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An examination of the slow inactivation and resting inactivation of Kv1.5 and ShakerIR Cheng, Yen May 2010

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 AN EXAMINATION OF THE SLOW INACTIVATION AND RESTING INACTIVATION OF Kv1.5 AND ShakerIR CHANNELS  by   Yen May Cheng  B.Sc., The University of British Columbia, 2003 M.Sc., The University of British Columbia, 2005    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)   September 2010      © Yen May Cheng, 2010 ii Abstract External H +  and Ni 2+  ions inhibit Kv1.5 channels by increasing current decay during a depolarizing pulse and reducing the peak current. A similarly accelerated current decay and reduced peak current amplitude have also been described in ShakerIR channels at low extracellular pH, and in the fast inactivating ShakerIR T449K, T449A and Kv1.5 H463G mutants in 0 K + o at pH 7.4. While the increased current decay may be attributed to an enhancement of the slow inactivation of open channels at depolarized potentials, the reduction of peak current cannot. Using standard whole-cell voltage-clamp techniques and fast perfusion changes, the hypothesis that the decreased peak current is due to the induction (by H + , Ni 2+ , or removal of K + ) of a resting inactivation process was investigated. It was found that exposure of resting Kv1.5 channels to H +  or Ni 2+  causes a decrease in the ability of Ba 2+  ions to move between the external solution and a deep binding site within the selectivity filter. This result is consistent with an outer pore constriction at rest, possibly similar to that which occurs during slow inactivation. An examination of the time courses for the onset of H + - or Ni 2+ -enhanced slow inactivation and resting inactivation of Kv1.5 showed that, compared to slow inactivation at +50 mV, the onset of resting inactivation at −80 mV is a relatively slow process. The bi- exponential recovery following H + - or Ni 2+ -enhanced slow inactivation or resting inactivation had time constants similar to those for recovery from control slow inactivation. Due to the greatly accelerated slow inactivation of ShakerIR at low pH and the relatively slower perfusion kinetics of our system, it was difficult to assess the contribution of resting inactivation to the H + - induced loss of ShakerIR current. However, analogous kinetic analyses performed on the fast- inactivating ShakerIR T449 and Kv1.5 H463 mutants in 0 K + o also showed that the recovery time courses for resting inactivation and slow inactivation were similar, suggestive of similar iii recovery pathways. Together, the results strongly suggest that, under specific conditions, the slow inactivation process in wild-type and mutant Kv1.5 and ShakerIR channels can uncouple from activation and occur at resting potentials. iv Preface This dissertation includes work that has been, or will be, published in peer-reviewed research journals. The introductory sections for Chapters 2 and 3, which contain the published work, have been re-worded to better integrate the work into the thesis as a whole. The remaining portions of Chapters 2 and 3 remain unaltered from the published versions, save for minor editorial or formatting changes made for consistency. Portions of the original introduction from each published article are now presented in Chapter 1. The following list details the relative contributions of the authors to the work presented.  Chapter 2: Cheng YM, Fedida D, Kehl SJ. 2008. External Ba 2+  block of human Kv1.5 at neutral and acidic pH: evidence for H + o-induced constriction of the outer pore mouth at rest. Biophysical Journal. 95(9): 4456-4468.  Yen May Cheng was responsible for ~80% of the work presented in this Chapter, including i) designing and planning experiments, ii) collecting and analysing the majority of the data, iii) writing the majority of the manuscript. Steven Kehl‟s contribution included i) conceiving of the project, ii) assisting in the collection of data used in the examination of the frequency dependency of Ba 2+  binding and developing the resulting model, iii) writing the section of the manuscript detailing the aforementioned model. David Fedida and Steven Kehl critically reviewed the manuscript and suggested revisions.  Chapter 3: Cheng YM, Fedida D, Kehl SJ. 2010. Kinetic analysis of the effects of H +  or Ni 2+  ions on Kv1.5 current shows that both ions enhance slow inactivation and induce resting inactivation. Journal of Physiology. 588(16): 3011-3030.  Yen May Cheng was responsible for ~60% of the work presented here, including i) collecting and analysing ~30% of the data, and ii) writing and revising the manuscript. The rest of the data was collected and analysed by Steven Kehl, who conceived of the project and also performed the numerical simulations presented in Figures 8 and 9. David Fedida and Steven Kehl critically reviewed and revised the manuscript.  Chapter 4: Resting inactivation of ShakerIR channels induced by low external pH or mutation of an outer pore residue.  Yen May Cheng contributed ~90% of the work presented in this chapter, a version of which will be submitted for publication in the future. The work performed included i) planning of the experiments and project management, ii) collecting and analysing the data, iii) generating and testing the T449 mutants, iv) writing the chapter. Steven Kehl conceived of the project and collected preliminary data on the H + -induced resting inactivation of wt ShakerIR. Calvin Tong performed the experiments examining the 0 K + o-induced resting inactivation of Kv1.5 H463G.   v Table of Contents Abstract ........................................................................................................................................... ii Preface............................................................................................................................................ iv Table of Contents .............................................................................................................................v List of Tables ................................................................................................................................. ix List of Figures ..................................................................................................................................x List of Abbreviations .................................................................................................................... xii Acknowledgements ...................................................................................................................... xiv Dedication ......................................................................................................................................xv 1 Introduction ............................................................................................................................1 1.1 OVERVIEW ......................................................................................................................1 1.2 THE STRUCTURE AND FUNCTION OF KV CHANNELS .........................................................2 1.2.1 Basic Kv channel architecture ....................................................................................2 1.2.2 The diverse family of Kv channels .............................................................................4 1.2.3 Detailed features of Kv channel structure ...................................................................5 1.2.3.1 The pore domain ...................................................................................................5 1.2.3.2 The voltage-sensing domain ...............................................................................11 1.2.4 An electromechanical view of activation and deactivation ......................................14 1.3 INACTIVATION OF KV CHANNELS ..................................................................................16 1.3.1 N-type inactivation....................................................................................................16 1.3.2 Slow or P/C-type inactivation ...................................................................................18 1.3.2.1 Slow inactivation involves the outer pore region ...............................................19 1.3.2.2 The role of the selectivity filter ...........................................................................22 1.3.2.3 Slow inactivation involves a localized conformational change in the selectivity filter ...................................................................................................24 1.3.2.4 The voltage sensor may be involved in slow inactivation ..................................26 1.3.3 U-type inactivation....................................................................................................28 1.3.4 Preferential closed-state inactivation of Kv4 channels .............................................29 1.4 RESTING INACTIVATION AND SCOPE OF THE THESIS ......................................................32 1.4.1 The effects of low pH on Kv1.5 and ShakerIR .........................................................32 1.4.2 The [K + ]o-dependent loss of availability of fast-inactivating ShakerIR T449 and Kv1.5 H463G mutants ..............................................................................................34 1.4.3 Objectives .................................................................................................................35 2 External Ba 2+  block of human Kv1.5 at neutral and acidic pH: evidence for H + - induced constriction of the outer pore mouth at rest ........................................................38 2.1 INTRODUCTION .............................................................................................................38 2.2 METHODS .....................................................................................................................40 2.2.1 Cell preparation .........................................................................................................40 2.2.2 Recording solutions ..................................................................................................40 2.2.3 Fast perfusion ............................................................................................................41 2.2.4 Electrophysiological procedures ...............................................................................41 2.2.5 Data analysis .............................................................................................................42 2.3 RESULTS .......................................................................................................................48 vi 2.3.1 Ba 2+  blocks closed Kv1.5 channels at pH 7.4 ...........................................................48 2.3.2 τblock is not a linear function of [Ba 2+ ] .......................................................................53 2.3.3 Ba 2+  block of Kv1.5 is dependent on stimulation frequency ....................................54 2.3.4 Ba 2+  causes a weak fast block of open Kv1.5 channels ............................................61 2.3.5 Ba 2+  block of Kv1.5 at rest and open channel dissociation ......................................63 2.3.6 Ba 2+  does not block Kv1.5 at low pH .......................................................................67 2.3.7 Ba 2+  unbinding is inhibited at low pH ......................................................................70 2.4 DISCUSSION ..................................................................................................................71 2.4.1 Ba 2+  block at pH 7.4 ..................................................................................................71 2.4.2 Ba 2+  block of Kv1.5 versus ShakerIR .......................................................................72 2.4.3 Low pH prevents movement of Ba 2+  to and from its deep binding site ...................74 2.4.4 Is the H + -induced pore constriction indicative of resting inactivation? ....................74 3 Kinetic analysis of the effects of H +  or Ni 2+  on Kv1.5 current shows that both ions enhance slow inactivation and induce resting inactivation. ......................................76 3.1 INTRODUCTION .............................................................................................................76 3.2 METHODS .....................................................................................................................78 3.2.1 Cell preparation .........................................................................................................78 3.2.2 Recording solutions ..................................................................................................78 3.2.3 Fast solution exchange ..............................................................................................78 3.2.4 Definitions of terms ..................................................................................................79 3.2.5 Signal recording and data analysis ............................................................................82 3.2.6 Numeric simulation ...................................................................................................83 3.3 RESULTS .......................................................................................................................84 3.3.1 External H +  and Ni 2+  enhance the current decay and reduce peak current ...............84 3.3.2 Decreasing external pH during a depolarization enhances slow inactivation ...........86 3.3.3 Current can recover from a transient decrease in pH during a depolarization ..........88 3.3.4 The onset of low pH-induced resting inactivation is slow ........................................89 3.3.5 Low pH-induced resting inactivation is sensitive to [K + ]o but not [K + ]i ..................90 3.3.6 Transient exposure to Ni 2+  during a pulse enhances slow inactivation ....................90 3.3.7 The time course of Ni 2+ -induced resting inactivation is slow ...................................93 3.3.8 Recovery from H + -enhanced slow inactivation or resting inactivation is fast ..........96 3.3.9 Recovery from Ni 2+ -enhanced slow inactivation or resting inactivation is slow ...100 3.3.10 Recovery from H + - and Ni 2+ -induced resting inactivation at depolarized potentials .................................................................................................................101 3.3.11 The time course for K + o-facilitated recovery from resting inactivation is the same as that for recovery from enhanced slow inactivation ...................................104 3.4 DISCUSSION ................................................................................................................108 3.4.1 Multiple inactivated states can explain the recovery overshoot during a depolarizing pulse ...................................................................................................111 3.4.2 Low pH and Ni 2+  also induce resting inactivation of Kv1.5 channels ....................111 3.4.3 Modelling the enhancement of resting inactivation by H +  and Ni 2+  .......................112 3.4.4 Are the effects of H +  or Ni 2+  on open and resting Kv1.5 channels related? ...........116 3.4.5 Evidence from other Kv channels for resting inactivation .....................................117 3.4.6 Concluding