikingly beyond the initial fast quenching component. A small secondary quenching of fluorescence emission is observed in Shaker A359C over short depolarizations (Claydon et al., 2007), and an opposite dequenching (increase in fluorescence emission) is seen in Kv1.5 A397C (Vaid et al., 2008). Both of these changes in emission have been shown to correlate with events occurring in the pore domain, like inactivation (Loots and Isacoff, 1998; Claydon et al., 2007; Vaid et al., 2008). In Kv1.2 channels a secondary slow quenching component, much larger than that observed in Shaker, and with a more negative voltage-dependence, correlates well with events involving the S1 and S2 region of the voltage sensor domain (Figures 2.5-2.10), in the absence of a resolvable gating charge correlate. Together, these findings suggest that despite similarities in ionic conductance and S1-S6 structure, aside from an initial common translocation of the S4 there are either dynamic  76  differences between regional interactions of Kv1 channels arising from differences in channel structure/gating, or the unique extracellular linkers of the different channels allow detection of slower changes in the channel during activation. This is not unprecedented, as it is known that in Shaker channels the S3-S4 linker length determines the magnitude and time course of fluorescence emission. Deletion of 26 of 31 amino acids in the S3-S4 linker reduces voltagedependent changes in TMRM emission from M356C by an order of magnitude (Sorensen et al., 2000), and the rate of fluorophore quenching is slowed compared to the same residue with the full-length linker (Mannuzzu et al., 1996; Cha and Bezanilla, 1997). To further investigate the role of the extracellular S1-S2 and S3-S4 linkers of Kv1.2, Kv1.5 and Shaker channels in reporting channel-specific changes in conformation via TMRM fluorescence emission, we constructed a series of chimeric channels involving these linker segments. We found that single or double linker substitutions into Kv1.2 prevented detection of the slow quenching component of A291C fluorescence, but that Kv1.2 linker insertion into Shaker did not allow detection of slow events in that channel. Fast fluorescence quenching was similar between all chimeras tested, independent of linker length or structure. From these data, it appears that secondary changes in S4 environment are dependent on the unique channel-specific linker composition, although other regions of the voltage-sensing domain certainly appear to be involved in these changes, based on incomplete transfer of these channel-specific fluorescence emissions.  77  Materials and methods  Molecular biology and RNA preparation Shaker Δ6-46 C245V A359C, Kv1.2 A291C and Kv1.5 C245V A397C channels were expressed in Xenopus laevis oocytes using a modified pBluescript SKII expression vector (pEXO) (a gift from A. Sivaprasadarao, University of Leeds, UK), as previously described (Vaid et al., 2008). S3-S4 linker chimera channels were generated using a two-step PCR protocol where overlapping primers encoding for the new linker sequence were paired with complementary flanking primers within the channel. Products of these reactions were then amplified through a second PCR using the same flanking primers, and the resultant channel fragments containing the core channel with new linker segment were then subcloned into the starting construct utilizing appropriate restriction endonuclease enzymes. For construction of the S1-S2 linker chimeras, both channels were first subcloned into a pBluescript SK- vector (EcoRI) in which the XbaI polylinker site had been removed (Klenow). Silent mutations were made in Kv1.2 for the purposes of addition of NsiI and XbaI sites into the equivalent positions to Shaker, flanking the S1-S2 linker.  After removal of unwanted  occurrences of these restriction sites at other locations in both channels (again through silent mutagenesis), the Shaker S1-S2 linker was subcloned into Kv1.2 and vice versa using BbsI and BsmI restriction endonucleases. The S1-S2/S3-S4 double linker chimera channels were generated with a similar procedure to the S3-S4 linker constructs, utilizing the S1-S2 linker chimera channels as starting sequences.  78  Oligonucleotide primers were synthesized by either Sigma-Genosys (Oakville, ON, Canada) or Integrated DNA Technologies (Coralville, IA), and mutations were generated using the Stratagene QuikChange kit (Stratagene, La Jolla, CA).  Mutations were confirmed by  sequencing the constructs using the core facility unit at the University of British Columbia. cRNA was synthesized from linear cDNA using the mMessage mMachine T7 Ultra cRNA transcription kit (Ambion, Austin, TX).  Oocyte preparation and injection Xenopus laevis frogs were terminally anesthetized by immersion in 2 mg/mL tricaine methanesulphonate (Sigma-Aldrich, Mississauga, ON, Canada).  Stage V-VI oocytes were  isolated and defolliculation was performed through a combination of collagenase treatment, involving mild agitation in 1 mg/mL collagenase Type 1a for approximately 1 hour, and manual defolliculation. Between defolliculation and injection, oocytes were incubated for 1-18 hours in Barth’s solution, which contained (in mM), 88 NaCl, 1 KCl, 2.4 NaHCO3, 0.82 MgSO4, 0.33 Ca(NO3)2, 0.41 CaCl2, 20 HEPES, titrated to pH 7.4 using NaOH. Injection of 50 nL of cRNA encoding the construct of interest was performed using a Drummond digital microdispenser (Fisher Scientific, Ottawa, ON, Canada), followed by incubation at 19°C in Barth’s solution. Currents were recorded 1-4 days after injection.  All animal protocols were performed in  accordance with Canadian Council of Animal Care guidelines. Unless otherwise stated, all chemicals were purchased from Sigma-Aldrich  79  Two-electrode voltage clamp Oocytes were placed in a bath chamber that was perfused with control ND96 external solution containing (in mM), 96 NaCl, 3 KCl, 1 MgCl2, 2 CaCl2, and 5 HEPES, titrated to pH 7.4 with NaOH. Microelectrodes were filled with 3 M KCl and had resistances of 0.2 to 2.0 MΩ. Unless otherwise stated, the holding potential between protocols was -80 mV. Voltage control and data acquisition were achieved with a Warner Instruments OC-725C amplifier (Hamden, CT), and Axon Digidata 1322 A/D converter (Axon Instruments, Foster City, CA), connected to a personal computer running pClamp9 software (Molecular Devices Corp.).  Voltage clamp fluorimetry Fluorimetry was performed simultaneously with two-electrode voltage clamp. Labelling of the oocytes with 5 µM tetramethylrhodamine-5-maleimide (TMRM; Invitrogen, Carlsbad, CA) dye was performed for 30 minutes at 10°C in a depolarizing solution containing (in mM), 98 KCl, 1 MgCl2, 2 CaCl2, and 5 HEPES, titrated to pH 7.4 using KOH. Signals were recorded via a Nikon TE300 inverted microscope with an Epi-Fluorescence attachment and a 9124b Electron Tubes photomultiplier tube (PMT) module (Cairn Research, Kent, UK). TMRM dye, exhibiting maximal light absorption at 542 nm, was excited by light from a 100 W mercury lamp filtered through a 525 nm band pass filter and reflected by a 560 nm dichroic mirror through a 20x objective lens focussed on the oocyte. Emitted light (maximal emission of ~567 nm) was collected by the objective lens and passed through the dichroic mirror to the PMT module. To record fluorescence intensity, voltage signals from the PMT were digitized using an Axon Digidata 1322 A/D converter and passed to a computer running pClamp9 software. To minimize fluorophore bleaching, a Uniblitz computer-controlled shutter (Vincent Associates, Ottawa, ON,  80  Canada) was used, and opened shortly prior to application of voltage-clamp pulses. For 100 ms pulses the fluorescence signal sampling frequency was 20 kHz; traces were averaged, with each signal representing the average of five sweeps, and were filtered offline at 1000 Hz. To account for any photobleaching of fluorophores that may occur during shutter opening, control fluorescence data were recorded in the absence of any change in voltage and subtracted from the voltage-dependent signal. Fluorescence records were normalized to baseline emission in the absence of a change in voltage, to control for cell-to-cell variability, and fluorescence-voltage (F-V) relationships are shown normalized to maximum and minimum levels of emission over the voltage ranges tested.  Data analysis Conductance-voltage (G-V) relationships were derived using normalized chord conductance, which was calculated by dividing peak current by the driving force based on the potassium equilibrium potential (internal potassium concentration was assumed to be 99 mM). F-V relationships were calculated based on the peak change in emission of the total trace or, in some stated cases, the individual components of the signal. All G-V, gating charge-voltage (QV), and most fluorescence-voltage (F-V) relationships (see below) were fit with a single Boltzmann function,  y=  1 ⎡V − V ⎤ 1 + exp ⎢ 1 / 2 ⎥ ⎣ k ⎦  81  where y is the conductance normalized with respect to the maximal conductance, V1/2 is the halfactivation potential, V is the test voltage, and k is the slope factor. Unless otherwise indicated, data reported throughout the text and figures are expressed as mean ± SEM.  82  Results  A homologous Shaker/Kv1.2 S4 residue produces different fluorescence emission Representative current and fluorescence traces for Shaker A359C-TMRM and Kv1.2 A291C-TMRM can be seen in Figures 3.1A and 3.1B respectively. Shaker and Kv1.2 both exhibit a rapid quenching of TMRM emission upon depolarization, which has been shown in both channels to reflect S4 movement upon channel activation and is well correlated with gating current measurements (Mannuzzu et al., 1996; Cha and Bezanilla, 1997; Figure 2.6). In addition, Kv1.2 fluorescence reports an additional slow quenching component that appears at very negative potentials (e.g. -120 mV and -60 mV in Figure 3.1B), and persists at very positive depolarizations (arrow) along with the fast component. A slow increase in Kv1.2 fluorescence is observed on repolarization that likely reflects reversal of the events underlying the slow quenching upon depolarization. Ionic currents and conductance-voltage (G-V) relationships (Figure 3.1C) are similar between Kv1.2 and Shaker, with half-activation potentials of -12.9 ± 2.8 mV and -14.8 ± 1.5 mV respectively, yet the fluorescence-voltage (F-V) relationship of Kv1.2 has a V1/2 ~27 mV negative to that of Shaker (-61.4 ± 1.2 mV compared to -34.7 ± 2.2 mV), largely due to the presence of this additional slow quenching component (Figure 2.8).  83  Figure 3.1. Shaker and Kv1.2 fluorescence from a homologous residue in the S3-S4 linker exhibit unique fluorescence effects upon depolarization. (A) Representative ionic (top) and fluorescence (bottom) records from Shaker A359C-TMRM channels in response to 100 ms changes in voltage between -120 mV and +60 mV. Data were collected every 10 mV, but only selected traces are shown for clarity. Cells were held at -80 mV. (B) Representative ionic (top) and fluorescence (bottom) records from Kv1.2 A291C-TMRM in response to the identical protocol used in (A). (C) Conductance-voltage (G-V, filled symbols) and fluorescence-voltage (F-V, hollow symbols) for Shaker A359C (circles) and Kv1.2 A291C (squares). Data are shown mean ± SEM (n = 7-10), and individual cell records were well fit with a Boltzmann function. Shaker half-activation (V1/2) and slope (k) parameters were -14.8 ± 1.5 mV and 21.4 ± 1.5 mV for the G-V and -34.7 ± 2.2 mV and 25.9 ± 2.4 mV for the F-V, respectively. In Kv1.2, mean V1/2 and k parameters were -12.9 ± 2.8 mV and 30.2 ± 1.3 mV, and -61.4 ± 1.2 mV and 22.8 ± 0.9 mV respectively. The arrow indicates the slow quenching phase of Kv1.2 A291C fluorescence. (D) ClustalW amino acid alignment of the transmembrane (S1-S6) and linker regions of Shaker and Kv1.2. Transmembrane regions S1-S6 are as noted, and the S1-S2 and S3-S4 linker regions are indicated by the grey boxes. TMRM fluorescence was recorded from Shaker/Kv1.2 channels labelled at A359C/A291C, at the point indicated by the black circle in the S3-S4 linker.  84  Shaker and Kv1.2 extracellular linkers differ in length and sequence The sensitivity of TMRM and other fluorophores to local microenvironments suggests that the changes in protein structure surrounding S4 during activation could be responsible for observed differences in TMRM fluorescence between Shaker and Kv1.2. An alignment of the membrane-spanning (S1 to S6) regions of both channels (Figure 3.1D) highlights large differences in the extracellular linker regions between the S1 and S2 transmembrane helices, and between S3 and S4 (grey boxes). Fluorescence measurements from Shaker channels in which the S3-S4 linker had been truncated resulted in profound changes in the observed phenotype (Sorensen et al., 2000); thus, this region in particular appears to be important in contributing to voltage-dependent changes in emission.  