remarks ................................................................................................118 4 An examination of the pH or K + o-dependent loss of current through wt ShakerIR or fast-inactivating ShakerIR and Kv1.5 mutant channels ...........................119 4.1 INTRODUCTION ...........................................................................................................119 vii 4.2 METHODS ...................................................................................................................121 4.2.1 Site-directed mutagenesis and cell preparation .......................................................121 4.2.2 Recording solutions ................................................................................................121 4.2.3 Fast solution exchange ............................................................................................122 4.2.4 Definitions of terms ................................................................................................122 4.2.5 Signal recording and data analysis ..........................................................................123 4.3 RESULTS .....................................................................................................................125 4.3.1 Low external pH enhances slow inactivation and reduces peak current of ShakerIR .................................................................................................................125 4.3.2 Slow inactivation is enhanced during a depolarization by a transient decrease in pH ............................................................................................................................127 4.3.3 The time course of H + -induced resting inactivation ...............................................129 4.3.4 Recovery from H + -enhanced slow inactivation is fast ............................................133 4.3.5 The open probability of fast-inactivating T449 mutants is [K + ]o-dependent ..........137 4.3.6 ShakerIR T449 mutants undergo time-dependent resting inactivation in 0 mM K + o ...........................................................................................................................139 4.3.7 Recovery of T449 mutants from 0 K + o-induced resting inactivation has the same time course as recovery from slow inactivation .....................................................141 4.3.8 Recovery from slow inactivation and resting inactivation follow the same time course in the Kv1.5 H463G mutant ........................................................................143 4.4 DISCUSSION ................................................................................................................146 4.4.1 Decreasing pH or [K + ]o enhances slow inactivation of wt or mutant ShakerIR channels, respectively .............................................................................................146 4.4.2 Low pH may induce resting inactivation of wt ShakerIR channels ........................147 4.4.3 Comparison with the H + -dependent current loss of Kv1.5 .....................................148 4.4.4 Enhanced slow inactivation in the ShakerIR T449 and Kv1.5 H463G mutants .....149 4.4.5 K + o-dependent resting inactivation of fast-inactivating ShakerIR and Kv1.5 mutant channels ......................................................................................................150 4.4.6 Comparisons with [K + ]o-dependent current loss in other channels ........................151 5 Discussion ............................................................................................................................154 5.1 SUMMARY OF RESULTS ...............................................................................................154 5.2 IN LOW pH OR Ni 2+  THE OUTER PORE OF KV1.5 IS CONSTRICTED AT REST ...................156 5.3 THE H + - OR Ni 2+ -INDUCED RESTING INACTIVATION OF KV1.5 OR ShakerIR ...............158 5.3.1 A comparison of the kinetics of slow inactivation and resting inactivation ...........159 5.3.2 Molecular mechanisms of H + -induced resting inactivation in Kv1.5 and ShakerIR .................................................................................................................161 5.3.2.1 Modulation of the turret ....................................................................................161 5.3.2.2 Modulation of the selectivity filter ...................................................................162 5.4 RESTING INACTIVATION IN FAST-INACTIVATING MUTANTS IS HIGHLY SENSITIVE TO [K + ]O ......................................................................................................................165 5.4.1 How does K + o modulate resting inactivation? ........................................................166 5.5 IMPLICATIONS OF RESTING INACTIVATION ..................................................................170 5.5.1 Coupling between activation and slow inactivation ...............................................170 5.5.2 Involvement of the voltage sensor in slow inactivation..........................................172 5.6 FINAL REMARKS ..........................................................................................................174 References ...................................................................................................................................176 Appendix .....................................................................................................................................193 viii A.1 THE FAST PERFUSION SYSTEM .....................................................................................193 A.1.1 The Warner Instruments SF-77B Perfusion Fast Step ............................................193 A.1.2 The custom fast application tool .............................................................................196 A.1.3 Characterizing the time course of the fast solution change. ...................................197  ix List of Tables Table 3.1. Low pH biases inactivation to a state from which recovery is fast. .............................99 Table 4.1. Low pH biases ShakerIR inactivation to a state from which recovery is fast. ...........136      x List of Figures Figure 1.1. The general structure of voltage-gated ion channel pore-forming α-subunits. .............3 Figure 1.2. The KcsA crystal structure provides a representation of the pore domain of Kv channels. .................................................................................................................7 Figure 1.3. The crystal structure of the Kv1.2 channel. .................................................................13 Figure 2.1. Ba 2+  blocks Kv1.5 in a concentration- and frequency-dependent manner. .................49 Figure 2.2. Sequence alignment of hKv1.5 and Shaker from the C-terminal end of S5 to the end of the selectivity filter. ...................................................................................53 Figure 2.3. The steady-state level of Ba 2+  block is dependent on the stimulation frequency and the [Ba 2+ ]. .............................................................................................................56 Figure 2.4. A model of the frequency-dependent Ba 2+  binding replicates the time- dependence of block onset and reversal. ....................................................................60 Figure 2.5. Fast application of Ba 2+  during a long depolarising pulse reveals very weak open channel block by 20 mM Ba 2+ . ..........................................................................62 Figure 2.6. Ba 2+  unbinding from the open state is voltage-dependent and inhibited by increasing the [K + ]o. ....................................................................................................65 Figure 2.7. Low pH inhibits both Ba 2+  binding and unbinding from Kv1.5. .................................69 Figure 3.1. Simplified gating schemes describing the putative actions of H +  or Ni 2+  ions on Kv1.5 channels at resting and depolarized potentials. ...........................................80 Figure 3.2. Effects of external H +  and Ni 2+  on Kv1.5 current. ......................................................85 Figure 3.3. The time course of enhanced slow inactivation and resting inactivation is pH dependent. ...................................................................................................................87 Figure 3.4. The time course of enhanced slow inactivation and resting inactivation is also [Ni 2+ ]-dependent. ........................................................................................................92 Figure 3.5. Ni 2+  prevents Ba 2+  entry and exit from the pore binding site. .....................................95 Figure 3.6. The kinetics of recovery from H + - or Ni 2+ -enhanced slow inactivation or resting inactivation. .....................................................................................................97 Figure 3.7. Current recovery during a test depolarizing pulse occurs following either H + - or Ni 2+ - enhanced slow inactivation or resting inactivation. ....................................102 Figure 3.8. The time course for recovery from 0 K + o-induced resting inactivation is the same as that for recovery from enhanced slow inactivation. ....................................107 Figure 3.9. Theoretical outcomes of the modulation of slow inactivation by ligand binding. .....................................................................................................................110 Figure 3.10. Numerical simulation of resting inactivation caused by Ni 2+  or low pH. ...............113 Figure 4.1. Decreases in external pH accelerate slow inactivation of ShakerIR and reduce peak current amplitude. ............................................................................................126 Figure 4.2. Current recovers from a transient decrease in pH applied during a depolarizing pulse. ....................................................................................................128 xi Figure 4.3. The time course of resting inactivation is time- and pH-dependent ..........................132 Figure 4.4. The recovery of ShakerIR channels from H + -enhanced slow inactivation is fast. ............................................................................................................................134 Figure 4.5. The effect of external [K + ]o on the amplitude of macroscopic currents through fast-inactivating ShakerIR mutants. ..........................................................................138 Figure 4.6. Removing external K +  induces a time dependent decline in current amplitude in fast-inactivating ShakerIR mutants. .....................................................................140 Figure 4.7. The recovery of T449K and T449A ShakerIR mutants from slow inactivation and resting inactivation induced by 0 K + o follows the same time course. ................142 Figure 4.8. Recovery at pH 7.4 of the Kv1.5 H463G mutant from slow inactivation and resting inactivation follow similar time courses. ......................................................145 Figure A.1. The Warner Instruments SF-77B Fast Step Perfusion system. ................................194 Figure A.2. Optimization of perfusion pipettes for use in the SF-77B Fast Step system. ...........195 Figure A.3. The custom fast application tool. ..............................................................................196 Figure A.4. Characterization of the perfusion kinetics. ...............................................................198  xii List of Abbreviations Amino Acid 3 letter code 1 letter code Alanine Ala A Arginine Arg R Asparagine Asn N Aspartate Asp D Cysteine Cys C Glutamate Glu E Glutamine Gln Q Glycine Gly G Histidine His H Isoleucine Ile I Leucine Leu L Lysine Lys K Methionine Met M Phenylalanine Phe F Proline Pro P Serine Ser S Threonine Thr T Tryptophan Trp W Tyrosine Tyr Y Valine Val V  A state - available state at −80 mV A0 - initial level of channel availability Arec,f - fast component of current recovery Arec,s - slow component of current recovery A-L state - ligand-bound available state at −80 mV δBa,off - electrical distance between the deep Ba 2+  binding site and the rate-limiting barrier for exit CL - cycle length FAT - fast application tool HEK-293 - human embryonic kidney cell line ID - inside diameter IKur - ultrarapid delayed rectifier current Inorm - normalized test current amplitude kx - rate constant KBa,d - apparent equilibrium dissociation constant of deep Ba 2+  binding site KBa,s - apparent equilibrium dissociation constant of superficial Ba 2+  binding site Kd - equilibrium dissociation constant Kv - voltage-gated potassium channel MEM - Minimum Essential Medium MTS reagents - methanethiosulfonate reagents NaV - voltage-gated sodium channel xiii nH - Hill coefficient O state - open state O-L state - ligand-bound open state OD - outside diameter OI state - open-but-slow-inactivated state OI-L state - ligand-bound open-but-slow-inactivated state Pinitial - initial proportion of Kv1.5 current blocked by Ba 2+  PB,SS - proportion or macroscopic current blocked by Ba 2+  at the steady-state Pd - proportion of channels blocked by Ba 2+  at deep site after prolonged depolarization pKa - negative logarithm of the apparent acid dissociation constant Po - open probability Pr - proportion of channels blocked by Ba 2+  at deep site after prolonged rest SCAM - substituted cysteine accessibility method ShakerIR  - fast-inactivation removed Shaker mutant τact,app - apparent activation time constant td - average amount of time that a channel is open during a depolarizing pulse tr - time spent at holding potential between test pulses τblock - time constant for the onset of Ba 2+  block τd - time constant for Ba 2+  unblock during depolarization τinact - time constant of slow inactivation onset τr - time constant for Ba 2+  block at the holding potential τRI - time constant of resting inactivation onset τunblock - time constant for recovery from Ba 2+  block TEA - tetraethylammonium U state - unavailable state at −80 mV U-L state - ligand-bound unavailable state at −80 mV v/v - volume/volume percentage solution wt - wild-type   xiv Acknowledgements Firstly, I am indebted to Dr. Steven Kehl for his gifts of patience, guidance and support. His unswerving work ethic and enthusiasm for research have been a continuous source of inspiration. I would also like to express my gratitude to the members of my graduate supervisory committee, Drs. David Fedida, Eric Accili and David Mathers, for their support and encouragement throughout my doctoral program. Special thanks are also extended to Dr. Ed Moore for his kind words and advice during my graduate training. During the course of my doctoral studies, I have been fortunate to have the support of the students and staff of the Department of Cellular and Physiological Sciences and, particularly, of the Cardiovascular Research Group at the Life Sciences Centre, for which I am very grateful. Thanks must also be extended to Fifi Chiu and Kyung Hee Park for their technical support in the lab. Thank you to my close friends for their encouragement and for helping me maintain a balance between school and the “real world”. To my family – Frank, Leng, Ming and Shawn, I am so very appreciative of your continued unconditional and “unquestioning” love and support these past five years. Finally, I am especially grateful to Andrew Horne for always knowing how to make me laugh and for sharing this and so many other adventures with me.  Financial support was provided by a Natural Sciences and Engineering Research Council of Canada – Postgraduate Scholarship (Doctoral) and a Michael Smith Foundation for Health Research Senior Graduate Studentship.  xv Dedication    To Frank and Leng     1 1 Introduction 1.1 OVERVIEW Electrical signalling requires the rapid movement of ions across the cell membrane. This is achieved by ion channels, which are macromolecular proteins that span the lipid bilayer and form selective pores through which ions can move down their electrochemical gradient. For example, voltage-gated potassium (Kv) channels allow K +  ions to flow out of the cell down their electrochemical gradient in response to membrane depolarization. Kv channels are highly selective for K +  over Na +  ions and have an important functional role in the activity of excitable tissues such as nerves and muscles, where they are responsible for repolarizing, or returning, the membrane potential to its resting level following a period of excitation, or depolarization (Pongs, 2008). Therefore, the opening and closing of Kv channels is crucial in the determination of action potential duration. The human Kv channel family is encoded by some 40 genes and represents the largest family of the K +  channels, underscoring their significance in the regulation of cellular excitability and other functions, such as Ca 2+  signalling or cell volume regulation (Gutman et al., 2005; Wulff et al., 2009; Harmar et al., 2009). That various cardiovascular and neurologic disorders are associated with inherited mutations and/or pharmacologic modulation of Kv channels (Gutman et al., 2005; Pongs, 2008; Wulff et al., 2009) further highlights the importance of understanding the mechanisms governing their activity. In this introduction, I will provide a brief overview of the structural and functional aspects of Kv channel gating, followed by an in-depth review of the different types of Kv channel inactivation. As detailed at the end of the chapter, the focus of the work described in this dissertation is on the processes involved in inactivation of members of the Kv1 family of Kv channels, specifically Kv1.5 and the fast-inactivation removed Shaker mutant (ShakerIR). 2 1.2 THE STRUCTURE AND FUNCTION OF KV CHANNELS 1.2.1 Basic Kv channel architecture The Shaker channel from Drosophila melanogaster was the first Kv channel to be cloned, in 1987 (Papazian et al., 1987; Kamb et al., 1987; Baumann et al., 1987), and its sequence bore a striking structural similarity to voltage-gated Na +  (NaV) and Ca 2+  channels that had also recently been cloned (Noda et al., 1984; Tanabe et al., 1987; for a review, see also Choe et al., 1999). From subsequent studies, Kv channels were shown, in their simplest form, to be comprised of 4 identical α-subunits arranged, with four-fold symmetry, around a central pore (Figs. 1A and B; Isacoff et al., 1990; Börjesson & Elinder, 2008). By comparison, voltage-gated Na +  and Ca 2+  channels have a single pore-forming subunit comprised of four non-identical and linked domains, or pseudosubunits, each of which is analogous to the Kv channel α-subunit motif (Noda et al., 1984; Tanabe et al., 1987). Each Kv channel α-subunit (and Na+ or Ca2+ channel pseudosubunit) has 6 transmembrane segments, S1 – S6; a re-entrant pore loop (P-loop), and cytoplasmic N- and C-terminal domains (Fig. 1A; Börjesson & Elinder, 2008). Notably, the S4 segment contains a set of regularly spaced basic residues, leading to the prediction and subsequent finding that S4 acts as a voltage sensor, while the S1 – S3 segments complete the voltage sensing domain and serve to help stabilize S4 within the lipid bilayer (Fig. 1A; Noda et al., 1984; Aggarwal & Mackinnon, 1996; for a review, see Börjesson & Elinder, 2008). The pore domain of each subunit has been shown to comprise the S5 to S6 regions, including the P-loop (Fig. 1A; MacKinnon & Yellen, 1990; MacKinnon et al., 1990; Yellen et al., 1991). Together, the voltage sensing domain confers changes in membrane voltage to the channel pore, which opens and closes to regulate K +  current (Fig. 1C). For a detailed review of these early studies in Kv channel topology and structure, readers are directed to a number of excellent review articles 3 (e.g. Choe et al., 1999; Hille, 2001; Yellen, 2002; Tombola et al., 2006; Börjesson & Elinder, 2008).   Figure 1.1. The general structure of voltage-gated ion channel pore-forming α-subunits.  A, Each α-subunit is composed of six transmembrane helices, a re-entrant pore loop, and intracellular N- and C-terminal domains. S1 – S4 forms the voltage sensing domain (blue). The S4 segment contains regularly spaced, positively charged residues. The pore domain (pink) is formed by S5 – S6, with the selectivity filter being comprised of the C-terminal region of the P- loop. B, Four α-subunits form an ion channel with a central pore-forming unit (pink) surrounded by four voltage sensing domains. The intracellular N- and C-terminal regions are removed for clarity. C, A change in membrane voltage leads to the movement of S4 charges in the voltage sensing domain that results in either opening or closing of the pore domain. Adapted with kind permission from Springer Science + Business Media: Börjesson SI & Elinder F (2008), Structure, function, and modification of the voltage sensor in voltage-gated ion channels, Cell Biochemistry and Biophysics, 52, 149-174, Figure 1.  4 1.2.2 The diverse family of Kv channels The cloning of the Shaker gene was quickly followed by the identification of other related channels, including three other Drosophila genes: Shal, Shab and Shaw, that code for Kv channels with different gating kinetics (Wei et al., 1990). In humans, Kv channels are encoded by 40 genes and, depending on their structural relatedness and predominant functional characteristics, can be grouped into 12 subfamilies (Gutman et al., 2005; Wulff et al., 2009; Harmar et al., 2009). Following a period of sometimes confusing terminology, a systematic method of naming mammalian Kv channels was adopted, wherein proteins are identified as Kvx.y, with the integers x and y indicating the subfamily and order of discovery in that particular subfamily, respectively (Gutman et al., 2005). An analogous nomenclature has been implemented for Kv channel genes, which are labelled as KCNMN, where M is a letter assigned to a particular subfamily and N is a number assigned to the particular channel. Correspondingly, the Drosophila channels Shaker, Shab, Shaw and Shal are now considered prototypes of the mammalian Kv1 to Kv4 subfamilies, which are derived from the KCNA to KCND gene families, respectively. The large number of genes encoding Kv channels speaks to their diverse nature. The Shaker gene alone has been shown to give rise to several channel isoforms with slightly different functional characteristics, due to alternative splicing of the mRNA (Pongs et al., 1988; Timpe et al., 1988; Iverson et al., 1988). Similarly, members of a Kv channel subfamily often have subtly different biophysical and/or pharmacological profiles that may be due to small differences in their primary sequences, particularly in the N- and C-termini and in the linkers between the transmembrane segments. Importantly, despite the sometimes large variations in their primary sequences, all Kv channels retain the same basic tetrameric structure based on an α–subunit with six transmembrane segments as outlined in the previous section, distinguishing them from other, 5 non-voltage-gated K +  channels, such as the inwardly rectifying (Kir) or two-pore (K2P) K +  channels. 1.2.3 Detailed features of Kv channel structure Key advances of our understanding of Kv channel structure came with the determination of the X-ray crystal structures of the bacterial K +  channels KcsA, KvAP (Doyle et al., 1998; Jiang et al., 2003) and the mammalian Kv1.2 channel (Long et al., 2005a; Long et al., 2005b; 2007). The atomic resolution of these structures confirmed the earlier conclusions regarding the basic tetrameric assembly of α-subunits, and provided further insight into the structure-function relationships of Kv channels. 1.2.3.1 The pore domain Although Kv channels, and K +  channels as a whole, are functionally and even structurally diverse, several factors point to a strong conservation of the general structure of the pore and mechanism of permeation. The strong selectivity for K +  ions coupled with the high throughput rate points to an environment that is highly specialized and favourable to the passage of K +  through the pore (Yellen, 2002). Figure 1.2A shows an alignment of the outer pore regions of Shaker, the human Kv1 subfamily and KcsA. The alignment highlights the conservation of residues within the outer pore, particularly in the P-loop, which contains the K +  channel signature sequence (TxxTxGYGD, highlighted in blue; please see the List of Abbreviations for amino acid codes) that forms the selectivity filter (Heginbotham et al., 1994; Doyle et al., 1998). Close inspection of the alignment reveals that, as expected, the primary sequence identity is highest amongst the Kv1 channel subfamily, including Shaker. However, following the elucidation of its crystal structure (Fig. 1.2B), the internal pH-sensitive KcsA channel isolated from Streptomyces lividans, has become a powerful tool in the study of permeation and 6 selectivity in the K +  channel pore. Although sharing relatively little primary sequence identity, the M1 and M2 transmembrane helices of the KcsA α-subunit are functionally analogous to the S5 and S6 helices of Kv channel α-subunits (Lu et al., 2001; Lu et al., 2002), and the KcsA structure is considered to provide a good representation of a Kv channel pore (Kurata & Fedida, 2006). 7   Figure 1.2. The KcsA crystal structure provides a representation of the pore domain of Kv channels.   8 Figure 1.2. The KcsA crystal structure provides a representation of the pore domain of Kv channels. A, Amino acid sequence alignment for the pore region (from the C-terminal end of S5 to the beginning of the cytoplasmic C-terminal domain) of Shaker, KcsA, the human Kv1 family, and a variety of other human Kv channel clones. The amino acid sequence of Shaker is listed in full; identical residues in other channels are represented by a dash (-). At the bottom of the alignment, conserved substitutions (i.e. replacement of an acidic residue with another acidic residue) are denoted at with (:); semi-conserved substitutions are marked with a (.). Blue amino acid symbols mark the K +  channel signature sequence; the dashed box identifies residues that contribute backbone carbonyl oxygen atoms to the selectivity filter. The red box highlights residues equivalent to Shaker T449. The purple box demarcates the conserved glycine forming the “glycine” hinge and the green boxes demarcate the PxP motif present in Kv channels, but absent in KcsA. B, Side view of two opposing subunits of KcsA. The S5-P linker is marked as the “turret”, while the N- and C-terminal halves of the P-loop are marked as the “pore helix” and “selectivity filter”, respectively. The M1 and M2 helices of KcsA are homologous to the S5 and S6 helices of Kv channels. Residues within the selectivity filter are shown in the “ball-and-stick” format and are magnified in panel C. K +  ions are shown as green spheres. Side chain carboxyl oxygen atoms of KcsA residue D80 (Shaker D447) are in close proximity to W67 (Shaker W434), and these two residues may form an intra-subunit hydrogen bond. D, Top-down view of the selectivity filter, showing how the side chain hydroxyl group of Y78 (Y445 in the Shaker GYG sequence) may be stabilized by hydrogen bonding with W68 (Shaker W435) of an adjacent subunit, and van der Waals interactions with W67 (Shaker W434) of the same subunit. Adapted from Figure 1 in Progress in Biophysics and Molecular Biology, Kurata HT & Fedida D (2006), A structural interpretation of voltage-gated potassium channel inactivation, 92, 185-208, with permission from Elselvier. 9 Figure 1.2B shows a side-view of two opposing α-subunits of the KcsA channel (Doyle et al., 1998; Kurata & Fedida, 2006). In addition to the positions of the transmembrane M1 and M2 helices (homologous to S5 and S6 in Kv channels), the structure shows that the M1-P loop linker (S5-P linker in Fig. 1.2A) forms an extracellular “turret”, while the P-loop can be separated into a pore helix and a region contributing to the selectivity filter. From the crystal structure, the mechanisms for K +  selectivity and high throughput become apparent. As predicted from earlier mutational analyses (Heginbotham et al., 1992; Heginbotham & MacKinnon, 1993; Heginbotham et al., 1994), the selectivity filter is formed by the K +  channel signature sequence in the C-terminal half of the P-loop. Amino acids in this region of each subunit (shown in “ball- and-stick” format, and magnified in Fig. 1.2C) provide a “backbone” of carbonyl oxygen atoms that project into the central axis of the pore (Doyle et al., 1998; Zhou et al., 2001b). This arrangement results in octahedral arrangements of oxygen atoms in the tetrameric channel that provide a close approximation of the hydration sphere for K +  ions in solution and confer high selectivity for K +  over much smaller Na +  ions (Zhou et al., 2001b). Notably, the high resolution of the KcsA crystal structures shows that the selectivity filter represents a multi-ion pore with several K +  ion binding sites (S1 to S4 shown in Fig. 1.2C), two of which may be occupied at any time (i.e. S1 and S3, or S2 and S4) (Zhou et al., 2001b; for a review, see Roux, 2005). Integral to the structure and function of the selectivity filter are the interactions between the amino acid residues comprising the K +  channel signature sequence and residues elsewhere in the pore region. It has been suggested that hydrophobic interactions, hydrogen bonds, and/or van der Waals interactions between residues in the selectivity filter and the pore helix help to stabilize the selectivity filter in its conducting state (Kurata & Fedida, 2006). Of particular note, Figs. 1.2C and D show the interactions of two adjacent tryptophan residues in the pore helix (W67 and W68 of KcsA, W434 and W435 in Shaker) with the tyrosine (Y78 in KcsA, Y445 in 10 Shaker) and aspartate residues (KcsA D80; Shaker D447), respectively, of the selectivity filter. The tryptophan residues form a so-called “aromatic cuff” around the selectivity filter, the importance of which is highlighted by the effects of mutations that disrupt these interactions, including changes in selectivity, gating, and inactivation kinetics (Perozo et al., 1993; Heginbotham et al., 1994; Yang et al., 1997). The crystal structure of KcsA has also provided insight into the location of the Kv channel activation gate, which regulates the access of K +  ions to the conducting pathway in response to changes in voltage. Figure 1.2B shows that the M2 helices intersect near the intracellular face of the channel, creating what is called the bundle crossing, and closing off the aqueous central cavity of the pore from the cytoplasm. In contrast, the crystal structure of a prokaryotic Ca 2+ -activated K +  channel (MthK) has M2 helices that are splayed outwards from the central axis, resulting in a continuum between the aqueous central cavity of the channel and the intracellular medium (Jiang et al., 2002a; Jiang et al., 2002b). Comparisons of the two structures led to the conclusion that the KcsA structure represents the closed or resting conformation of the pore and the MthK structure represents an open pore, and that a conserved glycine residue (purple residues in Fig. 1.2A) in the inner helix contributes to activation gating by acting as a hinge by which the helices may swing open and closed (Jiang et al., 2002a; Jiang et al., 2002b; Kurata & Fedida, 2006). Eukaryotic K +  channels have an additional PxP motif (green residues in Fig. 1.2A) located distal to the glycine hinge that was proposed, based on studies using the substituted cysteine accessibility method (SCAM) with thiol-reactive methanethiosulfonate (MTS) reagents (del Camino et al., 2000; del Camino & Yellen, 2001), to introduce a sharp bend in the S6 helix and form part of the activation gate. This was subsequently confirmed in the crystal structure of Kv1.2 (Long et al., 2005a). It remains somewhat unclear why eukaryotic K + channels require this PxP motif, given that prokaryotic channels can activate without it and that 11 the absolute positions of the proline residues in the S6 helices are not critical (Labro et al., 2003). Regardless, it is evident from these findings that the activation gate of Kv channels is formed by the intersection of the C-terminal ends of the four S6 helices. 