Transfer of Kv1.2 or Kv1.5 secondary quenching events are not conferred with the S3-S4 linker alone To investigate whether the S3-S4 linker confers channel specific fluorescence quenching patterns, we constructed chimeric channels in which this region of Shaker was inserted into Kv1.2 (Kv1.2/Shaker S3S4L, Figure 3.2A), and vice versa (Shaker/Kv1.2 S3S4L, Figure 3.2B). In Kv1.2, replacing the S3-S4 linker with that of Shaker maintained rapid voltage-dependent quenching, but caused a loss of the slow component of A291C fluorescence (Figure 3.2C). As a result, the F-V relationship was right-shifted relative to that of Kv1.2 A291C (V1/2 = -44.2 ± 3.2 mV), close to the voltage-dependence of Shaker A359C fluorescence (Figure 3.2E). However, when the S3-S4 linker of Kv1.2 was inserted into Shaker, no additional slow quenching component (Figure 3.2D) or shift in the F-V relationship (Figure 3.2E) was observed (V1/2 = -39.0 ± 1.9 mV).  Modest increases in the slow, pore-associated (Claydon et al., 2007)  component of Shaker fluorescence were observed in Shaker/Kv1.2 S3S4L, but this effect was  85  limited to potentials greater than 0 mV. In addition, changes in the structure and/or length of this linker also appear to have effects on the voltage-dependence of activation, as both chimeras showed a 20-30 mV positive displacement in their G-V relationships.  86  Figure 3.2. Kv1.2 slow fluorescence quenching is absent in S3-S4 linker chimera channels, and not conferred to Shaker with linker transfer. (A and B) Cartoon models of one potassium channel subunit demonstrating the Kv1.2 (red) and Shaker (blue) S3-S4 linker chimeras. (C and D) Representative ionic (top) and fluorescence (bottom) traces for Kv1.2/Shaker S3S4L “A291C” (C) and Shaker/Kv1.2 S3S4L “A359C” (D) in response to the same protocol as described in Figure 3.1A. (E) Conductance-voltage (G-V, filled symbols) and fluorescence-voltage (F-V, hollow symbols) for Kv1.2/Shaker S3S4L (circles) and Shaker/Kv1.2 S3S4L (squares). Data are shown mean ± SEM (n = 5-8), and individual cell records were well fit with a Boltzmann function. Kv1.2/Shaker S3S4L half-activation (V1/2) and slope (k) parameters were 7.9 ± 1.9 mV and 21.4 ± 1.3 mV for the G-V and -44.2 ± 3.2 mV and 20.1 ± 0.8 mV for the F-V, respectively. In Shaker/Kv1.2 S3S4L, mean V1/2 and k parameters were 21.1 ± 2.1 mV and 19.9 ± 0.6 mV, and -39.0 ± 1.9 mV and 19.4 ± 0.6 mV respectively.  87  In the Kv1.5 channel at depolarized potentials, TMRM at A397C reports rapid quenching changes in fluorescence similar to those seen in both Kv1.2 and Shaker, followed by a unique increase in fluorescence emission (Figure 3.3C) that appears to be a report of changes in pore stability (Vaid et al., 2008; Vaid et al., 2009). Insertion of the Kv1.5 S3-S4 linker into Shaker (Shaker/Kv1.5 S3S4L) did not transfer this rapid unquenching phenomenon (Figure 3.3B), and the F-V relationship overlaps that of Shaker alone (V1/2 = -32.8 ± 2.3 mV). The half-activation potential of the chimera was also right-shifted relative to Shaker A359C, with a half-activation potential of 8.9 ± 7.1 mV. Despite successful RNA translation, attempts to record from the reverse chimera (Kv1.5/Shaker S3S4L) were unsuccessful.  88  Figure 3.3. Secondary Kv1.5 fluorescence emission is not conferred by S3-S4 linker replacement in Shaker. (A and B) Representative ionic (top) and fluorescence (bottom) traces for Kv1.5 A397C (A) and Shaker/Kv1.5 S3S4L “A359C” (B) in response to the same protocol as described in Figure 3.1A. (C) Conductance-voltage (G-V, filled symbols) and fluorescencevoltage (F-V, hollow symbols) for Shaker/Kv1.5 S3S4L (circles) and Kv1.5 A397C (squares). Data are shown mean ± SEM (n = 3-8), and individual cell records were well fit with a Boltzmann function, in the case of Kv1.5 at potentials positive to -80 mV. Kv1.5 half-activation (V1/2) and slope (k) parameters were 8.9 ± 7.1 mV and 24.7 ± 3.3 mV for the G-V and -32.8 ± 2.3 mV and 27.3 ± 9.1 mV for the F-V, respectively. In Shaker/Kv1.5 S3S4L, mean V1/2 and k parameters were 11.3 ± 5.0 mV and 25.0 ± 1.2 mV, and -47.5 ± 5.0 mV and 18.4 ± 0.7 mV respectively. (D) ClustalW amino acid alignment of the S3-S4 linker regions of Shaker and Kv1.5. Portions of the transmembrane S3 and S4 regions are as noted, and the S3-S4 linker regions are indicated by the grey box. TMRM fluorescence was recorded from Shaker/Kv1.5 channels labelled at A359C/A397C, at the point indicated by the black circle in the S3-S4 linker.  Overall, insertion of foreign S3-S4 linker segments into Shaker strongly suggests that regions of Kv1.2 and Kv1.5 other than the linkers themselves confer their unique fluorescence properties. As well, S4 movement and translocation, which is tracked by the rapid emission quenching, is relatively unaffected by the linker exchanges, based on the similarities in the voltage-dependent fluorescence signals and the voltage-dependence of the F-V relationships in all of the Shaker chimeras.  89  Shaker S1-S2 linker transfer into the Kv1.2 channel prevents slow fluorescence quenching of the A291C microenvironment Characterization of the Kv1.2 fluorescence at A291C (Figure 2.8-2.10) revealed that the slow changes in emission were dependent upon the presence of the extracellular S1-S2 linker. In Figure 3.4, where a chimeric channel in which this linker region was replaced with that of Shaker (Kv1.2/Shaker S1S2L, Figure 3.4A), robust ionic currents were seen with no evidence of slow quenching in the fluorescence during depolarization (Figure 3.4C). The F-V relationship reflects the absence of this slow component at negative potentials (up to approximately -60 mV), but was similar to Kv1.2 WT at more depolarized potentials, with a half-activation potential of -58.7 ± 5.4 mV.  90  Figure 3.4. Kv1.2 slow fluorescence quenching is lost in S1-S2 linker chimera channels, and not conferred to Shaker with linker transfer. (A and B) Cartoon models of the Kv1.2 (red) and Shaker (blue) S1-S2 linker chimeras; only one subunit is shown for each construct. (C and D) Representative ionic (top) and fluorescence (bottom) traces for Kv1.2/Shaker S1S2L “A291C” (C) and Shaker/Kv1.2 S1S2L “A359C” (D) using the protocol described in Figure 3.1A. (E) Conductance-voltage (G-V, filled symbols) and fluorescence-voltage (F-V, hollow symbols) for Kv1.2/Shaker S1S2L (circles) and Shaker/Kv1.2 S1S2L (squares). Individual cell records were well fit with a Boltzmann function. Kv1.2/Shaker S1S2L half-activation (V1/2) and slope (k) parameters were 23.8 ± 4.8 mV and 23.5 ± 2.7 mV for the G-V and -58.7 ± 5.4 mV and 14.7 ± 1.8 mV for the F-V, respectively. In Shaker/Kv1.2 S1S2L, mean V1/2 and k parameters were 18.8 ± 5.4 mV and 20.5 ± 2.6 mV, and -25.2 ± 10.4 mV and 18.8 ± 1.8 mV respectively. Data are shown mean ± SEM (n = 4-5).  91  Similar to the S3-S4 linker studies, the reverse chimera inserting the Kv1.2 S1-S2 linker into Shaker (Figure 3.4B) did not confer a Kv1.2 phenotype. There was no perceivable slow quenching component, and no difference from the Shaker F-V relationship (Figure 3.4E; V1/2 = -25.2 ± 10.4 mV). This suggests that the S1-S2 linker itself does not facilitate detection or confer this slow environmental change in the external voltage sensing domain. Similar to our observations with the S3-S4 linker chimera channels, replacement of the native S1-S2 linker with that from another channel also resulted in significant shifts in the G-V relationship, with the Kv1.2/Shaker S1S2L construct having a V1/2 of 23.8 ± 4.8 mV, and 18.8 ± 1.8 mV in Shaker/Kv1.2 S1S2L channels, suggesting that the linker type affects channel open probability.  The Kv1.2 double (S1-S2 and S3-S4) linker swap lacks a slow quenching component It is possible that both Kv1.2 extracellular S1-S2 and S3-S4 linkers are required to detect slow fluorescence changes upon depolarization, either through their direct interaction or modification of the environment around A291C. To investigate this possibility, we constructed the double-linker swap Kv1.2/Shaker S1S2,S3S4L (Figure 3.5A) and Shaker/Kv1.2 S1S2,S3S4L chimera channels.  Kv1.2/Shaker S1S2,S3S4L records (Figure 3.5B) showed no slow  fluorescence component, and were identical to Kv1.2/Shaker S1S2L at positive potentials in both fluorescence phenotype and voltage-dependence (Figure 3.5C; V1/2 = -55.2 ± 1.5 mV). A rightward shift was also observed in the G-V relationship (V1/2 = 6.7 ± 3.0 mV), though this was not quite as large as the S1-S2L chimera alone (Figure 3.4E). The reverse double linker chimera, inserting the Kv1.2 extracellular linkers in Shaker, did not result in functional channels. This result suggests that the Kv1.2 linkers themselves are not responsible for slow quenching of fluorescence emission, since they cannot confer this phenotype onto the Shaker core channel.  92  Figure 3.5. Interactions between the S1-S2 and S3-S4 linkers do not alone confer unique channel fluorescence phenotypes. (A) Cartoon models of one potassium channel subunit demonstrating the constructed Kv1.2 (red) and Shaker (blue) double linker (S1S2,S3S4) chimera. (B) Representative ionic (top) and fluorescence (bottom) traces for Kv1.2/Shaker S1S2,S3S4L “A291C” in response to the same protocol as described in Figure 3.1A. (C) Conductancevoltage (G-V, filled symbols) and fluorescence-voltage (F-V, hollow symbols) for Kv1.2/Shaker S1S2,S3S4L. Data are shown mean ± SEM (n = 4-5), and individual cell records were well fit with a Boltzmann function. Half-activation (V1/2) and slope (k) parameters were 6.7 ± 3.0 mV and 23.4 ± 1.6 mV for the G-V and -55.2 ± 1.5 mV and 13.3 ± 0.3 mV for the F-V, respectively.  93  Discussion  Extracellular linkers alone cannot confer secondary fluorescence changes From our data, it appears clear that unique voltage-dependent secondary changes in TMRM emission from S4 in Kv1.2 and Kv1.5 channels upon depolarization (Figure 3.1, 3.3A) cannot be transferred into Shaker with replacement of the extracellular S3-S4 linker (Figure 3.2B, Figure 3.3B), the S1-S2 linker (Figure 3.4C), or both extracellular linkers (Figure 3.5C). Conversely, insertion of either the S1-S2 or S3-S4 linker from Shaker into Kv1.2 resulted in a loss of this slow component as sensed from TMRM attached to the A291C equivalent residue. The inability to simply confer secondary quenching properties from Kv1.2 or Kv1.5 into Shaker, or to maintain these properties with insertion of Shaker linker regions, suggests complex channel-specific differences in structure and function.  Chimera effects on activation are likely not responsible for shifts in fluorescence The large rightward shift in the G-V relationship observed with every one of the chimeras tested may reflect altered voltage sensitivity of the proteins, or an alteration in the coupling between gating charge movement and activation. Furthermore, comparable 20-30 mV rightward shifts in both the F-V and G-V of Kv1.2/Shaker S3S4L (Figure 3.2E) suggest that overlay with the Shaker WT F-V may be an artefact of this altered voltage-dependence. However, there are two main lines of evidence that argue against this as the sole explanation.  Firstly, rapid  fluorescence quenching reminiscent of S4 translocation observed and reported in Shaker, BKCa channels, and the isolated fast component of Kv1.2 fluorescence (Mannuzzu et al., 1996; Cha and Bezanilla, 1997; Savalli et al., 2006; Figure 2.6) is preserved in this and all of the chimera channels, and appears very similar in both phenotype and voltage-dependence. Based on this  94  finding, we suspect that linker structure does not affect the mobility of the primary gating charge movement, though gating current measurements would be required to completely dispel this notion. Secondly, the Shaker/Kv1.2 S3S4L chimera correlates well with the reverse chimera in terms of both fluorescence and ionic conductance (Figure 3.2E); thus, even if S4 movement were altered, transfer of the S3-S4 linker clearly confers unique voltage-dependence to the core channel. Similarly, S1-S2 linker chimera channels show large G-V shifts in the absence of large effects on the voltage-dependence of fluorescence emission, aside from Kv1.2/Shaker S1S2L at potentials negative to -50 mV (Figure 3.4E).  Therefore, our data appear to support the  conclusion that the extracellular S1-S2 and S3-S4 linkers affect fluorescence emission independently from the observed effects on ionic conductance.  Extracellular linker alterations and glycosylation can have significant effects on channel gating Extracellular linkers of the voltage-sensing domain have been shown to be important both structurally and functionally in Kv channel function. In Shaker, studies altering the structure and length of the S3-S4 linker have reported changes in activation and deactivation properties (Mathur et al., 1997) as well as shifts in the G-V relationship (Gonzalez et al., 2000; Gonzalez et al., 2001). Kv2.1 channels containing the S1-S2 linker from Kv1.2 exhibit a 34 mV leftward shift in the G-V relationship and 2-3 fold decrease in activation time constants, suggesting that the S1-S2 region is important in channel stability and capable of modulating gating (Koopmann et al., 2001). In Kv1.2 channels, glycosylation has been shown to modulate activation gating (Watanabe et al., 2007) and similar effects have also been shown for Kv1.5 but not other channels in the Kv1 family (Schwetz et al., 2010).  The Kv1.2 S1-S2 linker contains a  95  glycosylation site (Zhu et al., 2003) that is conserved in Shaker and Kv1.5. However, its positioning with respect to the transmembrane helices may differ based on the number of amino acids on either side, which may affect interactions with S4 in different Kv1 channels and its effects on channel function (Zhu et al., 2009).  Fluorophore quenching groups are altered by changes in linker length and structure Differences both in the structures of linkers and their interaction(s) with the voltage sensor domain could potentially confer channel-specific fluorescence phenotypes. Furthermore, these interactions may involve a number of different regions of the protein, such that with removal or alteration of even one of these components, the report of these secondary movements could be lost. Linker residues themselves have also been shown contribute to the fluorophore environment as quenching groups. Shaker channels in which the majority of the S3-S4 linker had been deleted (reduced from ~31 amino acids to 5) exhibited a 10-fold decrease in fluorescence emission changes from several different residues. In addition, unique fluorescence emission was observed from A359C and other residues, which the authors interpreted as a revelation of different quenching groups in the absence of the lengthy Shaker S3-S4 linker (Sorensen et al., 2000).  The Shaker S3-S4 linker may insulate the TMRM probe from secondary changes In the work of Sorensen and colleagues, there does appear to be some report of a slow change in emission from A359C in the S3-S4 deletion mutant at depolarized potentials (Sorensen et al., 2000), the origin of which is unknown. The Shaker S3-S4 linker is 11 amino acids longer than that in Kv1.2 and contains several proline residues, which may confer a more rigid  96  secondary structure to the region. Thus, assuming a similar slow reorientation of the voltage sensing domain occurs in Shaker, the native S3-S4 linker structure may serve to mask these changes from a S4 fluorophore, or lead to differences in linker position that may remove the probe from the local microenvironment. In contrast, a shorter Kv1.2 S3-S4 linker may not fully surround the S4 microenvironment, enabling a secondary report of movement from more distant regions of the voltage sensor domain. Our Kv1.2/Shaker S3S4L and Shaker/Kv1.2 S1S2L data are consistent with such an explanation, where the unique structure and/or additional length of the Shaker S3-S4 linker may be oriented in such a way as to block or buffer TMRM from experiencing slow emission changes associated with this movement.  Kv1.2 S1-S2 linker length and/or glycosylation may confer secondary fluorescence Kv1.2 glycosylation within the S1-S2 linker has been shown to modulate voltagedependence and channel gating (Watanabe et al., 2007), suggesting that the linker and/or attached carbohydrate may be in close proximity to the S4 helix. It is possible that some sort of allosteric modulation instead confers this effect, but our fluorescence data from Kv1.2 suggests that these regions are in close proximity to one another. To the best of our knowledge, there are no equivalent studies in Shaker supporting a close proximity of the S1-S2 linker and S4. The S1S2 linker of Shaker is 7 amino acids shorter than in Kv1.2, meaning it may not reside in the local microenvironment of a fluorescent probe attached to A359C. Fluorescent labelling of the S1-S2 linker of Shaker show mainly fast changes in TMRM emission (Pathak et al., 2007) including at T276C (Cha and Bezanilla, 1997), the homologous residue to T219C in Kv1.2 which shows exclusively slow voltage-dependent quenching (Figure 2.10).  From our data it is unclear  97  whether differences in Shaker gating, linker structure, or both may explain the lack of slow fluorescence observed in Shaker/Kv1.2 S3S4L and Kv1.2/Shaker S1S2L.  S4 translocation is unaffected by linker length or structure All five chimeric channels tested showed such a voltage-dependent decrease in TMRM emission similar to that observed in WT Shaker and correlated with S4 translocation. Thus, a fluorophore positioned at the external end of S4 is in an ideal position to report rapid changes in local environment, independent of S3-S4 linker identity.  The similarity in phenotype and  voltage-dependence of these constructs implies that it is global changes in the voltage sensor domain relative to itself or to the pore domain, and/or conserved portions of the S3-S4 linker, that contribute to this environmental change.  Summary Secondary changes in structure relating to other regions of the voltage sensor domain or pore appear strongly dependent on the identity of the extracellular linker segments. However, the insertion of a channel-specific linker is not sufficient to confer these secondary changes, suggesting important interrelations between the more stable core components of the individual channels, and/or variations in the precise mechanisms of gating within the Kv1 family. Rapid changes in fluorophore emission prevail in all chimeras tested, suggesting that the primary mechanism of reporting S4 translocation derives from conserved regions of the linker and/or the transmembrane helices of the channel.  98  Acknowledgements  This work was supported by grants from the Heart and Stroke Foundation of British Columbia and Yukon and the Canadian Institutes of Health Research to D. Fedida. We wish to thank Dr. Thomas Claydon for his collection of preliminary data, Dr. David Steele for help with designing chimera constructs, Dr. Sam Goodchild for helpful discussion, and Ms. Ka Kee Chiu and Mrs. Kyung Hee Park for assistance with cell culture.  99  References  Cha, A. and F. Bezanilla. 1997. Characterizing voltage-dependent conformational changes in the Shaker K+ channel with fluorescence. Neuron 19:1127-1140. Claydon, T. W., M. Vaid, S. Rezazadeh, S. J. Kehl, and D. Fedida. 2007. 4-aminopyridine prevents the conformational changes associated with P/C-type inactivation in Shaker channels. J Pharmacol Exp Ther 320:162-172. Doyle, D. A., J. M. Cabral, R. A. Pfuetzner, A. L. Kuo, J. M. Gulbis, S. L. Cohen, B. T. Chait, and R. MacKinnon. 1998. The structure of the potassium channel: Molecular basis of K+ conduction and selectivity. Science 280:69-77. Gonzalez, C., E. Rosenman, F. Bezanilla, O. Alvarez, and R. Latorre. 2000. Modulation of the Shaker K+ channel gating kinetics by the S3-S4 linker. J Gen Physiol 115:193-207. Gonzalez, C., E. Rosenman, F. Bezanilla, O. Alvarez, and R. Latorre. 2001. Periodic perturbations in Shaker K+ channel gating kinetics by deletions in the S3-S4 linker. Proc Natl Acad Sci U S A 98:9617-9623. Jiang, Y., A. Lee, J. Chen, V. Ruta, M. Cadene, B. T. Chait, and R. MacKinnon. 2003. X-ray structure of a voltage-dependent K+ channel. Nature 423:33-41. Koopmann, R., A. Scholle, J. Ludwig, T. Leicher, T. Zimmer, O. Pongs, and K. Benndorf. 2001. Role of the S2 and S3 segment in determining the activation kinetics in Kv2.1 channels. J Membrane Biol 182:49-59. Long, S. B., E. B. Campbell, and R. MacKinnon. 2005. Crystal structure of a mammalian voltage-dependent Shaker family K+ channel. Science 309:897-903. Loots, E. and E. Y. Isacoff. 1998. Protein rearrangements underlying slow inactivation of the Shaker K+ channel. J Gen Physiol 112:377-389. Mannuzzu, L. M., M. M. Moronne, and E. Y. Isacoff. 1996. Direct physical measurement of conformational rearrangement underlying potassium channel gating. Science 271:213216. Mathur, R., J. Zheng, Y. Y. Yan, and F. J. Sigworth. 1997. Role of the S3-S4 linker in Shaker potassium channel activation. J Gen Physiol 109:191-199. Pathak, M. M., V. Yarov-Yarovoy, G. Agarwal, B. Roux, P. Barth, S. Kohout, F. Tombola, and E. Y. Isacoff. 2007. Closing in on the resting state of the Shaker K+ channel. Neuron 56:124-140. Savalli, N., A. Kondratiev, L. Toro, and R. Olcese. 2006. Voltage-dependent conformational changes in human Ca2+- and voltage-activated K+ channel, revealed by voltage-clamp fluorometry. Proc Natl Acad Sci U S A 103:12619-12624.  100  Schwetz, T. A., S. A. Norring, and E. S. Bennett. 2010. N-glycans modulate Kv1.5 gating but have no effect on Kv1.4 gating. Biochim Biophys Acta 1798:367-375. Sorensen, J. B., A. Cha, R. Latorre, E. Rosenman, and F. Bezanilla. 2000. Deletion of the S3-S4 linker in the Shaker potassium channel reveals two quenching groups near the outside of S4. J Gen Physiol 115:209-221. Vaid, M., T. W. Claydon, S. Rezazadeh, and D. Fedida. 2008. Voltage clamp fluorimetry reveals a novel outer pore instability in a mammalian voltage-gated potassium channel. J Gen Physiol 132:209-222. Vaid, M., A. Horne, T. Claydon, and D. Fedida. 2009. Rapid outer pore movements after opening in a KV1 potassium channel are revealed by TMRM fluorescence from the S3S4 linker, and modulated by extracellular potassium. Channels (Austin) 3:3-5. Watanabe, I., J. Zhu, J. J. Sutachan, A. Gottschalk, E. Recio-Pinto, and W. B. Thornhill. 2007. The glycosylation state of Kv1.2 potassium channels affects trafficking, gating, and simulated action potentials. Brain Res 1144:1-18. Zhu, J., E. Recio-Pinto, T. Hartwig, W. Sellers, J. Yan, and W. B. Thornhill. 2009. The Kv1.2 potassium channel: the position of an N-glycan on the extracellular linkers affects its protein expression and function. Brain Res 1251:16-29. Zhu, J., I. Watanabe, A. Poholek, M. Koss, B. Gomez, C. Yan, E. Recio-Pinto, and W. B. Thornhill. 2003. Allowed N-glycosylation sites on the Kv1.2 potassium channel S1-S2 linker: implications for linker secondary structure and the glycosylation effect on channel function. Biochem J 375:769-775.  101  Chapter 4: The molecular basis for the actions of Kvβ1.2 on the opening and closing of the Kv1.2 delayed rectifier channel1  1  A version of this chapter has been published. Peters, C.J., Vaid, M., Horne, A.J., Fedida, D., & Accili, E.A. (2009). The molecular basis for the actions of Kvβ1.2 on the opening and closing of the Kv1.2 delayed rectifier channel. Channels 3(5), 1-9.  102  Introduction  In native tissue, the Shaker-related (Kv1) family of channels exists as pore-forming α−subunits in heteromeric complexes with auxiliary cytosolic proteins termed Kvβ-subunits (Parcej and Dolly, 1989; Scott et al., 1990; Parcej et al., 1992; Rhodes et al., 1995; Rhodes et al., 1996; Rhodes et al., 1997; Shamotienko et al., 1997; Orlova et al., 2003). According to recent crystallographic studies, these are positioned below the associated subunits in 1:1 stoichiometry with four-fold symmetry and form a so-called ‘hanging gondola’ (Gulbis et al., 2000; Long et al., 2005). To date, three mammalian Kvβ genes (Kvβ1-3) have been cloned (Rettig et al., 1994; Majumder et al., 1995; Heinemann et al., 1995; England et al., 1995a; England et al., 1995b; Heinemann et al., 1996), as well as a Kvβ homolog from Drosophila melanogaster called Hyperkinetic (Chouinard et al., 1995). The C-terminus is highly conserved among the three Kvβ genes and co-translationally forms the primary contacts with the T1 domain of the N-terminus of Kvα1 subunits (Sewing et al., 1996; Nakahira et al., 1996; Shi et al., 1996; Yu et al., 1996; Wang et al., 1996; Nagaya and Papazian, 1997; Gulbis et al., 2000). In contrast, the N-terminus is highly variable in primary structure among the three Kvβ genes, including three splice variants of Kvβ1, and does not form any long-lasting interactions with the Kv1 T1 domain (Accili et al., 1997). Elegant structural studies have shown that the N-termini of Kvβ1 subunits can reach the central cavity and inner pore of an N-terminally truncated Kv1.4 channel as an extended peptide to confer rapid channel block (Zhou et al., 2001), and that occupancy of this region is limited to one N-terminus at any given time. Rapid open channel block of Kv1.2 by the Kvβ1.2 Nterminus reduces whole-cell currents by up to ~90%, which can be explained completely by a corresponding reduction in open probability (Accili et al., 1997).  