1.2.3.2 The voltage-sensing domain The transmembrane helices S1 – S4 comprise the voltage sensing domain of Kv channels (Fig. 1.1). Of these segments, S4 is considered to be the main voltage sensor, primarily due to the presence of basic residues (Arg or Lys) spaced at regular intervals through the helix. Neutralization (Aggarwal & Mackinnon, 1996; Seoh et al., 1996a) and SCAM studies (Larsson et al., 1996) have shown that the outermost arginine residues (R1 – R4) are most important for voltage sensing and move across the membrane with changes in voltage. Due to the hydrophobic nature of the lipid bilayer, it had long been held that the S1 – S3 segments of the voltage sensing domain serve to shield the positive S4 charges from the lipid, particularly since these segments tend to contain negatively charged residues (reviewed in Börjesson & Elinder, 2008). However, the crystal structures of the mammalian Kv1.2 channel (Long et al., 2005a; Long et al., 2005b), and of the bacterial KvAP channel (Jiang et al., 2003), suggest that the S4 segments are not completely protected by the S1 – S3 segments. Indeed, the crystal structure of a chimeric Kv1.2/Kv2.1 channel indicates that lipid molecules and the voltage sensing domain segments, including the charged S4, are in close contact (Long et al., 2007). The crystal structure of Kv1.2 is shown in Figure 1.3. What is immediately apparent is that the voltage sensing domain of one subunit is separated from its own pore domain by the S4- S5 linker, and is in fact in close approximation with the pore forming S5-S6 segments of an adjacent subunit. Unfortunately, due to its static nature, the structure itself cannot show precisely how the S4 voltage sensor moves in response to changes in the transmembrane voltage to initiate 12 opening of the activation gate. It does, however, provide a structural framework with which to describe the electromechanical process of channel activation, as detailed in the next section. 13    Figure 1.3. The crystal structure of the Kv1.2 channel. A, Side view of a ribbon representation of the Kv1.2 crystal structure. The four α-subunits, including the transmembrane (TM) voltage sensing and pore domains and the intracellular T1 domain formed by the N-termini are shown, each in a different colour. Also shown are accessory β-subunits. B, Side view of a single α-subunit and its accessory β-subunit. Labels correspond to the transmembrane helices S1 – S6, the PVP sequence in S6, and the N- and C-termini of the α- and β-subunits. C, A view of the Kv1.2 crystal structure from the extracellular side of the pore; α-subunits are colour-coded as in A. Note how the voltage sensing domain (S1-S4) of one subunit is in close proximity to the pore domain (S5-S6) of an adjacent subunit. From Long SB, Campbell EB & MacKinnon R (2005a). Crystal structure of a mammalian voltage-dependent Shaker family K +  channel. Science 309, 897-903. Reprinted with permission from AAAS. 14 1.2.4 An electromechanical view of activation and deactivation In response to membrane depolarization, Kv channels activate and the intracellular activation gate opens to allow conduction of K +  current. The reverse process of deactivation and channel closing occurs upon repolarization. The question of how this change in membrane potential is “sensed” and translated into channel opening is one that was first explained in the seminal work of Hodgkin and Huxley (1952; reviewed in Hille, 2001), in which they proposed that Kv channels had four charged “particles” which responded to voltage by moving into either a permissive (activated) or a non-permissive (deactivated) state. Channel opening required that all four “particles” be in the permissive state, the probability of which increased with depolarization. Conversely, if one or more “particles” was in a non-permissive state, the channel would be in one of several possible closed states. While we now know that these charged “particles” predicted by Hodgkin and Huxley correspond primarily to the gating charges of the S4 voltage sensors, the exact way in which they move to open the channel remains somewhat unclear. There have been three major models for S4 movement in response to voltage. These include: i) the helical-screw or sliding helix model, in which S4 rotates about its axis whilst also moving translationally across the membrane to transfer charges across the electric field (Durell et al., 1998; Keynes & Elinder, 1999); ii) the transporter-like model, which involves a mostly rotational movement of S4 (and perhaps a rearrangement of the membrane) to essentially transport S4 charges from one side of the membrane to the other (Papazian & Bezanilla, 1997); and, iii) the paddle model, initially based on the crystal structure of KvAP, in which the voltage sensor paddle, comprised of S4 and the C-terminal half of S3, forms a rigid structure that moves through the membrane and in close proximity to S1 and S2 (Jiang et al., 2003). There continues to be some debate over which model is most accurate. In recent years the models have begun to 15 converge, such that in all three, S4 rotates while moving 6 - 15Å across the membrane (for reviews, see Tombola et al., 2006; Börjesson & Elinder, 2008). Regardless of the exact nature of S4 movement with depolarization, that movement must somehow result in the opening of the intracellular activation gate, likely by inducing a conformational change near either the conserved glycine hinge or the kink introduced by the PxP motif in S6 (see above), which causes the internal ends of S6 to swing away from the central axis of the pore and open the bundle crossing. The Kv1.2 crystal structure shows that the pore domain of each subunit, specifically the region of S6 below the PxP motif, is associated with the S4-S5 linker of the same subunit (Fig. 1.3), suggesting that this linker couples S4 movement to pore opening. This is consistent with findings from several functional studies showing that specific interactions between the S4-S5 linker and the S6 segment are critical for voltage-dependent gating (Lu et al., 2001; Chen et al., 2001; Lu et al., 2002; Tristani-Firouzi et al., 2002; Ding & Horn, 2002; Ding & Horn, 2003; Labro et al., 2008; Nishizawa & Nishizawa, 2009). Thus, the movement of S4 may be indirectly coupled to the S6 activation gate by the S4-S5 linker. Based on these findings, it has been proposed that an outward movement of S4 with depolarization creates tension in the S4-S5 linker that is, in turn, transmitted to S6. This results in the C- terminal end of S6 pivoting, from either the glycine hinge or the PxP motif, outwards from the central axis of the pore, which is then open. Conversely, inward movement of S4 with hyperpolarization would tend to push the S4-S5 linker radially inward and exert a force on the internal end of S6 so that the activation gate closes. In summary, the elucidation of the crystal structures of a number of K +  channels, particularly the mammalian Kv1.2 channel, has greatly enhanced our knowledge of Kv channel function, particularly in the areas of ion permeation and activation/deactivation gating. 16 1.3 INACTIVATION OF KV CHANNELS Following activation and channel opening, and provided the depolarization is long enough, many Kv channels undergo inactivation, defined as entry into a stable, non-conducting state that is structurally distinct from the non-conducting deactivated or closed states. Relative to the processes of activation and deactivation, the mechanisms of Kv channel inactivation are still poorly understood. Under voltage-clamp conditions, the inactivation process typically manifests as a time-dependent decay of current. Inactivation is considerably more variable than activation and deactivation, both in terms of phenotypic properties and the underlying mechanisms. Traditionally, in Kv channels, two major types of inactivation are thought to occur from the open state: N-type inactivation and P/C-type or slow inactivation. Relatively recently, there have also been reports (see below) of inactivation occurring from partially activated closed-states. This section provides a description of the biophysical properties of the various types of inactivation in Kv channels. 1.3.1 N-type inactivation Of the different forms of inactivation, N-type inactivation is arguably the best characterized. N-type inactivation is often referred to as “fast” inactivation and generally occurs on the timescale of tens of milliseconds. The mechanism for fast inactivation was first described for NaV channels, when it was shown that inactivation was inhibited by intracellular perfusion with the proteolytic enzyme pronase (Bezanilla & Armstrong, 1977). This led to the proposal that fast inactivation involved a tethered “ball-and-chain” mechanism, whereby an intracellular inactivation “ball” particle tethered to the channel via a peptide “chain” induced inactivation by occluding the inner pore of the channel. Pronase removes inactivation by cleaving the inactivation “ball and chain” at one or more points. It was subsequently shown that in NaV 17 channels the inactivation “ball” is actually a cluster of hydrophobic residues (IFM) in the intracellular linker between domains III and IV (West et al., 1992). Thus, fast inactivation of NaV channels is now better described by a “hinged-lid” mechanism whereby the IFM cluster, via hydrophobic interactions, latches the domain III/IV linker into a pore blocking position (West et al., 1992). A “ball-and-chain” mechanism for the N-type inactivation of Kv channels was suggested by studies with Shaker showing that N-type inactivation could also be removed by internal perfusion with trypsin, another proteolytic enzyme (Hoshi et al., 1990). Mutational deletion of residues 6-46 from the N-terminal region also removed fast inactivation, indicative of a role of the intracellular N-terminus in inactivation (Hoshi et al., 1990); this mutant has since become known as the fast inactivation-removed Shaker (ShakerIR) channel. The integral role of the N- terminal regions was confirmed by a subsequent finding that fast inactivation could be restored to ShakerIR channels by internal perfusion of a peptide with a primary sequence similar to that of the deleted N-terminus (Zagotta et al., 1990), leading to the term N-type inactivation (Hoshi et al., 1991). Not all Kv channels undergo N-type inactivation; those that do include Shaker, Kv1.4 and some Kv channels when expressed with β-subunits (reviewed in Kurata & Fedida, 2006). A conserved mechanism and structural basis for N-type inactivation amongst Kv channels is suggested by the transferability of N-terminal peptides from one N-type inactivating channel to another, and by the ability of these N-terminal peptides to confer inactivation on non-N-type inactivating channels such as Kv1.1 (Murrell-Lagnado & Aldrich, 1993; Stephens & Robertson, 1995; Antz & Fakler, 1998). The variability in the primary sequences of N-terminal inactivation peptides and their transferability between Kv channels suggests that the receptor site has fairly general structural requirements for the inactivation peptide. Indeed, the basic structural requirements for the N-terminal inactivation peptide include: i) a short sequence of hydrophobic 18 residues to form the “ball”; and, ii) a sequence of variable length with a net positive charge to form the “chain” (Hoshi et al., 1990). The current model of N-type inactivation involves the movement of one of the four inactivation “balls” through lateral openings between the S1 segments of the α-subunits and the N-terminal T1 domains (Fig. 1.3; MacKinnon et al., 1993; Gulbis et al., 2000; Sokolova et al., 2001; Long et al., 2005a) to a binding site in the central cavity of the channel where it may occlude the pore (del Camino et al., 2000). The positive charges of the “chain” may position the inactivation “ball” near the S1-T1 linkers by virtue of electrostatic interactions with the negative residues of these linkers (Gulbis et al., 2000). Given that N-type inactivation involves occlusion of the inner pore, it is perhaps unsurprising that inactivation is strongly coupled to activation and the N-terminal inactivation peptide behaves essentially as an open channel blocker (Demo & Yellen, 1991; Zhou et al., 2001a). In this respect, N-type inactivation is competitively inhibited by intracellular quaternary ammonium compounds, such as tetraethylammonium (TEA), that block open Kv channels at a binding site in the central cavity (Choi et al., 1991; del Camino et al., 2000). Additionally, recovery from N-type inactivation upon repolarization is accelerated in elevated external [K + ]o (Demo & Yellen, 1991), consistent with a “knock-off” effect of the inward K+ current on the inactivation particle. With repolarization, recovery from inactivation must occur prior to channel deactivation, resulting in a delay of the return of the S4 gating charges (referred to as charge immobilization) and a brief opening of the channel during recovery; this is likely due to the presence of the N-terminal “chain” that prevents closure of the activation gate (Demo & Yellen, 1991). 