103  Functional interactions between Kvβ1 and Kvα1 subunits are complex. All three Kvβ1 subunits convert Kv1 channels from the delayed rectifier phenotype observed with the αsubunits alone to a rapidly decaying transient outward current, and greatly slow channel closing upon repolarization (Rettig et al., 1994; Castellino et al., 1995; Accili et al., 1997). Fast inactivation and slowed channel closing can be abolished by the removal of the N-terminus, and the remaining Kvβ1 C-termini differentially regulate the surface trafficking of Kv1 channels, with a sizable upregulation of current and cell surface expression conferred upon Kv1.2 by Kvβ1 subunits (Shi et al., 1996; Accili et al., 1997; Accili et al., 1998). A negative shift in the activation curve is also observed when Kvβ1 subunits are co-expressed with Kv1 channels, but the structural mechanism underlying this effect is equivocal. Ablation of the unique N-terminus eliminates the negative shift produced by Kvβ1.2 on Kv1.2 (Accili et al., 1997) and mutations in Kvβ1.3 that remove fast inactivation make the shift in Kv1.5 activation less pronounced (Decher et al., 2008). Modeling of ionic current data suggests that open channel block by the N-terminus, in addition to causing fast inactivation, can produce saturation of activation at depolarized potentials and thus an apparent negative shift in activation (DeBiasi et al., 1997). Slowed channel deactivation is consistent with the inability of the channel gate to close while the Kvβ1 N-terminus resides in the pore, as has been demonstrated for fast inactivation produced by the Nterminus of Kv1.4 and Shaker channels, except that the extent of slowing induced by the Kvβ1 N-terminus is far greater. Allosteric interactions between Kvβ subunits and Kv1 channels may contribute to the shift in channel activation (Heinemann et al., 1996; Uebele et al., 1998) or to the slowing of channel deactivation, for example, by acting directly on the S4 segment or activation gate. Conformational changes in the outer pore that are associated with slow inactivation and 104  augmented by the Kvβ1 N-terminus (Morales et al., 1996; Accili et al., 1998) could also inhibit the closure of the activation gate as well as the return of the movement of the S4 helix to its resting position. However, there is no structural or functional evidence, as of yet, identifying any direct or indirect interaction of Kvβ with the voltage sensing elements. Do Kvβ1 subunits modify the movement of the S4 segment? If so, how is S4 movement modified and what parts of the Kvβ subunit are responsible? To answer these questions and establish the mechanisms underlying the effects conferred by the Kvβ1.2 subunit on Kv1.2 channel activation and deactivation, movement of the S4 segment and ion flow were independently tracked by combining voltage clamp fluorimetry and current recording. Voltage clamp fluorimetry permits the visualization in real time of protein conformational changes in response to stimuli, while simultaneously recording ionic currents with two electrode oocyte voltage clamp (Mannuzzu et al., 1996; Cha and Bezanilla, 1997). Our data demonstrate that distinct effects on Kv1.2 channel opening and closing, and on S4 movement, result from interactions with the N-terminus of Kvβ1.2.  105  Materials and methods  Molecular biology Human Kvβ1.2 was cloned from pcDNA3 into the vector pBluescript SK+ for expression in Xenopus laevis oocytes by digesting insert and vector with restriction enzymes XbaI and EcoRI (from New England Biolabs, Ipswich, MA). An N-terminal deletion mutant of Kvβ1.2 missing residues 1-77 (Kvβ1.2ΔN) was generated by simultaneous PCR amplification of the Kvβ1.2 C-terminus and introduction of an EcoRI recognition sequence at its 5’ end, after which the PCR product was digested with XbaI and EcoRI and re-inserted into pBluescript SK+. The modified pBluescript vector pEXO was used to express rat Kv1.2 in oocytes. Cysteine residues were introduced at specific sites (M288 and A291) in the S3–S4 linker for fluorophore labeling. Point mutations were introduced using Quikchange (Stratagene, Cedar Creek, TX) with primers synthesized by Integrated DNA Technologies (Coralville, IA), and were sequenced at the University of British Columbia core facility.  cRNA was synthesized using the mMessage  mMachine T7 Ultra transcription kit (Ambion, Streetsville, ON) from cDNA templates linearized with SacII (Kv1.2) or NotI (Kvβ1.2 and Kvβ1.2ΔN). For co-injections of α and β subunit RNA, β-subunit RNA was pre-mixed with α-subunit RNA at a 50:1 ratio, to maximize the number of co-assembled channels, as was characterized previously (Accili et al., 1997).  Oocyte preparation Xenopus laevis oocytes were prepared and isolated as has been described previously (Claydon et al., 2007). Following removal of the follicular layer, oocytes were injected with 50 nl (10–200 ng) of cRNA and incubated in Barth's solution, which contained (in mM) 88 NaCl, 1 KCl, 2.4 NaHCO3, 0.82 MgSO4, 0.33 Ca(NO3)2, 0.41 CaCl2, 20 HEPES (pH 7.5), for 1–3 days at 106  19°C. Prior to recording, injected oocytes were labelled with a reactive fluorescent dye, 5 µM tetramethylrhodamine-5-maleimide (TMRM), in a depolarizing solution containing (in mM) 99 KCl, 1 MgCl2, 2 CaCl2, and 5 HEPES (pH 7.5), for 30 min at 10°C.  Two-electrode voltage clamp electrophysiology and fluorimetry Ionic currents and fluorescence signals were recorded simultaneously using two-electrode voltage clamp fluorimetry as described previously (Claydon et al., 2007). The bath solution contained (in mM) 96 NaCl, 3 KCl, 1 MgCl2, 2 CaCl2, and 5 HEPES (pH 7.5). Voltagedependent fluorescence changes were measured from TMRM bound via a stable carbon-sulfur bond to cysteine residues introduced in the S3–S4 linker. Excitation and emission light were filtered with 525-nm bandpass and 560-nm longpass filters, respectively. Emitted light was detected using a 9124b Electron Tubes photomultiplier tube (ET Enterprises, Uxbridge, UK). Acquired signals (ionic current and fluorescence) were sampled at 20 kHz and low-pass filtered off-line at 1-3 kHz. Fluorescence traces recorded from TMRM represent the average of at least five sweeps. Recording microelectrodes contained 3M KCl and had resistances between 0.2 and 0.8 MΩ.  Data analysis Conductance–voltage (G-V) and fluorescence–voltage (F-V) relations were fitted with a single Boltzmann function, y=  1 ⎡V − V ⎤ 1 + exp ⎢ 1 / 2 ⎥ ⎣ k ⎦  107  where y is the conductance or fluorescence amplitude normalized with respect to maximal conductance or fluorescence amplitude, V1/2 is the half-activation potential, V is the test voltage, and k is the slope factor. Data are shown as mean ± SEM.  Statistical differences were  determined using the Student’s t-test. Where mean values for fits (Boltzmann and exponential functions) are quoted, they represent the averages of fits to data from individual oocytes in order to obtain values for SEM.  108  Results  The Kvβ1.2 N-terminus induces fast inactivation, a negative shift of activation and slowed deactivation in Kv1.2 In order to establish the effects of the Kvβ1.2 subunit on Kv1.2 in our system, oocytes were injected with cRNA encoding Kv1.2 alone, or mixed with either full length Kvβ1.2, or a mutant of Kvβ1.2 lacking the first 77 residues, called Kvβ1.2ΔN (Figure 4.1A). The sequence of Kvβ1.2ΔN corresponds to a C-terminal region that is identical among the three Kvβ1 splice variants and thus is referred to as Kvβ1 C-terminus in previous studies from our group (e.g. Accili et al, 1997). The cysteine-substituted Kv1.2 mutants were utilized for these ionic studies as well as for the fluorescence-tracking experiments which were performed in parallel. Typical delayed rectifier currents were recorded from Kv1.2-expressing oocytes and showed little to no inactivation over the 100 ms pulse duration (Figure 4.1B). In the presence of Kvβ1.2, but not the truncated Kvβ1.2ΔN, ionic currents from Kv1.2 measured at depolarized potentials were converted from delayed outward rectifying currents to transient outward currents followed by a rapid decay to a steady state value, reminiscent of N-type inactivation (Figure 4.1B). Tail current amplitudes, measured immediately following the pulse to -40 mV, were normalized to the maximum amplitude, plotted against test voltage and fitted to a Boltzmann function (Figure 4.1C). We noted a leftward shift of the activation curve, along with markedly slower kinetics in the ionic tail currents, when Kvβ1.2 was co-injected with Kv1.2 (Figure 4.1D). When Kvβ1.2ΔN was used in place of the full length Kvβ1.2, the recorded ionic currents were kinetically indistinguishable from those recorded from Kv1.2 injected alone, though they were significantly larger in amplitude (data not shown), consistent with previous observations of Kv1.2 current upregulation by Kvβ1 and Kvβ2 in Xenopus laevis oocytes (Accili et al., 1997;  109  Accili et al., 1998). Upregulation of current has been used as functional evidence for a direct interaction, which has been demonstrated in a number of studies utilizing biochemical assays (see Introduction). These data show that the fast inactivation, activation shift, and slowed deactivation observed when Kv1.2 was co-expressed with Kvβ1.2 were due to its N-terminus, as was shown previously (Accili et al., 1997). Moreover, the similarity of these data with those obtained from wild type Kv1.2 channels show that the cysteine substitutions do not alter the interaction between these subunits.  110  Figure 4.1. Kvβ1.2 confers a spike-and-decay “fast inactivation” to Kv1.2 currents, a hyperpolarizing shift of the activation curve, and a slowing of current deactivation. (A) The Kvβ1.2 constructs used in this study. Kvβ1.2ΔN lacks the first 77 amino acids that contain the region responsible for fast inactivation. The primary sequence of the deleted amino acids of Kvβ1.2 is shown in the right panel. (B) Current traces recorded from oocytes expressing Kv1.2 A291C alone, with Kvβ1.2 or with Kvβ1.2ΔN. Traces were recorded in response to depolarizing test pulses from a holding potential of -80 mV to voltages ranging from -120 mV to +60 mV in 10 mV steps, then to -40 mV to elicit outward tail currents, before being returned to the holding potential. (C) A plot of normalized tail current amplitudes versus test voltage, fitted with a single order Boltzmann function. This relation (the G-V curve) is steepened and shifted to more negative potentials significantly (p<0.05, Student’s t-test) when Kv1.2 A291C is co-injected with Kvβ1.2 (circles, dashed line; V1/2 = -17.50 ± 1.69 mV, k = 13.30 ± 0.61 mV, n = 16) as compared with Kv1.2 A291C expressed alone (squares, solid line; V1/2 = -5.78 ± 2.78 mV, k = 15.15 ± 0.64 mV, n = 23), or with Kvβ1.2ΔN (triangles, dotted line; V1/2 = -8.30 ± 3.26 mV, k = 14.92 ± 0.44 mV, n = 9). (D) A plot of deactivation time constants determined from single exponential fits of tail currents recorded at -40 mV following test pulses from -20 mV to +40 mV, in 10 mV increments. The values for Kv1.2 A291C + Kvβ1.2 (circles) are considerably larger at all potentials (p<0.0001 at all potentials tested, Student’s t-test, n values as in C) compared to for Kv1.2 + Kvβ1.2ΔN (triangles) or Kv1.2 A291C alone (squares).  111  The voltage dependences of fluorescence deflections upon depolarization from TMRM-labelled Kv1.2 S4 mutants are unaffected by Kvβ1.2 To track the movement of the S4 segment independently from ionic current, two sites at the extracellular end of the S4 helix were labelled with TMRM dye, and voltage clamp fluorimetry recording was performed alongside two-electrode voltage clamp. As mentioned in the Experimental Procedures, cysteine residues were introduced at sites M288 and A291, as the equivalent residues (M356 and A359; Figure 4.2A) were found to faithfully track S4 movement in Shaker (Mannuzzu et al., 1996). The addition of TMRM to wild type channels produced little fluorescence quenching in response to depolarizing test pulses (Figure 4.2B), despite the presence of an external cysteine residue (C181) in wild-type Kv1.2. We generated a mutant Kv1.2 C181V A291C, and found that fluorescence signals therefrom were kinetically equivalent to Kv1.2 A291 with C181 intact (see overlapping traces in Figure 4.2C). However, channel surface expression was reduced for the C181V A291C mutant. Therefore, C181 was left intact for these studies.  112  Figure 4.2. Kv1.2 current and/or fluorescence kinetics are not modified by substitutions required to record fluorescence. (A) An alignment of the S3-S4 linkers of Shaker and rat Kv1.2 potassium channel α-subunits, with expected topology. Unlike the long linker present in Shaker, Kv1.2 has only a short external S3-S4 loop. Residues mutated to cysteine for voltage clamp fluorimetry, M288 and A291, are indicated by arrows. (B) (Top) Current traces recorded from oocytes expressing wild-type Kv1.2 labelled with TMRM in response to depolarizing test pulses to voltages ranging from –120 mV to +60 mV in 10 mV steps, from a holding potential of -80 mV. (Bottom) Fluorescence trace recorded at +60 mV in the above protocol (see the corresponding current trace above indicated in black). No discernible changes in fluorescence signals were observed over 100 ms at +60 mV (or any other potential tested). (C) Normalized fluorescence recordings from oocytes expressing Kv1.2 C181V A291C or Kv1.2 A291C during 100 ms pulses to +60 mV, which show little difference in activation and deactivation kinetics. The increased noise in Kv1.2 C181V A291C is due to the lower level of surface expression and the larger amplification of the signal with normalization.  TMRM-labelled Kv1.2 M288C and Kv1.2 A291C yielded voltage-dependent fluorescence deflections in a voltage-clamp fluorimetry configuration (Figure 4.3A). Here, oocytes were subjected to a protocol similar to the one used to generate G-V curves in Figure 4.1, except that after depolarization, oocytes were returned to -100 mV to allow complete channel deactivation and return of the voltage sensor. As seen in traces from both M288C and A291C, robust fluorescence that varied as a function of applied voltage could be recorded under these conditions. In order to examine effects of the Kvβ1.2 N-terminus on voltage sensor movement 113  and on channel activation simultaneously and separately, we co-injected cRNA encoding Kvβ1.2 (or Kvβ1.2ΔN) with Kv1.2 M288C or A291C, and recorded fluorescence deflections. To compare the voltage dependencies of the fluorescence deflections associated with voltage sensor movement, signal amplitudes from Kv1.2 A291C and M288C were normalized to those at 0 mV, plotted against membrane potential and fit with a Boltzmann function to generate fluorescence-voltage (F-V) curves, from which values of mid-activation voltage and slope factor were determined.  For Kv1.2 A291C and Kv1.2 M288C, no differences in the voltage  dependence of fluorescence were observed when Kvβ1.2, or Kvβ1.2ΔN, were co-expressed with the α-subunit as compared to the α-subunit injected alone (Figure 4.3B,C). In all cases, the F-V curves span a range of voltages more negative than those of the G-V curves (the fitted G-V curve for Kv1.2 from Figure 4.1B is shown in grey in Figure 4.4), as would be expected for a signal correlated to voltage dependent S4 movement that precedes channel opening. Negatively shifted S4 fluorescence and gating charge as functions of test voltage have been demonstrated for the closely related Shaker channels (Bezanilla et al., 1991; Mannuzzu et al., 1996; Cha and Bezanilla, 1997) and Kv1.5 channels (Chen et al., 1997; Vaid et al., 2008). Interestingly, the F-V curve for A291C is markedly left-shifted and less steep than that for M288C as well as those published with the equivalent construct (A359C) in Shaker (Bezanilla et al., 1991; Mannuzzu et al., 1996), suggesting A291C may be additionally reporting on a separate aspect of gating from Kv1.2 M288C.  114  Figure 4.3. Voltage-dependence of S4 movement upon depolarization in Kv1.2 is unaffected by Kvβ1.2. (A) Fluorescence traces at indicated voltages for 100 ms, from a holding potential of -80 mV, recorded from oocytes expressing Kv1.2 M288C (above) or Kv1.2 A291C (below) alone (left), with Kvβ1.2 (middle), or with Kvβ1.2ΔN (right). Oocytes were subjected to a series of depolarizing potentials in 10 mV steps, then returned to -80 mV, while simultaneously recording fluorescence deflections. Some traces are removed for clarity. For A291C, this protocol followed a brief -160 mV prepulse to yield directionally consistent fluorescence. (B) (left) Plots of fluorescence signals, normalized to fluorescence deflection amplitudes at 0 mV (ΔF0 mV), recorded from oocytes expressing Kv1.2 M288C versus test voltage, and fitted with a single order Boltzmann function. This relation (the F-V curve) was unaffected (p>0.05, Student’s t-test) by the full length Kvβ1.2 (circles, dashed line; V1/2 = -42.12 ± 0.77 mV, n = 7) or Kvβ1.2ΔN (triangles, dotted line; V1/2 = -45.73 ± 0.56 mV, n = 9), compared to Kv1.2 M288C alone (squares, solid line; V1/2 = -46.75 ± 1.35 mV, n = 8). (right) F-V curves generated of Kv1.2 A291C alone (squares, solid line; V1/2 = -60.43 ± 0.62 mV, n = 14), are unaffected (p>0.05) by full length Kvβ1.2 (circles, dashed line; V1/2 = -62.15 ± 0.92 mV, n = 8) and Kvβ1.2ΔN (triangles, dotted line; V1/2 = -63.72 ± 0.26 mV, n = 11).  115  Slowing of both current deactivation, and fluorescence return to baseline levels upon channel closing, are induced by the Kvβ1.2 N-terminus and correlated to the proportion of channels undergoing fast inactivation Notably, the fluorescence tails were considerably slowed by full length Kvβ1.2, resulting in the imposition of a second, slower phase of fluorescence at some potentials (Figure 4.3A, 4.4A). These observations are consistent with a slowed second phase of fluorescence in the full length Shaker channel (Savalli et al., 2007). To examine the voltage dependence of the onset of slowed fluorescence tails, the fluorescence tails were fitted with a double exponential function, and amplitudes for each component were determined. We reasoned that the slower component was due to the pore-localized Kvβ1.2 N-terminus and that the amplitude of this component reflects the proportion of available channels that had entered an inactivated state. When the ratios of the slower τ amplitudes to the total fluorescence tail amplitudes were normalized, and plotted against voltages of the preceding pulses (Figure 4.4B), they overlaid well with the G-V curve of Kv1.2 (from Figure 4.1C). This suggests that slowing depends on the fraction of open channels that are available for fast block by the Kvβ1.2 N-terminus or, alternatively, for a slower inactivation process.  116  Figure 4.4. The voltage dependence of the fraction of the fluorescence that is slowed upon repolarization is correlated with the Kv1.2 G-V curve. (A) Deactivation fluorescence tails from M288C with and without Kvβ1.2 at -80 mV following a test pulse to 0 mV are normalized to their respective minima and maxima, and fitted with a single (M288C alone) or a double (with Kvβ1.2) exponential function (overlaid in grey). (B) Plot of amplitudes of the slow τ from Kv1.2 M288C + Kvβ1.2 fluorescence deactivation tails (from Figure 4.3A), as a fraction of the total amplitude (fast and slow) and normalized to the maximum fractional value, versus test voltage (circles, n = 5). These are overlaid with a Kv1.2 G-V curve (squares, from Figure 4.1C).  If the slower component were due to the pore-localized Kvβ1.2 N-terminus and fast block, then the channels entering the fast inactivated state should leave this state slowly and the proportion of channels inactivated would depend upon how quickly they entered this state. Therefore, depolarizing pulses of varying duration were delivered and the amplitudes of slow components of the tails as a function of time upon relaxation to negative potentials were determined. Samples of fluorescence tails at -80 mV from Kv1.2 alone, or with Kvβ1.2 or Kvβ1.2ΔN following depolarizing pulses to +60 mV for 5 ms-40 ms are shown in Figure 4.5A. As before, fluorescence tails recorded with Kvβ1.2 were fitted with a double exponential  117  function from which amplitudes for the slow component were determined. Using a similar protocol, the amplitudes for the slow component were determined at -40 mV for ionic tail currents from Kv1.2 with Kvβ1.2 (Figure 4.5B). These amplitudes, which likely represent the proportion of channels in the inactivated state, were plotted against prepulse duration (Figure 4.5C). Although the maximum value was larger for ionic current, single exponential fits of these data yielded almost identical values of τ of 8.5 ± 1.3 ms (n = 6) for fluorescence and 8.8 ± 1.2 ms (n = 8) for ionic currents. These values likely reflect the progressive entry of channels into the fast inactivated state because they are similar to the order of onset of fast current inactivation at +60 mV (6.0 ± 0.8 ms, n = 11), a sample of which is overlaid in grey in Figure 4.5C.  118  Figure 4.5. Current deactivation and the return of fluorescence to baseline are coupled and slowed by the Kvβ1.2 N-terminus. (A) Fluorescence traces from oocytes expressing Kv1.2 M288C alone, or with Kvβ1.2 or Kvβ1.2ΔN, recorded at -80 mV following prepulses to +60 mV for variable durations between 5-40 ms (5, 20 and 40 ms traces are shown). Traces were fitted with a single (M288C alone or with Kvβ1.2ΔN) or a double (with Kvβ1.2) exponential function, which are overlaid in grey. (B) Current traces elicited from oocytes expressing Kv1.2 M288C with Kvβ1.2 as for fluorescence traces in A except at -40 mV to visualize them in the outward direction. (C) Plots of amplitudes of the slow τ from Kv1.2 M288C + Kvβ1.2 fluorescence (squares, n = 6) and current (circles, n = 8) deactivation tails (from Figure 4.5A,B), as a fraction of the total amplitude (fast and slow) versus prepulse duration. Each set of values is fitted with a single exponential function (solid, fluorescence; dashed, ionic). To better compare these fitted curves, the fit for the fluorescence data was normalized to the maximum value for the ionic current fit and overlaid in grey.  119  Discussion  Our data demonstrate that distinct effects on Kv1.2 channel opening and closing, and on S4 movement, result from transient interactions with the N-terminus of Kvβ1.2. The voltage dependence of S4 movement was unaffected by that subunit despite the apparent shift of the G-V curve to more negative voltages. This apparent shift is thus likely due to the progressive saturation of ionic tail currents from which channel activation is determined at depolarized membrane potentials; this is consistent with a model of open channel block to describe interactions of the Kvβ1 N-terminus with Kv1 channels (DeBiasi et al., 1997; Accili et al., 1997), and rules out a direct effect of the Kvβ1 C-terminus on the channel voltage sensor. Along with a corresponding slowing of current deactivation, the fluorescence deflections recorded during deactivation of Kv1.2 exhibited a dramatic slowing by Kvβ1.2. This slowing correlated with the proportion of open channels entering the fast inactivated state, on a time scale too quick for a slower inactivation process. These findings compare closely with those obtained using Shaker channels, which show that gating charge is “immobilized” during the intrinsic fast inactivation invoked by its own N-terminus; this occludes the pore and prevents gate closure (Bezanilla et al., 1991). Immobilization likely results from physical associations between the activation gate and the S4 segments. Although it is possible that the Kvβ1.2 N-terminus may have inhibited fluorescence return by acting directly on the S4 segment, there are three reasons to suggest that this does not occur. First, the voltage-dependence of on-fluorescence movement is unaffected by Kvβ1.2. Second, the observation that the slowing of ionic and fluorescence tails during deactivation are similar to the rate of onset of inactivation can be explained most easily by open channel block by the N-terminus. Third, previous structural studies have shown that the Nterminus interacts predominantly with the pore and not with the S4 segment (Zhou et al., 2001), 120  although it may be that such a conformation cannot be resolved by current structural methods. Therefore, although an interaction between the Kvβ1.2 N-terminus and the S4 segment is possible, it does not seem likely based on the evidence to date. When Kvβ1.2 was co-expressed with Kv1.2, the amplitude of the slow time constant for the recovery rate of fluorescence following a depolarizing pulse of increasing length made up a smaller proportion of the total amplitude compared to that for the ionic traces: even after the longer prepulses, a significant fraction of the fluorescence recovery remained fast. This implies that the fluorescence signal is reporting on an additional or separate component of movement of the S4 segment when the channel closes that is not slowed by the Kvβ1 N-terminus. This is consistent with studies showing that the return of the S4 segment may be partitioned into several steps, some of which may not necessarily correspond temporally with activation gate closure (Wang and Fedida, 2002). Alternatively, the Kvβ1 N-terminus may not inhibit the returning movement of all four S4 segments, similar to what has been reported for rapid inactivation in the voltage-gated sodium channel Nav1.4, where occlusion of the pore (albeit by a different mechanism) preferentially slows the return of only two of four S4 segments to their resting position during channel closure (Cha et al., 1999). Although the influence on Kv1.2 gating behaviour can be explained by open channel block by the Kvβ1.2 N-terminus, the blocking process itself may be more complex and consist of at least two separate stages (Zhou et al., 2001). The binding of the N-terminus to the cytoplasmic surface may occur initially, placing the channel in a pre-inactivated state. This initial interaction would be followed by the movement of the N-terminus into the pore as an extended peptide, such that the channel becomes blocked and fully-inactivated. The slowed return of fluorescence and current deactivation in our studies of Kv1.2 could result from an interaction of the Kvβ1.2 121  N-terminus with either the cytoplasmic surface or the inner pore but, because slowing is voltage independent, the binding of the extended peptide to the cytoplasmic face, an interaction on the outside of the membrane electric field, may make the most sense intuitively.  