1.3.2 Slow or P/C-type inactivation Removal of N-type inactivation uncovers a second inactivation process in ShakerIR (Hoshi et al., 1990). Based on the initial finding with ShakerIR C-terminal splice variants that 19 the time course of this inactivation was dependent on the nature of residue 463 (i.e., either an Ala or a Val residue) in S6 at the level of the central cavity, the term “C-type inactivation” was coined (Hoshi et al., 1991). Shortly thereafter, it was found that mutations in the outer pore region of Kv2.1 and/or a Kv2.1/Kv3.1 chimeric channel, particularly at position 369 at the base of the pore helix in Kv2.1, induced an inactivation process with characteristics that appeared distinct from both N-type and C-type inactivation (De Biasi et al., 1993). Thus, the term “P-type inactivation” was introduced, to emphasize the involvement of the outer pore and to distinguish it from C-type inactivation. In the years since, it has been shown that C-type inactivation is complicated and likely involves conformational changes in the outer pore and selectivity filter (see below). Consequently, the definitions of P- and C-type inactivation have evolved and the current view is that P-type inactivation describes an initial inactivation process involving the outer pore and selectivity filter, while C-type inactivation refers to a stabilization of this inactivated conformation and immobilization of the S4 gating charges in an activated-inactivated state (Olcese et al., 1997; e.g. Loots & Isacoff, 1998). However, these operational definitions of P- and C-type inactivation remain ambiguous, and the terms are now often conflated into “P/C- type inactivation”. Because P/C-type inactivation typically occurs on a slower (100 – 1000 ms) timescale than N-type inactivation (although see Hoshi et al., 1991), the term “slow inactivation” has also been used. The term “slow inactivation” is used here, to avoid the sometimes confusing aspects of the P/C-type inactivation terminology. 1.3.2.1 Slow inactivation involves the outer pore region After its first description in ShakerIR channels (Hoshi et al., 1991), several findings implied an important role for the outer pore region in slow inactivation. Unlike N-type inactivation, slow inactivation was not inhibited by internal TEA (Choi et al., 1991). Instead, the onset of slow inactivation in ShakerIR and other channels was inhibited by externally applied 20 TEA (Grissmer & Cahalan, 1989;  Choi et al., 1991) and by elevated [K + ]o (Hoshi et al., 1990; López-Barneo et al., 1993). Additionally, it was found that the Shaker T449 residue near the outer mouth of the selectivity filter (see Fig. 1.2A) played a critical role in the rate and [K + ]o- dependence of slow inactivation (López-Barneo et al., 1993). As detailed below, these features all point to an involvement of the outer pore in slow inactivation. The inhibitory effects of external TEA on the slow inactivation of ShakerIR and other channels (Grissmer & Cahalan, 1989; Choi et al., 1991; Molina et al., 1997) has been likened to a “foot-in-the-door” effect, whereby the presence of TEA in an outer pore binding site inhibits the conformational changes underlying slow inactivation. Mutational studies have suggested that the putative external TEA binding site is near residue T449 located near the outer mouth of the selectivity filter (MacKinnon & Yellen, 1990; Heginbotham & MacKinnon, 1992; Molina et al., 1997). The sensitivity of TEA binding to the residue at position 449 (or its equivalent) likely explains why some channels (e.g. human (h)Kv1.4, K531) do not exhibit TEA-sensitive slow inactivation (Molina et al., 1997). The inhibitory effects of elevated [K + ]o on the rate of slow inactivation has also been attributed to a similar mechanism of action. It was initially suggested that the slow inactivation of ShakerIR was relatively insensitive to elevations in [K + ]o (López- Barneo et al., 1993). However, subsequent findings indicated that this was an artifact resulting from the effects of K +  accumulation in the outer pore mouth due to outward K +  flux, such that the effective [K + ]o during a depolarizing pulse was ≈15 mM (Baukrowitz & Yellen, 1995). Prevention of this accumulation, either with specialized perfusion or voltage protocols, or by the presence of an intact N-type inactivation mechanism, significantly enhanced the [K + ]o sensitivity of ShakerIR and wild-type (wt) Shaker slow inactivation (Baukrowitz & Yellen, 1995). Like external TEA block, and consistent with a role of the outer pore mouth, the inhibitory effects of K + o on ShakerIR slow inactivation are very sensitive to mutations at position 21 449 (López-Barneo et al., 1993). Intriguingly, the sensitivity of T449 mutants to [K + ]o was correlated with the effects of the mutation on the slow inactivation rate, further highlighting the importance of the residue at this position to slow inactivation. Specifically, mutation of T449 to Glu (E), Lys (K), Ala (A), Gln (Q) or Ser (S) drastically accelerated slow inactivation, and the time constants for slow inactivation of the T449E, -K, and -A mutants also exhibited an enhanced sensitivity to elevated [K + ]o (López-Barneo et al., 1993; Schlief et al., 1996). Conversely, mutations to a His (H), Val (V), or Tyr (Y) resulted in a slowing or removal of slow inactivation and a decreased sensitivity to changes of [K + ]o (López-Barneo et al., 1993; Schlief et al., 1996). The T449H mutation also introduced an enhanced sensitivity of slow inactivation to extracellular acidification and, at low pH, an inhibitory effect of K + o on slow inactivation was restored (López-Barneo et al., 1993). Among the Shaker-related Kv1 channels, the identity of the residue at the position equivalent to Shaker T449 is not well conserved (Fig. 1.2A) and the slow inactivation properties of these channels do not always match those of the corresponding ShakerIR T449 mutant (reviewed in Kurata & Fedida, 2006). For example, Kv1.4 has a lysine at this position, but slow inactivates at a rate >20 slower than that of the fast-inactivating ShakerIR T449K mutant (López- Barneo et al., 1993; Rasmusson et al., 1995). However, the importance of residues at the position homologous to Shaker T449 in the slow inactivation of Kv1 channels is suggested by the ability of mutations at this position to modulate other manoeuvres that affect slow inactivation. For example, the enhancement of the slow inactivation rate of hKv1.5 by external H +  or Ni 2+  ions (reviewed in detail below) is prevented by the R487V (analogous to T449V) mutation, which has minimal effect on slow inactivation under control conditions (Fedida et al., 1999). 22 1.3.2.2 The role of the selectivity filter The ability of external K +  to inhibit slow inactivation via a proposed foot-in-the-door mechanism raised the possibility of an involvement of the selectivity filter in the conformational change underlying slow inactivation. This was reinforced by the finding that the ability of external monovalent cations to inhibit slow inactivation was dependent on their relative permeability through Kv channels (Pardo et al., 1992; López-Barneo et al., 1993; Fedida et al., 1999). This is consistent with the idea that the occupancy of one or more binding sites within the selectivity filter impedes slow inactivation. In a series of studies, Baukrowitz and Yellen (1995; 1996) showed that slow inactivation of Shaker depends on the occupancy of a highly selective K +  binding site that is likely to be near the external end of the selectivity filter, such that the rate of slow inactivation is correlated with the exit rate of the last ion from the selectivity filter. Additionally, it was shown that slow inactivation can be enhanced by use-dependent intracellular blockers or N-type inactivation, which prevents refilling of the selectivity filter from the intracellular milieu and facilitates emptying of the K +  binding site (Baukrowitz & Yellen, 1996). This also agrees with the aforementioned inhibitory effects of elevated [K + ]o and external TEA, which would reduce the egress of K +  from the pore and/or the probability of a prolonged vacancy of the binding site. Further support for a role of the selectivity filter in slow inactivation comes from the observation of changes in the Na +  permeability of slow inactivated channels such that appreciable Na +  currents through Shaker or related channels may be recorded in the absence of K +  (Starkus et al., 1997; Kiss & Korn, 1998; Kiss et al., 1999; Ogielska & Aldrich, 1999; Wang et al., 2000). The ability of slow inactivated channels to conduct Na +  ions argues against a complete collapse or closure of the selectivity filter and is instead supportive of conformational changes that sustain the Na +  permeability of the selectivity filter whilst preventing K +  conduction 23 (Starkus et al., 1997). Based on the similarities between the inhibitory effects of external K + on the rate of slow inactivation and on the Na +  current through inactivated channels, it was inferred that the K +  binding site responsible for modulating the rate of inactivation reported in earlier studies (see above; e.g. Baukrowitz & Yellen, 1996) also resides within the selectivity filter (Kiss & Korn, 1998; Ogielska & Aldrich, 1999). Various mutations near the selectivity filter modulate inactivation, with the Shaker W434F mutation in the pore helix being one of the most notable. The W434F mutation confers permanent (or exceedingly rapid) slow inactivation on Shaker (Yang et al., 1997) such that Na +  currents like those observed in the slow inactivated wild-type channel can be recorded (Starkus et al., 1998). The homologous W472F mutation has the same effect in hKv1.5 (Chen et al., 1997; Wang & Fedida, 2001). Shaker W434 is the positional equivalent of W67 in KcsA and together with W435 forms the “aromatic cuff” (Fig. 1.2) that is proposed to help stabilize the selectivity filter in a conducting conformation (Doyle et al., 1998). Conceivably, disruption of the hydrogen bond network around the selectivity filter facilitates the partial pore collapse underlying slow inactivation. Intriguingly, the T449V mutation, which inhibits slow inactivation (López-Barneo et al., 1993), is able to “rescue” Shaker W434F current such that the W434F-T449V double mutant is conducting (Kitaguchi et al., 2004). This supports the conclusion that the non- conducting W434F mutant represents a permanently (or rapidly) slow inactivated channel and underscores the importance of interactions between the selectivity filter, the pore helix and the outer mouth of the pore in slow inactivation. Finally, recent studies in KcsA have suggested that an interaction between E71 in the pore helix and D80 of the GYGD sequence destabilizes the selectivity filter (Fig. 1.2) and promotes a constriction of the selectivity filter and slow inactivation (Cordero-Morales et al., 2006; Cordero-Morales et al., 2007). The V370E mutation in Kv1.2 (homologous position to 24 KcsA E71) enhances slow inactivation, purportedly by interacting with D379 (equivalent to KcsA D80), leading to the proposal that a similar mechanism may be possible (Cordero-Morales et al., 2007). However, given that eukaryotic channels have a conserved valine residue at the position equivalent to E71, whether this interaction is functionally significant in wild-type eukaryotic channels remains questionable. 1.3.2.3 Slow inactivation involves a localized conformational change in the selectivity filter While it is clear that slow inactivation results in a loss of K +  conductance, the ability of inactivated channels to conduct Na + , as mentioned above, precludes a complete collapse of the selectivity filter and/or outer pore. However, several factors are suggestive of a localized constriction or conformational change of the selectivity filter as the underlying mechanism of slow inactivation. For example, Yellen et al. (1994) showed that substitution of a cysteine residue (T449C) at the outer aspect of the ShakerIR selectivity filter resulted in a strongly state- dependent sensitivity to external Cd 2+  and Zn 2+  ions. Relative to the open state, the slow inactivated state of ShakerIR T449C demonstrated a 45,000-fold increase in affinity for Cd 2+ , implying a structural change in this region that increased the external accessibility of and/or decreased the distance between the T449C residues, allowing for co-ordination of the Cd 2+  by the cysteine residues (Yellen et al., 1994). In a follow up to the above study, it was found that the accessibility of substituted cysteine residues at positions M448, T449 and P450 in Shaker to modification by MTS reagents was also state-dependent, such that modification was much faster in the slow inactivated state (Liu et al., 1996). Furthermore, inter-subunit bridging and disulfide cross-linking between M448C residues was demonstrated with a thiol-reactive compound and an oxidizing reagent, respectively, and this reactivity was also strongly accelerated in the slow inactivated state (Liu et al., 1996). A correlation between the slow inactivation rate in T449 mutants and chloramine-T oxidation of M448C has also been reported (Schlief et al., 1996). The 25 potentially close approximation of intersubunit T449 or M448 residues during slow inactivation is consistent with the hypothesis that the changes in the outer pore occur in a highly cooperative manner (Ogielska et al., 1995; Panyi et al., 1995; Ogielska & Aldrich, 1999). Conformational changes in the outer pore region of Shaker have also been indirectly observed with voltage-clamp fluorimetry studies, whereby thiol-reactive fluorescent dyes are used to tag cysteine residues introduced at specific external sites on the channel. Any changes in the immediate environment of the dye, i.e. due to conformational changes, are reflected as changes in the fluorescence emission. Upon depolarization, fluorescent tags at T449C, S424C in the turret, and other regions of the outer pore in Shaker show changes in emission with kinetics that parallel that of the ionic current decay associated with slow inactivation (Cha & Bezanilla, 1997; Loots & Isacoff, 1998; Loots & Isacoff, 2000; Gandhi et al., 2000). While it can sometimes be difficult to discern whether the conformational changes underlying these changes in fluorescence emission are occurring at the site of the fluorophore or if they are due to the indirect effects of conformational changes elsewhere, it is clear that the outer pore mouth experiences environmental alterations in response to slow inactivation. Further evidence of the localized nature of the outer pore constriction is supplied by various studies showing ion-trapping within the pore of inactivated channels. Accessibility studies with externally applied Ba 2+  ions and ShakerIR channels have shown that slow inactivation inhibits the access of Ba 2+  to the deeper of two binding sites within the selectivity filter and, alternatively, that Ba 2+  already bound at this site can become trapped by slow inactivation (Basso et al., 1998; Harris et al., 1998). On a related note, work by Ogielska and Aldrich (1999) showed that in a ShakerIR A463C mutant, slow inactivation can proceed with a K +  ion bound at a deep binding site in the selectivity filter, perhaps akin to the deep Ba 2+  binding site. Conversely, K +  occupancy of a more superficial site inhibited slow inactivation, consistent 26 with the earlier findings of Baukrowitz and Yellen (1996) and the hypothesis that the active site for K +  inhibition of slow inactivation lies in the outer region of the selectivity filter (Ogielska & Aldrich, 1999). Similarly, it has been suggested that a K +  (or other permeant ion) can become trapped within the central cavity of slow inactivated Kv1.3 channels upon hyperpolarization, and the subsequent movement of this ion into the selectivity filter aids recovery from slow inactivation (Ray & Deutsch, 2006). Together, the ability of permeant ions to reside within the selectivity filter of slow inactivated channels points to a localized constriction or conformational change, likely near the outer mouth of the selectivity filter. 