A two-step  unblocking process also fits with our data showing that a fast fluorescence component remains, which may correspond to a fast exit of the Kvβ1.2 N-terminus from the inner pore. While our results are in strong accordance with the findings of Zhou et al. using Kv1.4 and Kvβ1.1, the variability in primary structure within the N-termini among different Kvβ1 subunits and the involvement of other cellular elements may contribute to the complexity of the block by altering Kvβ interaction at both the outer and inner pore mouth, with potentially different consequent effects on voltage sensor movement or its coupling to channel opening. For example, the Kvβ1.3 N-terminus, unlike Kvβ1.1, may enter the pore as a hairpin and the inactivation conferred by this subunit may be modified by the cellular metabolite PIP2 (Decher et al., 2008). So while some similarity in mechanism seems evident from our data and those of others, there are undoubtedly significant differences, which makes comparative studies among the Kvβ1 splice variants important in the future. One ongoing challenge in constructing a cohesive picture of Kvβ subunit behaviour, and understanding the overall consequences of Kv α- and β-subunit associations, is the difficulty in performing unambiguous functional studies in native tissue. Indeed, the three Kvβ proteins and their splice variants, which lack any measurable electrical behaviour of their own, are known to interact promiscuously with known Kv1α subunit isoforms to alter trafficking and gating, as well as to enhance their sensitivity to redox environment (Wang et al., 1996; Bahring et al., 2001; Weng et al., 2006; Pan et al., 2008). Model systems, such as the oocyte expression technique used in this study, provide critical insights into possible behavioural roles by driving interactions 122  mainly between Kvβ and Kvα proteins in a cellular context. These findings can be used to generate hypotheses about their function in native tissue but studies such as global Kvβ1 knockouts in mice have yielded, not surprisingly, data that are consistent with these subunits having a role more complex than has been observed in heterologous cells (Giese et al., 1998; Murphy et al., 2004; Aimond et al., 2005). Interactions among Kvβ subunits (Accili et al., 1997; Gulbis et al., 1999) as well as between Kvβ subunits and Kv channels from other subfamilies e.g. Kv4.2 and 4.3 (Nakahira et al., 1996; Rhodes et al., 1997; Perez-Garcia et al., 1999; Yang et al., 2001; Aimond et al., 2005) further increase the potential for pleiotropism and require further analysis in both heterologous expression systems and native tissue. Approaches to better define the molecular compositions of channels, and conditional knockout experiments that target specific tissues and stages of development, and thus limit compensation of any changes induced by the deletion of genes, will also greatly aid in the understanding of the role for Kvβ subunits, and their functional interactions with Kv1 channels, in native tissue.  123  Acknowledgements  The human Kvβ1.2 subunit was a kind gift from Dr. B. Wible (Case Western Reserve University, Cleveland, OH). The modified pBluescript vector pEXO was a kind gift from Dr. A. Sivaprasadarao, (University of Leeds, UK). We would like to thank F. Chiu for assistance in construct cloning, and T. Claydon for feedback on initial experiments. C.J.P. received support from a Graduate Fellowship from the Michael Smith Foundation for Health Research, A.J.H. is supported by a NSERC Post-Graduate Scholarship, D.F. is supported by a Career Investigator award from the Heart and Stroke Foundation of British Columbia and Yukon, E.A.A. is supported by the Heart and Stroke Foundation of British Columbia and Yukon, and is also the recipient of a Tier II Canada Research Chair.  124  References  Accili, E. A., J. Kiehn, Q. Yang, Z. G. Wang, A. M. Brown, and B. A. Wible. 1997. Separable Kvβ subunit domains alter expression and gating of potassium channels. J Biol Chem 272:25824-25831. Accili, E. A., Y. A. Kuryshev, B. A. Wible, and A. M. Brown. 1998. Separable effects of human Kvβ1.2 N- and C-termini on inactivation and expression of human Kv1.4. J Physiol 512 (Pt 2):325-336. Aimond, F., S. P. Kwak, K. J. Rhodes, and J. M. Nerbonne. 2005. 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NAB domain is essential for the subunit assembly of both α-α and α-β complexes of Shaker-like potassium channels. Neuron 16:441-453. Zhou, M., J. H. Morais-Cabral, S. Mann, and R. MacKinnon. 2001. Potassium channel receptor site for the inactivation gate and quaternary amine inhibitors. Nature 411:657-661.  128  Chapter 5: Discussion  129  Kv1.2 is expressed in excitable cells throughout the body  Functional studies have shown abundant Kv1.2 expression in the brain and nervous system of multiple species (Stuhmer et al., 1989; Sheng et al., 1993; Wang et al., 1993; Sheng et al., 1994; Wang et al., 1994; Sobko et al., 1998; Glazebrook et al., 2002; Gutman et al., 2003), including humans (Coleman et al., 1999). In KCNA2 (Kv1.2) knockout mice, the average lifespan was reduced to 17 days due mainly to an increased susceptibility to spontaneous and induced seizures (Brew et al., 2007). Interestingly, hypoexcitability was observed in auditory neurons lacking Kv1.2, and was attributed to the loss of subunit contribution within heterologous channels leading to altered neuron kinetics, implicating the importance of this subunit as both a homomer and heteromultimer. Kv1.2 has also been observed in airway smooth muscle (Adda et al., 1996), and is well demonstrated in the cardiovascular system (Paulmichl et al., 1991; Roberds and Tamkun, 1991; Barry et al., 1995; Bertaso et al., 2003). The prevalence of Kv1.2 throughout physiological systems underscores its importance in the repolarization of excitable cells, and as such it is of great interest to develop a detailed understanding of its voltagedependent gating.  Scientific rationale for investigating Kv1.2 gating  Despite the incredible amount of insight afforded to us by the crystallization of Kv1.2 (Long et al., 2005), and later the Kv1.2/Kv2.1 “paddle” (S3b-S4) chimera (Long et al., 2007), many questions remain unanswered with regard to gating dynamics of Kv1.2 and potassium channels in general. The structure as we know it represents a single moment in time for this otherwise dynamic protein, with minimal information about how it is gated to or from its captured state.  Furthermore, crystallization was carried out in the absence of a potential  130  difference (0 mV), making it unclear exactly what channel state was obtained.  Both the  activation gate and voltage sensor appear to be displaced from their closed state positions predicted by cysteine accessibility studies (Larsson et al., 1996; Yusaf et al., 1996; Baker et al., 1998) and histidine proton current (Starace and Bezanilla, 2001; Starace and Bezanilla, 2004) data, but a moderate shift in the position of S4 is necessary to fit with other experimental data obtained from open-state histidine cross-linking studies (Lewis et al., 2008), taking into account the distance over which such interactions are formed. Thus, it is uncertain whether the crystal structure portrays the channel in the physiological open state or in further modified conformations (physiological or not) as a result of prolonged depolarization (Bezanilla, 2008). Alternatively, different parts of the channel may have been captured in different states. While there are still uncertainties with regards to the Kv1.2 crystal structure, there is no doubt that it represents a great leap forward in the quest to understand potassium channel gating, and provides a measure of insight lacking in many other voltage-gated proteins. In the absence of crystal structures for Shaker or other Kv channels, functional data obtained from these channels is often interpreted using the Kv1.2 channel crystal structures (Long et al., 2005; Long et al., 2007). Electrophysiologically, Kv1.2 has not been studied as extensively as Shaker and many other Kv channels, so our knowledge of its functional properties is somewhat limited. However, the studies that have been conducted show significant and important differences in the gating, structure and function of Kv1.2 that suggest such comparisons should be made with caution. Kv1.2 does not appear to undergo significant inactivation of ionic current, even with long depolarizations (Paulmichl et al., 1991), suggesting that known interactions between S4 and the pore domain (Loots and Isacoff, 2000; Elinder et al., 2001) may not occur in this channel. Glycosylation effects (Zhu et al., 2003; Watanabe et al., 2007; Zhu et al., 2009) and pH  131  sensitivity (Steidl and Yool, 1999; Ishii et al., 2001) differ significantly between Kv1.2 and other channels, even closely related channels such as Shaker. As well, in mammalian cells Kv1.2 has been shown to exhibit a bimodal gating phenotype with widely disparate half-activation potentials and activation time constants, that has not been observed in Shaker or Kv1.5 channels (Rezazadeh et al., 2007). The purpose of the present series of experiments was to expand our knowledge of Kv1.2 gating and structure using electrophysiological and fluorimetric methods, to develop a more detailed understanding of protein dynamics that can then hopefully be applied to information gleaned from the crystal structure.  In addition, understanding similarities and differences  between Kv1.2 and other potassium channels will go a long way towards assessing the suitability of using this channel in referencing and interpreting data from other channels, and of Kv1.2 as a universal template of Kv structure in general.  Kv1.2 gating currents correlate with rapid fluorescence quenching  When labelled with TMRM at an introduced cysteine at the extracellular end of S4 (A291C), the probe exhibits a rapid change in fluorescence emission, followed by a very prominent slow quenching component (Figure 2.2).  This is in contrast to Shaker, where  labelling of A359C lacks such a prominent and negatively left-shifted slow component. When the rapid Kv1.2 component is plotted as a function of membrane potential, it exhibits a voltagedependence very similar to Shaker A359C. This fast component in Shaker has been wellcorrelated with gating currents (Mannuzzu et al., 1996; Cha and Bezanilla, 1997), but no such records have ever been obtained for Kv1.2 channels. Here, in the first report of Kv1.2 gating currents, we show that the voltage-dependence and time course of S4 movement correlate  132  extremely well with that of the rapid fluorescence quenching component from TMRM-A291C (Figure 2.6). Furthermore, the Kv1.2 Q-V relationship is similar to previous reports for Shaker gating currents (Perozo et al., 1994; Claydon et al., 2007b). Therefore, it appears that both channels have similar kinetics associated with the major element of S4 translocation, as reported by both gating current and fluorescence measurements. However, there is an additional facet to Kv1.2 gating uniquely uncovered by fluorimetry, suggesting additional slow environmental changes in this region that will be discussed below.  Slowed fluorescence upon repolarization is consistent with delayed S4 return  In addition to the rapid Shaker-like report of channel gating currents, Kv1.2 A291C fluorescence reports a prominent slow quenching component, with a very negative voltagedependence (V1/2 = -73.9 ± 1.4 mV) and a time constant of ~24 ms at all potentials tested. Importantly, there appears to be no measurable gating charge correlate to this movement, which may be the primary reason such a movement within the Kv1.2 voltage sensor domain has never before been reported. We have shown that this slow component, while not obviously affecting the kinetics of activation, decays with a time course overlapping that of ionic current deactivation upon repolarization (Figure 2.7), suggesting that the movement may need to reverse before channel closure can occur. Fluorophores at locations within the S1-S2 linker uniquely report this slow quenching, in the absence of rapid S4-associated changes in emission (Figure 2.10). In chimera channels replacing the Kv1.2 S1-S2 linker, the S3-S4 linker, or both linkers with those from other channels (Kv1.5 or Shaker) this slow component was lacking, and these constructs exhibited faster deactivation and off-fluorescence kinetics compared to Kv1.2 A291C, suggesting a complex interplay between these protein moieties throughout the voltage sensing domain.  133  To further investigate the properties of these fluorescence components, we looked for changes in both on- and off-fluorescence in the presence of an N-type inactivation-conferring Kvβ1.2 subunit. We found that on-fluorescence from A291C and M288C in S4 was unaltered in the presence of the auxiliary subunit (Figure 4.3); rather, the leftward shift of the Kv1.2 G-V curve appears to result from increased channel block at greater potentials, artificially shifting the conductance. The fluorescence data, combined with the absence of this effect with an Nterminal deleted Kvβ1.2 subunit support previous studies suggesting Kvβ1.2 acts through a poreblocking mechanism, independent of the voltage sensing domain (DeBiasi et al., 1997; Accili et al., 1997). However, off-fluorescence was significantly slowed in the presence of Kvβ1.2, but not in the deletion mutant (Figure 4.4, 4.5), suggesting that the same interaction of the Kvβ1.2 N-terminus that affects channel availability also prevents deactivation and S4 return. Interactions with inactivation-conferring Kvβ subunits have been theorized to allosterically inhibit gating charge return through effects on the activation gate similar to the native N-terminal inactivation moiety of the full-length Shaker channel (Bezanilla et al., 1991). Recent data has also provided evidence that the open activation gate can inhibit S4 return until the channel can deactivate (Batulan et al., 2010), and complementary effects have been observed where activation gate opening is prevented pharmacologically with 4-aminopyridine, and S4 return is accelerated compared to the drug-free state (McCormack et al., 1994; Loboda and Armstrong, 2001). Therefore, it appears likely that this slow change in emission reports on interactions limiting the return of the S4 gating charge, on a slower time scale than expected for off-gating of Kv1 channels, consistent with return from a more stable activated state.  