1.3.2.4 The voltage sensor may be involved in slow inactivation There has been a long-standing association between slow inactivation and the S4 voltage sensors. Indeed, like fast inactivation in NaV channels (see above), one of the classical descriptors for slow inactivation is an apparent “immobilization” of S4 gating charge following prolonged depolarization and slow inactivation in ShakerIR and Kv1.5 (Olcese et al., 1997; Chen et al., 1997). In other words, gating current measurements revealed that the amount of gating charge returned upon repolarization following slow inactivation appeared to be significantly less than that moved with the initial depolarizing step. In actuality, charge immobilization represents a left-shift or hysteresis of the voltage-dependence of S4 movement following prolonged depolarization and/or slow inactivation, and likely reflects a change of the energetic landscape for voltage sensor movement (Olcese et al., 1997). The similar time courses for the recovery from charge immobilization and the recovery of ionic current following slow inactivation further strengthened the association between the voltage sensors and slow inactivation (Olcese et al., 1997). Additionally, as with slow inactivation, charge immobilization was found to be sensitive to the presence of permeant ions and the channel blocker 4-aminopyridine (4-AP) and correlated 27 with the development of Na +  tail currents (Fedida et al., 1996; Chen et al., 1997; Wang & Fedida, 2001). A link between voltage sensor movement and slow inactivation was also supported by voltage-clamp fluorimetry studies in Shaker, which were suggestive of an interaction between the top of S4 and the outer pore region (Loots & Isacoff, 1998; Loots & Isacoff, 2000; Gandhi et al., 2000). Specifically, it was observed that following slow inactivation, fluorophores attached at S424C (and other regions) in the turret were able to report on fast movements of the voltage sensor that were otherwise undetected at this site in open, non-inactivated channels (Loots & Isacoff, 1998; Loots & Isacoff, 2000). Conversely, fluorophores attached to A359C at the top of the S4 segment reported on two phases of movement: a rapid component associated with S4 activation upon depolarization and a slower component with kinetics matching those of slow inactivation (Loots & Isacoff, 1998). The exact nature of any interactions between the voltage sensing and pore domains during slow inactivation remains unknown. Based on studies examining cysteine accessibility and disulfide bond formation, a conserved glutamate residue (E418 in Shaker) at the top of S5 has been implicated in electrostatic interactions with G452 in the P-S6 linker (Elinder & Arhem, 1999; Larsson & Elinder, 2000; Elinder et al., 2001). It has been proposed that upon depolarization the voltage sensor exerts an allosteric effect on S6, causing the P-S6 linker to rotate and resulting in a breaking of the hydrogen bonds between E418 and G452 that stabilize the conducting conformation of the pore, allowing slow inactivation to occur (Larsson & Elinder, 2000; Elinder et al., 2001). Although charge immobilization and S4-pore interactions appear to go hand-in-hand with slow inactivation, there is also evidence to suggest that the coupling between the voltage sensor and slow inactivation may not be absolute. One example is the recently cloned voltage- 28 dependent phosphatase from Ciona intestinalis (Ci-VSP), which lacks a pore domain and hence, cannot undergo outer pore rearrangements associated with slow inactivation, yet it still exhibits charge immobilization in response to depolarization (Villalba-Galea et al., 2008). Conversely, the non-conducting ShakerIR W434F mutant, which is considered permanently inactivated, does not exhibit charge immobilization unless subjected to a prolonged depolarizing pulse (Olcese et al., 1997; Yang et al., 1997). These findings raise the question, therefore, of whether charge immobilization is an obligatory component of slow inactivation or if it reflects an independent and inherent property of voltage sensors at depolarized holding potentials. Villalba-Galea and colleagues (2008) have suggested the latter, arguing that charge immobilization reflects an inherent movement of S4 voltage sensors from an unstable activated state to a “relaxed” state. A recent study showing that charge immobilization is related to an interaction between the S4-S5 linker and S6 that tends to stabilize the open state, supports the proposal that immobilization is not a consequence of slow inactivation (Batulan et al., 2010). In summary, removal of N-type inactivation uncovers a slow inactivation process that involves a localized constriction near the outer mouth of the selectivity filter and/or outer pore region of Kv channels. Slow inactivation is a complex process, as evidenced by the continued uncertainty over the precise molecular machinery initiating and underlying the constriction of the selectivity filter, and whether or not it involves a contribution from the voltage sensors. 1.3.3 U-type inactivation A number of Kv channels exhibit inactivation properties that differ from those typical of N-type or slow inactivation. For example, the inactivation of Kv2.1 and Kv3.1 shows sensitivities to external TEA and K +  that are opposite to that of slow inactivating channels such as Shaker; that is, increases in the [TEA]o or [K + ]o actually accelerate inactivation (Klemic et al., 1998; Klemic et al., 2001) A similar behaviour has also been observed in an N-terminal 29 truncated form of Kv1.5, even though the full-length channel shows properties generally consistent with those ascribed to slow inactivation (Kurata et al., 2001; Kurata et al., 2002; Kurata et al., 2005). An additional characteristic of this inactivation process, from which it derives its name, is the “U-shape” of the inactivation-voltage relationship, whereby inactivation is more rapid and complete at intermediate voltages (e.g. 0 mV) associated with significant dwell times in partially activated closed states (Klemic et al., 1998; Klemic et al., 2001). This also accounts for the observation that, relative to a single prolonged depolarization, U-type inactivation is enhanced by short, repetitive voltage steps that cause the channel to spend proportionally more time in partially activated states (Klemic et al., 2001). In contrast, slow inactivation as described for Shaker and related channels is typically fastest and most complete at high voltages (e.g. +50 mV) associated with fully activated and open channels (Hoshi et al., 1991; Kurata et al., 2001), such that slow inactivation is thought to be strongly coupled to activation and the open state (Olcese et al., 1997). U-type inactivation is still fairly poorly understood, and it remains unclear whether it represents a variant of slow inactivation. It has recently been suggested that the tendency of ShakerIR channels expressed in Xenopus oocytes to undergo classical slow inactivation or U-type inactivation depends on whether voltage-clamp recordings are made from excised patches or intact cells, and that both slow inactivation and U- type inactivation, with the latter tentatively proposed to involve closure of the activation gate despite sustained depolarization, can contribute to current decay (González-Pérez et al., 2008). 1.3.4 Preferential closed-state inactivation of Kv4 channels Although N-terminal deletion in Kv4.1 and Kv4.2 channels can slow or abolish the initial fast component of inactivation (Jerng & Covarrubias, 1997; Bähring et al., 2001; Barghaan et al., 2008), this fast inactivation does not share the typical features of N-type inactivation such as inhibition by internal TEA or a dependency on basic residues in the N-terminus (Jerng & 30 Covarrubias, 1997; Gebauer et al., 2004; for a review see Jerng et al., 2004). Interestingly, C- terminal deletions in Kv4.1 can mimic the effects of N-terminal deletion, suggestive of a concerted action between the N- and C-termini that evokes the fast phase of inactivation (Jerng & Covarrubias, 1997). A similar process has not been described in Shaker. It is currently thought that, in the absence of accessory subunits, the N-terminus of Kv4 channels mediates unstable open-state inactivation via occlusion of the intracellular pore mouth, perhaps facilitated by interactions with the C-terminus (Jerng et al., 2004; Covarrubias et al., 2008). There are also several major differences between the classical model of slow inactivation for Shaker and the inactivation properties of Kv4 channels. For example, in ShakerIR and, to a lesser extent, the related Kv1 channels, the presence of a valine residue at position T449 (or its equivalent), tends to inhibit slow inactivation (López-Barneo et al., 1993; Jäger et al., 1998). Kv4 channels have a native valine residue at the position homologous to Shaker T449, yet still undergo robust inactivation. Additionally, as with U-type inactivation, high external [K + ] or [TEA] accelerate inactivation and inhibit recovery from inactivation in Kv4 channels (Jerng & Covarrubias, 1997; Shahidullah & Covarrubias, 2003; Kaulin et al., 2008). While the exact mechanism by which Kv4 channels inactivate remains unclear, it has become evident that a considerable proportion of channels are able to inactivate from partially activated or pre-open closed states with modest depolarizations (e.g. from −80 to −40 mV) (Jerng et al., 1999; Bähring et al., 2001; Beck & Covarrubias, 2001; Wang et al., 2005; Skerritt & Campbell, 2007). Importantly, in Kv4 channels the weakly voltage-dependent concerted opening step is assumed to be strongly reverse-biased, in contrast to Kv1 channels such as Shaker (Jerng et al., 1999; Bähring et al., 2001; Beck & Covarrubias, 2001; Shahidullah & Covarrubias, 2003; Patel et al., 2004; Wang et al., 2005; Skerritt & Campbell, 2007; Dougherty et al., 2008; Kaulin et al., 2008). Given the unstable nature of the open state and that inactivation 31 from the closed state is absorbing, with a prolonged depolarization channels eventually accumulate in the closed inactivated state, leading to the term preferential closed-state inactivation (Covarrubias et al., 2008; Dougherty et al., 2008; Kaulin et al., 2008). Under conditions of greater membrane depolarization (i.e. positive to −30 mV), the increased open probability (Po) allows for significant inactivation from the open state. The popular conception is that the open-inactivated state is unstable and can only recover through the open state. This gating behaviour, coupled with the reverse-biased opening transition, leads to increased residency of the inactivation-permissive pre-open closed state and preferential closed-state inactivation (Dougherty et al., 2008; Kaulin et al., 2008). It should be noted that alternative models have been proposed where open-state inactivation is prominent and absorbing at sufficiently depolarized potentials (Patel et al., 2004; Skerritt & Campbell, 2007) and, in some cases, where direct transitions between the open- and closed-inactivated states are not excluded (Wang et al., 2005). Although the mechanistic basis is still poorly understood, Kv4 channels clearly undergo inactivation from one or more closed states. Recent gating current measurements from Kv4.2 channels are suggestive of significant charge immobilization related to inactivation at hyperpolarized potentials (Dougherty et al., 2008), leading to a proposal that Kv4 channels inactivate via a mechanism analogous to that proposed for closed-state inactivation of sea urchin HCN channels (Covarrubias et al., 2008; Dougherty et al., 2008; Kaulin et al., 2008). That is, inactivation involves an uncoupling or “slippage” of the mechanical linkage between the voltage sensors and the internal activation gate, such that the S4 segments can be immobilized in the activated position while the activation gate remains closed (Shin et al., 2004). 32 1.4 RESTING INACTIVATION AND SCOPE OF THE THESIS In this Chapter, the slow inactivation of Shaker and related Kv1 family members has been reviewed and shown to occur during a prolonged depolarization and to involve a localized constriction of the outer pore. In contrast to U-type and preferential closed-state inactivation, where inactivation occurs from partially activated closed states in the activation pathway, slow inactivation is considered to be strongly coupled to channel activation and opening. However, several findings suggest that decreases in channel availability caused by the effects of external H +  on wt Kv1.5 and ShakerIR channels and/or by the modulatory effects of external K +  on ShakerIR T449 mutants and Kv1.5 H463G mutants, may be associated with inactivation occurring from the fully closed, or resting, state. As detailed below, the examination of this apparent resting inactivation, and whether it represents slow inactivation that has been uncoupled from activation, forms the overarching theme of this dissertation. 1.4.1 The effects of low pH on Kv1.5 and ShakerIR The Kv1.5 channel, a mammalian homologue of Shaker, underlies the ultrarapid delayed rectifier current (IKur) in human atrial myocytes (Fedida et al., 1993) and undergoes slow but not fast inactivation. We (Kehl et al., 2002; Fedida et al., 2005; Claydon et al., 2007) and others (López-Barneo et al., 1993; Pérez-Cornejo, 1999; Steidl & Yool, 1999; Jäger & Grissmer, 2001; Starkus et al., 2003) have shown that low extracellular pH increases the rate and the extent of Kv1.5 and ShakerIR current decay during a depolarizing pulse. Although the pKa for this effect differs by over 1 pH unit between Kv1.5 and ShakerIR, it has nonetheless been attributed in both channel types to an enhancement of the slow inactivation process. Acidification also causes a marked reduction of peak Kv1.5 current, which was attributed by Steidl and Yool (1999) to an activity-dependent accumulation of channels in the slow inactivated state. However, we found 33 that the decline of the peak macroscopic current persisted at very low stimulus frequencies where cumulative slow inactivation was precluded (Kehl et al., 2002). In single channel recordings, this effect manifests as a concentration-dependent increase of the number of null sweeps during step depolarizations (Kwan et al., 2004; Kwan et al., 2006; Claydon et al., 2007). Fluorescence measurements from ShakerIR channels have also provided evidence that protons induce inactivation of channels from closed, resting states (Claydon et al., 2007). Thus, we proposed that in ShakerIR and Kv1.5 the reduction of the peak current amplitude arises, at least in part, from a H + -induced decrease of channel availability at −80 mV, which is defined here as resting inactivation. An unresolved issue is whether the two effects of H + , namely the apparent enhancement of slow inactivation and the induction of resting inactivation, are mechanistically related and whether either effect represents modulation of the slow inactivation process observed with depolarization at pH 7.4. A number of previous findings imply a possible mechanistic link between the slow inactivation process and the effects of low pH. For example, as with slow inactivation in ShakerIR (López-Barneo et al., 1993; Baukrowitz & Yellen, 1995; Kiss & Korn, 1998), the resting inactivation induced by low pH is antagonized by elevated [K + ]o (Kd = 1 mM; Jäger & Grissmer, 2001; Kehl et al., 2002). Additionally, mutation of a specific outer pore residue (Kv1.5 R487 or ShakerIR T449) to a valine or tyrosine, which is implicated in the modulation of slow inactivation, markedly attenuates the enhanced current decay and resting inactivation observed at low pH (López-Barneo et al., 1993; Kehl et al., 2002; Trapani & Korn, 2003; Starkus et al., 2003; Claydon et al., 2007). Although in ShakerIR the site of action for the enhanced slow inactivation observed at low pH has been suggested to be a conserved aspartate residue in the GYGD sequence of the selectivity filter (Pérez-Cornejo, 1999; Starkus et al., 2003), in Kv1.5 the pH sensitivity is dramatically reduced by the mutation to glutamine of the 34 H463 residue in the turret of each α-subunit (Steidl & Yool, 1999; Kehl et al., 2002). Based on these findings, and the fact that Ni 2+ , another histidine ligand, qualitatively reproduces most of the effects of extracellular acidification (Perchenet & Clément-Chomienne, 2001; Kwan et al., 2004), we have proposed that H +  or Ni 2+  binding at one or more H463 residues in Kv1.5 facilitates, at rest, a K + o-sensitive process involving the outer pore and which might be related to slow inactivation (Kehl et al., 2002; Kwan et al., 2004). 