134  Kv1.2 inactivation is limited, but the channel shows fluorescence consistent with a relaxed state of S4  In many Kv channels, lengthy depolarization can result in the progression of channels from activated states into multiple inactivated states involving structural changes in both the pore (Hoshi et al., 1991; DeBiasi et al., 1993; Lopez-Barneo et al., 1993; Ogielska et al., 1995; Liu et al., 1996; Kiss and Korn, 1998; Larsson and Elinder, 2000; Loots and Isacoff, 2000; Claydon et al., 2007a) and voltage sensor domains (Olcese et al., 1997; Loots and Isacoff, 1998). Inactivation of ionic current in other channels such as Shaker has been shown to be associated with immobilization of S4 gating charge, which subsequently alters the time course and voltage dependence of its return upon hyperpolarization (Olcese et al., 1997; Loots and Isacoff, 1998). While these events have been thought to be correlated to changes in the pore domain (Loots and Isacoff, 2000; Elinder et al., 2001; Batulan et al., 2010), studies in the Ci-VSP protein, which has no pore domain, have shown similar results (Villalba-Galea et al., 2008), suggesting that these or other secondary changes involved in S4 stabilization may occur independent of the pore, and are inherent to the voltage sensing domain. In the absence of β-subunit modulation, Kv1.2 does not show N-type inactivation properties, and does not appear to undergo any significant degree of slow inactivation in response to prolonged depolarization (Paulmichl et al., 1991), certainly when compared to other channels in the Kv1 family. In fact, the channel contains a valine at a critical position near the selectivity filter that, in Shaker (T449V), has been shown to inhibit inactivation (Lopez-Barneo et al., 1993).  However, we report slow changes in the voltage sensor domain upon both  depolarization and repolarization, which appear to track ionic current deactivation and affect the rate of S4 return, based on VCF measurements. Such a slow reorganization of the voltage sensor domain is consistent with a Ci-VSP-like depolarization-induced stabilization of the activated S4  135  helix that must reverse for ionic current deactivation to occur, referred to in this protein as a relaxed activated state (Villalba-Galea et al., 2008). Alternatively, deactivation may be required to occur before S4 charge reversal can fully occur; from our data, it is unclear which motion is the rate-limiting step in the sequence, or how the S1-S2 region is involved in this process. However, our data support changes in the stability of the voltage-sensor domain independent of effects on pore conformation.  Role of the extracellular linkers in slow fluorescence-associated S4 stabilization  To better understand the specific molecular entities contributing to this slow quenching of Kv1.2 fluorescence emission, we investigated the roles of the different extracellular linkers by constructing Kv1.2/Shaker linker swap channels. Surprisingly, we were unable to confer this slow component to Shaker, or preserve this component in Kv1.2 with replacement of either extracellular linker. Attempts to investigate linker-linker interactions as a putative mechanism were unsuccessful, as the Shaker channel with Kv1.2 S1-S2 and S3-S4 extracellular linkers failed to produce functional channels, or gave no voltage-dependent fluorescence changes. Nonetheless we can make certain inferences or hypotheses about the roles of these linkers based on chimeras for which we were able to record fluorescence. For example, the absence of a slow quenching component in Shaker A359C, Kv1.2/Shaker S3S4L, or Shaker/Kv1.2 S1S2L channels led us to consider the role of the Shaker S3-S4 linker as a barrier to seeing secondary changes in the voltage sensing domain. It is quite possible that the longer S3-S4 linker in Shaker (31 amino acids compared to 20 in Kv1.2) is oriented around S4 in such a way as to prevent distant movements within the voltage sensor domain from altering the local probe environment. Of course, this presumes that secondary movements occur in Shaker which, if they do, are not  136  reported in our VCF measurements. In contrast, the absence of this component in Kv1.2/Shaker S1S2L and Shaker/Kv1.2 S3S4L may relate to different compositions of the S1-S2 linker, which we have shown to be important in reporting this reorganization. One potential explanation is that only the longer S1-S2 linker in Kv1.2 may extend into the S4 environment, in contrast to the shorter Shaker S1-S2 linker, which does not come into proximity of the fluorescent probe. These two hypotheses would explain the quantitative results presented here, but they of course would require further study in order to garner further support. Alternatively, changes in linker length and structure may also alter the positions of S2 and S3 and their negatively charged residues relative to S4, which may alter fluorescence in an unpredictable manner, and/or affect channel gating (which was observed in all of the chimera channels). From the standpoint of the associated S4 stabilization, assuming that Shaker gating charge immobilization and our report of slow Kv1.2 fluorescence are similar mechanisms, it appears that interactions between the extracellular linkers are not the primary facet of this reorganization. Rather, interactions with a probe attached at S4 may report on linker movements associated with more distant reorganizations within the voltage sensing domain.  Further studies needed to understand the nature of this reorganization  While our data are consistent with the Kv1.2 S4 segments entering a relaxed activated state, from which return is slow, further studies are needed to better understand these events and rule out alternate explanations. Others have shown in Shaker, using gating currents (Olcese et al., 1997), and we have shown via VCF (Figure 2.4), that S4 mobility is affected by changes in holding potential as indicated by a leftward shift in the voltage-dependence of Q-V and F-V relationships. In Kv1.2, no shift was observed when holding at potentials between -120 mV and  137  -50 mV. It would be of great interest to continue to explore this effect using more depolarized holding potentials, as well as the effect of membrane potential on the isolated fast fluorescence component reporting on S4 translocation alone in both Kv1.2 and chimera channels. Similar experiments exploring these effects on gating currents would be expected to shed further light on S4 mobility as a function of potential, ideally concomitantly with fluorescence through the use of a cut-open oocyte preparation. The data presented above paint a qualitative picture of Kv1.2 gating involving several aspects of the voltage sensing domain, but many quantitative questions remain unanswered. VCF provides little insight to the extent of protein movements. Furthermore, where two protein moieties are involved, VCF can sense relative changes but cannot discern which region is moving more, or not at all. These are very important questions that need to be answered in order to fully understand Kv1.2 activation, and the emerging role of all components of the voltage sensing domain. Lanthanide resonance energy transfer (LRET), which measures changes in photon emission between separate donor and acceptor probes, is highly sensitive to even small changes in donor-acceptor distance (Cha et al., 1999; Posson et al., 2005), and thus can provide quantitative measurements of changes in fluorophore environment as a function of convergence or divergence between two protein moieties and their tethered donor and acceptor probes. Strategic placement of these probes in Kv1.2 and the linker swap chimera channels throughout the S1-S2 linker, the S3-S4 linker and the pore domain would help to localize changes in the protein structure, and piece together the underlying molecular rearrangements.  From the  collection of data from multiple donor-acceptor pairs in these separate regions of the protein, it is very likely that we could obtain our best picture yet of how the entire voltage sensing domain responds to changes in transmembrane potential with both fast and slow rearrangements;  138  hopefully, such a detailed study would further resolve the discrepancies between what our data shows and what the crystal structure predicts.  Reconciling VCF data with the Kv1.2 crystal structure  Although our data are consistent with the S1-S2 linker residing in close proximity to S4, according to the crystal structure of the Kv1.2/2.1 chimera (Long et al., 2007), this region may not reside in a position to directly influence S4 environment (Figure 2.11). Such proximity to a fluorophore would need to be on the order of angstroms in order to quench, or alter the energy of an excited fluorophore, though indirect interactions could occur over a larger scale. However, fluorophore positioning in our model is approximate, and may be somewhat flexible in its movement, so it is difficult to rule out any possible interactions. In addition, the non-native Kv2.1 S3-S4 linker differs in length and sequence from the native channel, and may have had further secondary effects on the structural orientation of the voltage sensor domain, particularly S1 and S2. Indeed, S3-S4 linker replacement in our hands showed significant effects on channel gating (Figure 3.3). Kv1.2 itself has not been crystallized with its extracellular linkers in a native orientation (Long et al., 2005) due to the flexibility of these regions, and has not been crystallized in the resting state, but our data are consistent with a state-dependent S1-S2 and S3S4 linker interaction; further investigation would help elucidate whether these regions do in fact reside in close proximity to one another, and whether these interactions primarily occur in the activated or resting state. Computer modelling of the closed state Kv1.2 chimera (YarovYarovoy et al., 2006; Pathak et al., 2007) does not show these regions in close proximity (Figure 2.11), but until a true crystal structure of Kv1.2 in the closed state can be obtained, the possibility that the S1-S2 linker interacts with the S4 environment cannot be excluded.  139  Kv1.2 activation: more than just S4?  The high density of positively charged residues within the S4 helix make it an ideal candidate for conferring voltage-sensitivity to voltage-gated ion channels, a fact supported by gating current measurements from charge-deletion mutants (Seoh et al., 1996; Aggarwal and MacKinnon, 1996). It is known from these same studies that at least one residue in the S2 helix, E293 in Shaker, is important in gating charge movement as well (Seoh et al., 1996); however, what is less well understood is whether this residue is important in stabilizing positively charged S4 arginines within the lipid bilayer, or whether E293 traverses a fraction of the electric field during activation. Our fluorescence data from Kv1.2 channels supports the idea that S2 may undergo conformational changes during depolarization, relative to the external environment of the channel, and that the entire voltage-sensing domain plays a role in responding to changes in membrane potential. Voltage-dependent changes in the S1-S2 region, while not associated with any resolvable degree of gating charge movement, are very important to consider as far as their potential interactions with the S4 helix and, according to our data, stabilizing activated states of the voltage sensor. This is line with other work highlighting point mutations in the Kv1.2 voltage sensing domain, outside of S4, with profound effects on activation gating (Minor et al., 2000; Rezazadeh et al., 2007). While our data are applicable to Kv1.2, we hope that further studies in Shaker and other Kv channels will elucidate the complete role of the voltage sensing domain in contributing to channel activation.  Summary  Conformational changes in voltage-gated proteins remain the subject of great debate, and many different techniques exist for resolving the specific mechanics behind the different  140  processes. Study of the Kv1.2 channel is of great importance for attempting to resolve the intricacies of channel gating, due to our enhanced understanding of channel structure. In this thesis I have utilized VCF to investigate voltage-dependent changes in the voltage sensing domain. I have demonstrated that two separate independent processes occur upon depolarization – a rapid S4-associated movement that correlates well with Kv1.2 gating currents, and a slow conformational change that can be localized to other regions of the voltage sensing domain. These secondary changes do not appear to outwardly affect ionic current, but upon repolarization are rate limiting for channel deactivation, suggesting they may report on delayed S4 return associated with a stabilized activated conformation. We believe that these slow movements may be a report of movements found in Shaker and other voltage-gated proteins, but uniquely observable in Kv1.2 due to its extracellular linker structure. Corroborative observations were made in the presence of an auxiliary Kvβ1.2 subunit, which has been shown to delay S4 return through its interactions with the activation gate, and accentuates the report of slow fluorescence on repolarization. 