1.4.2 The [K+]o-dependent loss of availability of fast-inactivating ShakerIR T449 and Kv1.5 H463G mutants As mentioned above, amino acid substitutions at position 449 in ShakerIR can have a major effect on the time course of slow inactivation. In the ShakerIR T449A, -E or -K mutants, slow inactivation is dramatically accelerated and, despite an increase of the driving force for outward K +  movement, there is a large decrease of the peak current when the [K + ]o is reduced from 140 to 2 mM (López-Barneo et al., 1993). To account for these findings, it was suggested that external K +  exerted a foot-in-the-door effect to antagonize slow inactivation and, secondly, that the mutations enhanced a (closed state) resting inactivation process that shared, at least qualitatively, the [K + ]o-sensitivity of the slow inactivation observed during a depolarizing pulse (López-Barneo et al., 1993). Interestingly, a reduction of macroscopic current amplitude is also observed in 0 K + o in wt Kv1.4 channels, which have a Lys residue at the position equivalent to Shaker T449, and it has also been suggested that this may be due to a decrease in channel availability associated with resting inactivation (Pardo et al., 1992; Jäger et al., 1998). However, the properties of the putative K + -dependent resting inactivation, either in ShakerIR or Kv1.4 channels, has not been examined in detail. In the investigation of the H + -induced reduction of the maximal conductance of Kv1.5 channels, it was found that mutations of residue H463 in the outer turret had profound effects on 35 slow inactivation (Kehl et al., 2002). While the H463Q mutation inhibited the effects of low pH, it was found that the H463G mutant exhibited accelerated slow inactivation at pH 7.4 (Kehl et al., 2002). Additionally, and reminiscent of the properties of the ShakerIR T449 mutants described above, the availability and consequently the peak conductance of H463G mutant channels was highly sensitive to the removal of K + o such that outward K +  current was virtually eliminated in 0 K + o (Kehl et al., 2002; Zhang et al., 2005). However, in K + -free solutions, robust Na +  tail currents with properties similar to those observed through slow inactivated channels could be observed (Zhang et al., 2005). These results imply that the loss of Kv1.5 H463G availability in 0 K + o may also be due to a resting inactivation process. 1.4.3 Objectives The results of the previous studies described above suggest that the activity-independent decrease of peak current amplitude of wt and mutant Kv1.5, ShakerIR and Kv1.4 channels at low pH or in low [K + ]o might be due a loss of channel availability associated with resting inactivation. However, it is unclear from these studies whether the mechanism underlying the H + -, Ni 2+ - or low-[K + ]o-mediated loss of channel availability at rest actually involves an outer pore inactivation process analogous to that of slow inactivation and, thus, whether the putative resting inactivation truly represents a form of slow inactivation occurring from the fully closed state. Consequently, the main aim of the experiments presented here was to elucidate whether slow inactivation can occur at −80 mV (i.e. resting inactivation) in Kv1.5 and ShakerIR channels under conditions of low pH, external Ni 2+ , or low [K + ]o. In the first study (Chapter 2), we pose the question of whether the decrease in peak macroscopic current and channel availability exhibited by Kv1.5 at low external pH is associated with constriction of the selectivity filter. To do this, externally applied Ba 2+  ions were used to probe the outer pore under resting (−80 mV) conditions at pH 7.4 and pH 5.5. Following the characterization of Ba 2+  binding to Kv1.5 36 channels, which has not been reported before, we show that Ba 2+  movement to and from a high affinity site deep within the resting channel is inhibited by low pH. This finding is consistent with the previously described properties of slow inactivation in ShakerIR (Basso et al., 1998; Harris et al., 1998), and suggests that in Kv1.5 external acidification induces constriction of the selectivity filter at rest that is perhaps similar to that which occurs during slow inactivation. In the second series of experiments (Chapter 3), a comparative analysis of the effects of low pH and external Ni 2+  ions on the kinetics of slow inactivation at +50 mV and resting inactivation at −80 mV was performed. These experiments were predicated on the hypothesis that if slow inactivation and resting inactivation involved the same mechanism (e.g. constriction of the selectivity filter), then the recovery kinetics following either process would be similar. Through the use of fast perfusion changes, we confirm that the accelerated current decay at +50 mV in low pH or external Ni 2+  is due to an enhancement of the slow inactivation process. By monitoring the time course of the decrease in peak macroscopic current, a robust and unequivocal resting inactivation of Kv1.5 induced by external H +  and Ni 2+  is also confirmed. The similarities of the time constants for recovery from enhanced slow inactivation and resting inactivation suggests that similar recovery pathways and mechanistic processes are involved. Using methodologies similar to those applied in Chapter 3, the concluding study (Chapter 4) examines whether resting inactivation occurs in ShakerIR and the fast-inactivating ShakerIR and Kv1.5 mutants. First, the effects of low pH on ShakerIR were assessed. It was found that, as in Kv1.5, the enhancement of slow inactivation in ShakerIR cannot account for all of the reduction in peak current, and that external H +  may also promote resting inactivation. The kinetics of the onset and recovery from the loss of availability of the ShakerIR T449 and Kv1.5 H463G fast-inactivating mutants in 0 K + o were also examined. From these experiments, the time course for recovery from resting inactivation induced by 0 K + o was identical to that for recovery 37 from depolarization-induced slow inactivation, which is strongly suggestive of a link between inactivation at +50 mV and −80 mV.  38 2 External Ba2+ block of human Kv1.5 at neutral and acidic pH: evidence for H + -induced constriction of the outer pore mouth at rest 2.1 INTRODUCTION As detailed in Chapter 1, several factors point to a H + -induced loss of Kv1.5 channel availability at −80 mV, and we propose that this could be due to a resting inactivation process. However, whether the H + -induced resting inactivation involves the same conformational changes of the selectivity filter and outer pore that occur during slow inactivation remains unknown. In this chapter, we have used externally applied Ba 2+  as a molecular probe to examine possible changes in the accessibility of the outer pore of Kv1.5 under acidic conditions. A Ba 2+  ion has almost the same crystal radius as a K +  ion (1.15 Å vs. 1.13 Å, respectively) and is able to enter the pore of several classes of K +  channels from either side of the membrane (Armstrong et al., 1982; Neyton & Miller, 1988a; Neyton & Miller, 1988b; Hurst et al., 1995; Harris et al., 1998; Somodi et al., 2004; Gibor et al., 2007). Regardless of the side of the channel to which it is applied, Ba 2+  binding is thought to occur within the selectivity filter (Neyton & Miller, 1988b; Hurst et al., 1995; Jiang & MacKinnon, 2000) and, perhaps because of its greater charge, Ba 2+  tends to have a long mean dwell time (relative to K + ), resulting in a block of K +  current. In ShakerIR channels, Ba 2+  has been shown to bind to two sequential sites within the pore: a low affinity superficial site that gives rise to a near-instantaneous component of block and a deep site with a higher affinity for Ba 2+  that is associated with a comparatively much slower block onset and offset (Hurst et al., 1995). It has been shown that the conformational change in the outer pore associated with slow inactivation reduces the ability of externally applied Ba 2+  to reach this deep binding site and, conversely, Ba 2+  already bound can become trapped at this site with slow inactivation (Basso et al., 1998; Harris et al., 1998). 39 Here, we first characterize the block of Kv1.5 by externally applied Ba 2+  and show that in resting channels there are two sequential binding sites and that the affinity of the deep binding site is much higher than reported for ShakerIR. Again in contrast to ShakerIR, there is only weak fast block of open channels by Ba 2+  in Kv1.5. We then assess whether Ba 2+  can either access or exit the deep binding site of resting channels at pH 5.5 and find that low external pH can prevent both the loading and unloading of the deep pore site. These results suggest that the decrease of channel availability associated with external acidification is caused by a constriction of the outer end of the selectivity filter and adds support to the hypothesis that an outer pore inactivation process occurs at rest at low pH. 40 2.2 METHODS 2.2.1 Cell preparation Wild-type hKv1.5 currents were recorded from channels stably expressed in a human embryonic kidney cell line (HEK-293; American Type Culture Collection, Rockville, MD, USA), as reported previously (Kehl et al., 2002). Cells were passaged using trypsin-EDTA and maintained in Minimum Essential Medium (MEM) supplemented with 10% foetal bovine serum, 1% (v/v) penicillin-streptomycin and 0.5 mg ml −1  geneticin at 37ºC in an atmosphere of 5% CO2 in air. All tissue culture supplies were obtained from Invitrogen (Burlington, ON, Canada). 2.2.2 Recording solutions Unless otherwise stated, the standard, nominally K + -free bath solution contained (in mM): 143.5 NaCl, 2 CaCl2, 1 MgCl2, 5 glucose, 10 HEPES and was titrated at room temperature to pH 7.4 with NaOH. In experiments where K + o was present, KCl was substituted for NaCl. Ba 2+ -containing solutions were prepared by iso-osmotic substitution of NaCl with BaCl2. The standard low pH bath solution was prepared using 10 mM MES instead of HEPES and titrated to pH 5.5 with NaOH. The patch pipette solution contained (in mM): 130 KCl, 4.75 CaCl2 (pCa 2+  = 7.3), 1.38 MgCl2, 10 EGTA, 10 HEPES and was adjusted to pH 7.4 with KOH. Chemicals were obtained from the Sigma Aldrich Chemical Co. (Mississauga, ON, Canada). In most experiments, a section of glass coverslip to which cells had attached was placed in a saline-filled recording chamber (0.5 ml volume) and perfused with control solution (2 ml min −1 ). We found no difference between experiments using continuous perfusion and those where control and test responses were recorded without perfusion (not shown). 41 2.2.3 Fast perfusion For experiments requiring rapid solution exchange, a Warner Instruments (Hamden, CT, USA) SF-77B Perfusion Fast Step system was used (see Appendix). Perfusion pipettes were made using borosilicate theta glass tubing (2 mm outside diameter, OD; Warner Instruments) that was pulled in one step to an outer diameter at the tip of approximately 300 m and then cut into two parts using a ceramic wafer (for details, see Appendix; also, Jonas, 1995). Deactivated fused silica tubing (0.18 mm inside diameter, ID; Agilent Technologies Inc., Mississauga, ON, Canada) was then inserted into the back ends of the pipette barrels, advanced as far as possible towards the tip and fixed in place by back-filling with epoxy glue. The free ends of the silica tubing were then connected to the solution reservoirs via polyethylene tubing and the manifolds provided with the Fast Step system. While the bath was constantly perfused with control solution, the assembled perfusion pipette was positioned as close as possible to the cell of choice, so that it was exposed to solution outflow from only one barrel. Rapid switching of the pipette position (200 m movement) exposed the cell to the test solution that flowed continuously from the second barrel. Using a method described by Panyi and Deutsch (2006), the time course of solution exchange, estimated from the decay in current amplitude at 0 mV when the [K + ] of the perfusion solution was switched from 0.5 to 140 mM K +  (not shown), was approximately 150 ms (see Appendix for details). 2.2.4 Electrophysiological procedures Whole-cell currents were recorded at room temperature (20 - 25ºC) using an EPC-8 patch clamp amplifier and Pulse + Pulsefit software (HEKA Electronik, Lambrecht/Pfalz, Germany), connected through an ITC-18 digital interface (Instrutech, Port Washington, NY). Patch electrodes were made from thin-walled borosilicate glass (1.2 mm OD; World Precision Instruments, Sarasota, FL) and had resistances of 1.0 – 2.5 M measured in the bath with 42 standard internal and external solutions. Capacitance and series resistance were compensated (typically 80%) using the circuitry of the amplifier. Leak subtraction was achieved using the online P/N protocol in Pulse, for which the holding potential was −100 mV and the scaling factor was −0.25. Current signals filtered at 3 kHz (−3dB, 8 pole Bessel filter) were digitised (18 bit) at a sampling interval of 100 s. Voltages were corrected for liquid junction potentials. Cells were held at −80 mV and a test potential of +50 mV was standard. Test pulses were 1.5 s long in experiments examining open channel Ba 2+  dissociation. In experiments involving repetitive stimulation at different frequencies, the pulse duration was 10 ms and the current amplitude was measured at the end of the voltage pulse. Where applicable, current amplitudes were normalized to control measurements done at the beginning of an experiment. Data from cells showing <85% recovery to control levels were discarded. 