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Thornhill. 2009. The Kv1.2 potassium channel: the position of an N-glycan on the extracellular linkers affects its protein expression and function. Brain Res 1251:16-29. Zhu, J., I. Watanabe, A. Poholek, M. Koss, B. Gomez, C. Yan, E. Recio-Pinto, and W. B. Thornhill. 2003. Allowed N-glycosylation sites on the Kv1.2 potassium channel S1-S2 linker: implications for linker secondary structure and the glycosylation effect on channel function. Biochem J 375:769-775.  146  Appendix A: Supplemental Material for Chapter 21  1  A version of this chapter has been submitted for publication as Supplemental Material to Chapter 2 of this thesis. Andrew J. Horne, Christian J. Peters, Thomas W. Claydon, & David Fedida. (2010). Fast and slow voltage-sensor rearrangements during activation gating in Kv1.2 channels detected using tetramethylrhodamine fluorescence. The Journal of General Physiology 136(1):83-99.  147  Supplemental Results  Kv1.2 WT channels do not show voltage-dependent changes in fluorescence emission Kv1.2 homotetramers carry a delayed rectifier outward current very similar in kinetics and voltage-dependence to that from Shaker channels and other members of the Kv1 family. In Xenopus oocytes, robust ionic currents can be recorded from cRNA-injected cells, as shown in Figure A.1A during 100 ms pulses to hyperpolarized and activating potentials. From a holding potential of -80 mV, activation occurs with depolarizations to approximately -30 mV, with greater depolarizations resulting in faster activation and larger currents. There was no observed reduction in current during the pulse, suggesting that minimal inactivation occurred during this time period. Kv1.2 WT channels have one external cysteine, C181, located near the outer end of S1. Although this residue is buried in the lipid membrane in the crystal structure (Long et al., 2007), it is unknown whether this residue is available for TMRM modification in the open or closed state and, if labelling can occur, whether this could produce a voltage-dependent deflection. Labelled WT channels showed ionic currents similar in size and activation to unlabelled Kv1.2 (Figure A.1B). No changes were observed between the conductance-voltage (G-V) relationships (Figure A.1C) or activation time constants (Figure A.1D) of unlabelled and labelled channels. Half-activation potentials, calculated to be 1.2 ± 1.6 mV for the unlabelled channels and -1.7 ± 1.5 mV for labelled Kv1.2, are both in agreement with other studies in mammalian cells (Watanabe et al., 2007) and oocytes (Russell et al., 1994; Ishii et al., 2001). At +60 mV, unlabelled Kv1.2 WT currents activated with a time constant of 3.4 ± 1.2 ms, similar to values observed in other studies (Paulmichl et al., 1991). Taken together, these data suggest that if  148  attachment of TMRM at residue C181 can occur based on secondary structure in the region, channel gating is not affected. To address whether a fluorophore attached at C181 can report voltage-dependent changes in channel structure, we recorded the fluorescence from labelled WT Kv1.2 channels (Figure A.1E).  We tested 21 cells, and found no evidence of any deviation from the baseline  fluorescence at any voltage, despite robust ionic currents at depolarized potentials. With longer (6.3 s) depolarizations, a small decaying fluorescence component was observed in some cells (Figure A.1F) even after attempts to control for fluorophore photobleaching. This decrease from baseline was only evident after approximately 2 s depolarizations at +60 mV, and may either reflect incomplete correction for photobleaching during long pulses, or a very slow voltagedependent fluorescent change detected by C181. The experiments of the present study utilized depolarizations of 100 ms or less, which allowed these very slow changes in fluorescence to be ignored.  149  Figure A.1. TMRM labelling of WT Kv1.2 channels does not alter activation kinetics, or give voltage-dependent fluorescence deflections. (A and B) Representative current traces recorded from unlabelled control (A) and TMRM-labelled (B) Kv1.2 WT channels (Kv1.2 WTTMRM) are shown for 100 ms pulses from -120 mV to +60 mV, from a holding potential of -80 mV. Although traces were recorded at 10 mV increments, only every third trace is shown for clarity. (C) Conductance-voltage relationships were calculated from peak current at the end of each pulse, and are shown as mean ± SEM for Kv1.2 WT (filled circles) and Kv1.2 WT-TMRM (hollow circles) channels. Boltzmann fits to the data give a V1/2 of 1.2 ± 1.6 mV and -1.7 ± 1.5 mV and slope factors of 18.0 ± 0.9 mV and 17.1 ± 0.8 mV for Kv1.2 WT and Kv1.2 WT-TMRM respectively (n = 7-8). (D) Mean activation time constants and standard error were calculated from -50 mV to +100 mV, for Kv1.2 WT and Kv1.2 WT-TMRM channels (n = 7-10). (E and F) Voltage-dependent fluorescence deflections were absent from cells expressing TMRM-labelled Kv1.2 WT channels at 100 ms (E), only surfacing as a minor deflection with long depolarizations (F), shown here with a 6.3 s depolarizing pulse. Dotted lines refer to the baseline level of fluorescence emission.  Previous work with fluorimetry in Shaker has looked at voltage-dependent changes sensed by TMRM at the extracellular end of S4, in the S3-S4 linker (Mannuzzu et al., 1996; Cha 150  and Bezanilla, 1997; Cha and Bezanilla, 1998; Claydon et al., 2007a; Claydon et al., 2007b). Kv1.2 was labelled in this region by introducing the A291C mutation, which is the homologue of Shaker A359C and is located 3 residues upstream of the first arginine in S4 (Figure 2.1). Despite the lack of a voltage-dependent fluorescence deflection at C181 alone (Figure A.1), we first measured Kv1.2 fluorescence from A291C in the background of the C181V mutation (Figure A.2). Deflections from Kv1.2 C181V A291C are shown in Figure A.2A during 100 ms pulses to voltages between -120 mV and +60 mV. Fluorescence signals were small, but at depolarized potentials exhibited a fast initial decrease in emission, followed by a slower quenching phase over the remainder of the pulse; this is best seen in the isolated waveform shown in the inset, obtained at +30 mV. Expression of Kv1.2 C181V A291C, as measured from ionic currents, was less than the Kv1.2 WT construct in oocytes, and the fluorescence signals, as seen in Figure A.2A, were quite small and difficult to resolve. In contrast, the fluorescence signals from channels containing the endogenous cysteine (Kv1.2 C181 A291C) were larger and allowed resolution of a fast and slow component at depolarized potentials (Figure A.2B). Overlay of normalized C181 A291C and C181V A291C Kv1.2 fluorescence deflections at +30 mV (insets panels of Figures A.2A and A.2B) shows that the fluorescence signals were identical (Figure A.2C), as has been previously reported by our lab (Peters et al., 2009). Furthermore, no differences were observed in the fluorescence-voltage (F-V) relationships (Figure A.2D). Taken together, these data suggest that removal of the endogenous cysteine residue in the outer S1 helix was not required to observe changes in the fluorescence environment around S4, and that its presence did not alter the overall fluorescence phenotype. Given the greater expression and larger fluorescence signals observed  151  with Kv1.2 A291C channels, all further experiments were carried out using the WT Kv1.2 (C181) background, into which S3-S4 linker mutations were introduced.  Figure A.2. A TMRM fluorophore attached to residue A291C reports identical voltagedependent changes in fluorescence in C181 and C181V channels. Representative Kv1.2 C181V A291C (A) and Kv1.2 C181 A291C (B) fluorescence deflections are shown for voltages between -120 mV (top trace) and +60 mV (bottom trace), from a holding potential of -80 mV. The inset depicts the change in fluorescence emission at +30 mV, with the scale bars as in the main panel. Arrows denote the baseline level of fluorescence emission. (C) An overlay of normalized fluorescence deflections following depolarization to +30 mV shows no difference in the Kv1.2 C181V A291C and Kv1.2 C181 A291C phenotypes. (D) Fluorescence-voltage relationships for Kv1.2 C181V A291C (filled circles) and Kv1.2 C181 A291C (hollow circles). Data are shown as mean ± SEM (n = 4-8).  Substitution of the Kv1.5 S2 transmembrane segment does not affect Kv1.2 fluorescence Figure A.3 shows representative fluorescence emission from TMRM attached to A291C in the Kv1.5-S23L-Kv1.2 chimera, in which the voltage sensing domain of Kv1.2 had been 152  replaced up to the intracellular end of S3.  Representative fluorescence emission and F-V  relationships (data not shown) were identical to Kv1.5-S12L-Kv1.2 A291C (Figure 2.8), suggesting that the fluorescence modifying portion of the Kv1.5 structure was limited to S1 or the S1-S2 linker, common to both chimera channels tested, rather than either the S2 helix or S2S3 linker segment.  Figure A.3. Representative fluorescence traces from the Kv1.5-S23L-Kv1.2 A291C chimera are similar in phenotype to the shorter chimera replacement. Data were collected using the same protocol to that in Figure 2.2.  Fluorescence scanning of the Kv1.2 voltage sensor reveals information about the slow fluorescence component A scan of a small portion of the S3-S4 linker of both Kv1.2 and Shaker revealed that the prominent slow quenching event originally observed from TMRM attachment to A291C was not exclusive to this site (Figure 2.3). Figure A.4 shows a model of Kv1.2 in the closed state (Pathak et al., 2007), highlighting residues found to give voltage-dependent fluorescence deflections in Kv1.2.  153  Figure A.4. Mapping of the S3-S4 linker scan residues onto the Kv1.2 closed state. Cartoon of the closed state Kv1.2 model generated by Pathak et al. (Pathak et al., 2007), with residues M288 (orange), S289 (cyan), L290 (magenta), A291 (red) and I292 (dark green) labelled and showing side chain residues. Blue residues correspond to I187 and T219, residues also shown to give voltage-dependent deflections upon TMRM labeling.  154  References  Cha, A. and F. Bezanilla. 1997. Characterizing voltage-dependent conformational changes in the Shaker K+ channel with fluorescence. Neuron 19:1127-1140. Cha, A. and F. Bezanilla. 1998. Structural implications of fluorescence quenching in the Shaker K+ channel. J Gen Physiol 112:391-408. Claydon, T. W., M. Vaid, S. Rezazadeh, S. J. Kehl, and D. Fedida. 2007a. 4-aminopyridine prevents the conformational changes associated with P/C-type inactivation in Shaker channels. J Pharmacol Exp Ther 320:162-172. Claydon, T. W., M. Vaid, S. Rezazadeh, D. C. Kwan, S. J. Kehl, and D. Fedida. 2007b. A direct demonstration of closed-state inactivation of K+ channels at low pH. J Gen Physiol 129:437-455. Ishii, K., K. Nunoki, T. Yamagishi, H. Okada, and N. Taira. 2001. Differential sensitivity of Kv1.4, Kv1.2, and their tandem channel to acidic pH: Involvement of a histidine residue in high sensitivity to acidic pH. J Pharmacol Exp Ther 296:405-411. Long, S. B., X. Tao, E. B. Campbell, and R. MacKinnon. 2007. Atomic structure of a voltagedependent K+ channel in a lipid membrane-like environment. Nature 450:376-382. Mannuzzu, L. M., M. M. Moronne, and E. Y. Isacoff. 1996. Direct physical measurement of conformational rearrangement underlying potassium channel gating. Science 271:213216. Pathak, M. M., V. Yarov-Yarovoy, G. Agarwal, B. Roux, P. Barth, S. Kohout, F. Tombola, and E. Y. Isacoff. 2007. Closing in on the resting state of the Shaker K+ channel. Neuron 56:124-140. Paulmichl, M., P. Nasmith, R. Hellmiss, K. Reed, W. A. Boyle, J. M. Nerbonne, E. G. Peralta, and D. E. Clapham. 1991. Cloning and expression of a rat cardiac delayed rectifier potassium channel. Proc Natl Acad Sci USA 88:7892-7895. Peters, C. J., M. Vaid, A. J. Horne, D. Fedida, and E. A. Accili. 2009. The molecular basis for the actions of K(V)beta1.2 on the opening and closing of the K(V)1.2 delayed rectifier channel. Channels (Austin ) 3:314-322. Russell, S. N., N. G. Publicover, P. J. Hart, A. Carl, J. R. Hume, K. M. Sanders, and B. Horowitz. 1994. Block by 4-aminopyridine of a Kv1.2 delayed rectifier K+ current expressed in Xenopus oocytes. J Physiol 481.3:571-584. Watanabe, I., J. Zhu, J. J. Sutachan, A. Gottschalk, E. Recio-Pinto, and W. B. Thornhill. 2007. The glycosylation state of Kv1.2 potassium channels affects trafficking, gating, and simulated action potentials. Brain Res 1144:1-18.  155  

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