2.2.5 Data analysis Data are presented as mean  S.E.M., except for the values derived from non-linear least- squares fitting routines, which are expressed as mean  S.D. (Igor, Wavemetrics Inc., Portland, OR); n represents the number of cells. Due to the sigmoidal nature of current activation under control (0 mM Ba 2+ ) conditions, the time course of activation was estimated by fitting the portion of the trace between 50 – 100% of the peak current to a single exponential function (using a 1.5 s pulse). After Ba 2+  exposure, the entire rising phase of the first current trace obtained upon the return to control conditions could be fit to an exponential function. The time courses of currents obtained using short pulses (10 ms) were not quantified. The analysis of the relationship between the blocking rate (1/block) and [Ba 2+ ]o was based on a sequential two-site binding model:  Ba 2+  + Site1-Site2                          Ba 2+ :Site1-Site2                           Site1-Site2:Ba 2+   (SCHEME 2.1) k1 k-1 k-2 k2 43 where Site 1 and Site 2 are, respectively, the superficial and deep sites, k1 and k2 are the respective binding rates at Site 1 and Site 2, and k−1 and k−2 are the respective rates of unbinding from Site 1 and Site 2. This same model has been used previously to describe the concentration dependence of the slow block of ShakerIR channels by Ba 2+  (Hurst et al., 1995). At the superficial site the binding rate is proportional to [Ba 2+ ]. The deeper site is not directly exposed to the external bulk solution or the internal bulk solution by virtue of the closed activation gate and, as a consequence, k2 is proportional to the probability that Site 1 is occupied. We make the assumption that k−2 is independent of the occupancy of Site 1; this arose from the assumption in the original model that a coulombic interaction precludes simultaneous occupation of the two sites by Ba 2+ . Even if this constraint is removed, there is little change in the outcome of the model, because k−2, derived independently from the time dependence of the current recovery in Ba 2+ -free medium (Fig. 2.1E) is very small relative to k2 at Ba 2+  concentrations that produce a significant degree of block. Indeed, making the off-rate a function of the probability that the superficial site is not occupied (i.e., k−2 x 1/(1 + [Ba 2+ ]o/KBa,s)) has minimal effect on the fit parameters (not shown). Thus, the data in Fig. 2.1C were fit to the equation: 2 o 2 , 2block ]Ba[ 1 1    /1                  k K k sBa    (Equation 2.1) where k2 and k−2 are, respectively, the binding and unbinding rates at the deep site, KBa,s is the apparent equilibrium dissociation constant of the superficial site, and 1/(1 + KBa,s /[Ba 2+ ]o) is the probability that Ba 2+  is bound to the superficial site. Primarily to allow a comparison with the results of others, the [Ba 2+ ]o-dependence of the steady-state residual current at a given pulse frequency was also quantified by fitting the data to the Hill equation: 44 H , o 2 , ]Ba[ 1 1 1 n dBa SSB K P                  (Equation 2.2) where PB,SS is the proportion of macroscopic current blocked at the steady-state, KBa,d is the apparent equilibrium dissociation constant of the deep Ba 2+  binding site and nH is the Hill coefficient reflecting the number of Ba 2+  ions binding per channel. To model the frequency dependence of the Ba 2+  block of Kv1.5, we employed Equations 2.3-2.7, which were originally applied by Starmer et al. (1986) to describe the block of NaV channels by the quaternary lidocaine derivative QX222. Briefly, a bimolecular scheme involving unblocked channels (U), drug (D) and blocked channels (B) is:             (SCHEME 2.2) For a given channel conformation (i.e. open or closed) the steady-state level of block is kon/(kon + koff) and the time constant for the block is (kon + koff) −1 . However, if the channel conformation affects the binding equilibrium, by virtue of an effect on the accessibility of the site and/or a change of kon and/or koff, and if the lifetime of a particular conformation is short-lived with respect to the blocking time constant, then the proportion of channels blocked can be critically influenced by the frequency and duration of test pulses that cycle channels in and out of different conformations. In the case of QX222, block is enhanced by increasing the pulse frequency, which increases the proportion of time spent in one or more conformations favouring drug binding and decreases the time spent in conformations favouring drug unbinding. As will be shown, binding and unbinding of Ba 2+  exhibits the opposite relationship: channel opening during test pulses to +50 mV promotes Ba 2+  unbinding whereas the reaction is biased towards binding in a closed channel at −80 mV. Consequently, increasing the frequency of depolarizing test 45 pulses decreases the fractional block by Ba 2+ . An equation describing the blockade with repetitive stimulation (for details of the derivation see the Appendix of Starmer et al. 1986) is: )exp(1 exp1expexp1 ,                                         r r d d r d d d SSB tt P t P P             (Equation 2.3) where PB,SS is the proportion of channels blocked at the steady-state, Pd is the proportion of channels blocked at the deep site after a prolonged depolarization, Pr is the proportion of channels blocked at the deep site after a prolonged rest at the holding potential, td is the average amount of time that a channel is open during a depolarizing pulse, d is the time constant for unblock during a depolarization to the test voltage, tr is the time at the holding potential between test pulses, r is the time constant for the block at the holding potential, and λ is equal to the unitless quantity (td/d + tr/r). With the assumption that Pd is zero (Hurst et al., 1995; Harris et al., 1998), Equation 2.3 simplifies to: )exp(1 exp1exp ,                          r r d d r SSB tt P P                      (Equation 2.4) Since Ba 2+  can be trapped in open-inactivated channels (Basso et al., 1998; Harris et al., 1998) this equation is valid for prolonged depolarizations only if the value for td is appropriately adjusted, otherwise the degree of unblock would be overestimated. With the 10 ms test depolarizations used in the quantification of the Ba 2+  block, inactivation is not a significant mitigating factor and the estimate of td is calculated from the open probability estimated from prior single channel recordings (see Results). The fractional block at the end of the x th  pulse of a train (PB,x) is: 46     ...2,1,exp  ,,,  xxPPPP SSBinitialSSBxB           (Equation 2.5) where Pinitial is the proportion of the current blocked at the point in time when the pulse frequency is changed,  is as defined for Equation 2.3, and x is the pulse number in the stimulus train. In words, Equation 2.5 indicates that changing the stimulus frequency causes the proportion of current blocked during a pulse train to relax mono-exponentially to a new steady- state level.  The relationship between the time constant () for that relaxation and  is:  t x ee             (Equation 2.6) Given that t, the time elapsed since the change of the pulse frequency, is equal to the product of the pulse number (x) and the cycle length (CL), then the time constant can be calculated by:     CLxCL x yields                                      (Equation 2.7) To describe the [K + ]o-dependence of the open channel Ba 2+  unbinding rate (see Fig. 2.6F), the Ba 2+  dissociation rates measured at +50 mV with different [K + ]o were fit to the Hill equation, H d o max, ]K[ 1  n off off K k k               (Equation 2.8) where koff is the apparent rate of Ba 2+  unbinding from open channels at +50 mV (taken as 1/act,app, where act,app is the apparent activation time constant; see Fig. 2.6), koff,max is the unbinding rate with 0 mM K + o, Kd is the apparent equilibrium dissociation constant for the K + o inhibition of Ba 2+  unbinding and nH is the Hill coefficient reflecting the number of K +  ions binding per channel to slow Ba 2+  unbinding. 47 To quantify the voltage-dependence of the open channel Ba 2+  dissociation rate (see Fig. 2.6D), koff  was measured over a range of test potentials (with a fixed [K + ]o and [K + ]i), plotted against the test potential and fit to the Woodhull model: koff (V) = koff (0mV) x exp (2δBa,offFV/RT)                   (Equation 2.9) where koff(V) is the Ba 2+  unbinding rate at the test voltage, koff (0 mV) is the unbinding rate at 0 mV, V is the test potential, δBa,off is the electrical distance between the deep Ba 2+  binding site and the rate-limiting barrier for exit, and F, R, and T have their usual meaning (Woodhull, 1973).  48 2.3 RESULTS 2.3.1 Ba2+ blocks closed Kv1.5 channels at pH 7.4 Our first objective was to test the hypothesis that Ba 2+  is able to access and bind within the pore of Kv1.5 channels and block current. We began by characterizing the effects of external Ba 2+  on Kv1.5 currents at pH 7.4 using a stimulus protocol comparable to that used to investigate Ba 2+  block of ShakerIR channels (Hurst et al., 1995). Figure 2.1A illustrates a representative experiment examining the effect of 5 mM Ba 2+  on currents evoked by 10 ms pulses to +50 mV applied at a frequency of 1 Hz from a holding potential of –80 mV. Tail current was recorded at −80 mV. Scaling of the steady-state current evoked in 5 mM Ba2+ and superimposition on the control current (Fig. 2.1A, inset) revealed that the activation and deactivation kinetics of the steady-state current observed with 1 Hz stimulation were slightly affected by the 5 mM Ba 2+  treatment.  49  Figure 2.1. Ba 2+  blocks Kv1.5 in a concentration- and frequency-dependent manner.  50 Figure 2.1. Ba 2+  blocks Kv1.5 in a concentration- and frequency-dependent manner. A, With 0 mM K + o, pH 7.4 solution, 10 ms test pulses to +50 mV were applied at 1 Hz without (uppermost trace) and with 5 mM Ba 2+ . Shown are 65 consecutive traces from the time of Ba 2+  application, which began immediately after the control trace. Inset, The trace at the steady-state level of block was scaled 1.87x to match the peak amplitude of the control current. There was a small effect of Ba 2+  on the onset of the K +  current. B(i), Trace 1 was recorded using a 10 ms pulse to +50 mV after a 1 min exposure to 1 mM Ba 2+  in 0 mM K + o at −80 mV. Traces 2-5 show the current from subsequent pulses applied at 1 Hz in the continued presence of Ba 2+ . Trace 25 shows the steady-state current in1 mM Ba 2+  with 1 Hz stimulation and Trace 100 is the steady- state current following recovery in 0 mM Ba 2+ . B(ii), Traces 1-5 and 25 are normalized with respect to Trace 100. There was a marked slowing of the rising phase of the current elicited by the first pulse after Ba 2+  exposure but this effect is diminished with subsequent pulses in the continued presence of Ba 2+ . C, The time course of the onset and offset of the Ba 2+  block. The average current amplitude in the last 1 ms of each pulse was normalized to the Ba 2+ -free control current. Down- and up-arrows indicate the time of fast Ba 2+  application and withdrawal, respectively. Black circles () represent the normalized current amplitudes for the experiment shown in A. Single exponential fits to the onset of and recovery from Ba 2+  block give block = 6.84 s and unblock = 19.21 s, respectively. When the experiment was performed with the same stimulus protocol but with 20 mM Ba 2+  (▲), block onset was slightly faster (block = 4.97 s) and the normalized steady-state current was smaller (0.37 vs. 0.50 for 5 mM Ba 2+ ), but the time course of current recovery was the same. If the [Ba 2+ ] was kept constant at 5 mM, but the pulse frequency decreased to 0.2 Hz (open diamonds; ), the normalized steady-state level was also reduced (steady-state normalized current = 0.27). Although the onset of the block was not affected by the decrease in stimulation frequency, recovery with a 0.2 Hz pulse frequency was much slower (unblock = 63.20 s) than that seen at 1 Hz. D, The blocking rate (1/block) is a non- linear function of the [Ba 2+ ]. block values were assessed in 0 mM K + o, with 10 ms pulses to +50 mV applied at 1 Hz. The solid line represents the best fit of the data to a sequential 2-site binding model (Equation 2.1; see text for fit parameters). Data points are from a total of 46 cells, 3-8 cells per point. E, The unblocking rate is linearly related to the pulse frequency. Values for unblock were assessed with 0 mM K + o, 20 mM Ba 2+ o, using 10 ms pulses to +50 mV applied at 1, 0.7, 0.2, 0.066, or 0.033 Hz. The extrapolated off-rate without pulsing (0 Hz) is 0.001  0.002 s −1 . Data points are from a total of 53 cells, 5-24 cells per point. 51 For the currents shown in Fig. 2.1B(i), 1 mM Ba 2+  was applied for 1 min in the absence of pulses before initiating a train of 1 Hz pulses to +50 mV. The current in the first pulse had a small initial amplitude and slowly activating time course, which is consistent with the block of virtually all of the channels at the start of the pulse, followed by a slow unblock during the pulse (Harris et al., 1998). With continued pulsing in the presence of Ba 2+  the degree of block at equilibrium (Trace 25) was nearly identical to that in Fig. 2.1A. Trace 100 is the steady-state recovery response obtained in Ba 2+ -free medium. For Fig. 2.1B(ii) select traces were normalized with respect to Trace 100 to better illustrate their relative activation time courses. These data suggest that repetitive pulsing has a substantial effect on the steady-state level of block of Kv1.5 by Ba 2+ , consistent with the observation made by Armstrong and colleagues on the Ba 2+  block of the squid giant axon delayed rectifier channels (Armstrong et al., 1982). A detailed analysis of the frequency-dependence of Ba 2+  block of Kv1.5 is presented below. To illustrate the time dependence of the onset and offset of Ba 2+  block, the normalized mean current in the last millisecond of each test pulse from the experiment described by Fig. 2.1A has been plotted against time in Fig. 2.1C (); the downward arrow indicates the time at which the 5 mM Ba 2+  fast perfusion began. In five experiments of this type the relaxation of the normalized test current (Inorm) to a steady-state level of 0.52  0.01 was well fitted by a single exponential with a time constant (block) of 7.2  0.6 s. After returning to Ba 2+ -free medium, marked by the upward black arrow, the time course for complete reversal of the Ba 2+  block was also mono-exponential (unblock = 18.5  2.6 s). In an experiment with a four-fold higher concentration of Ba 2+  (20 mM, , Fig. 2.1C) but with the same pulse frequency of 1 Hz, the onset of the block was only slightly faster (block = 4.8  0.5 s; n = 9), and the steady-state block (1 − Inorm = 0.61  0.01) was ≈20% greater. Note that after switching from 20 mM Ba 2+  back to 52 control solution unblock was 12.8  2.1 s, which was not significantly different (P > 0.05) from unblock measured following the 5 mM Ba 2+  treatment. For the third set of responses illustrated in Figure 2.1C (), a 5 mM concentration of Ba 2+  was used but the stimulus frequency was 0.2 Hz. The block (9.2  0.3 s; n = 5) was similar to that observed with a 1 Hz pulse frequency using either 5 or 20 mM Ba 2+ . However, the degree of block (0.74  0.01) was greater and unblock was slower (unblock = 60.5  2.9 s) than with the 1 Hz test pulse frequency following channel loading using either 5 or 20 mM Ba 2+ . Thus, the Ba 2+  block of Kv1.5 is similar to that reported for ShakerIR (Hurst et al., 1995; Harris et al., 1998) in that the steady-state level of block and the kinetics of block onset are dependent on both the Ba 2+  concentration and frequency of test pulses applied, while the time course of current recovery from Ba 2+  block is sensitive only to the pulse frequency. Despite these qualitative similarities in the features of the Ba 2+  block of Kv1.5 (Fig. 2.1A,C) and ShakerIR currents, the results of Fig. 2.1 also reveal substantial qualitative and quantitative differences, which is perhaps unexpected given the similarity of their pore structures (see Fig. 2.2). For example, in ShakerIR with 20 mM Ba 2+ , the inhibition of current consists of a very rapid phase, attributable to fast block of open channels during the test pulse, as well as a slow phase that relaxes to the steady state with a time constant of approximately 100 s (see Fig. 2 of Hurst et al., 1995). In contrast, but similar to findings in Kv1.3 (Somodi et al., 2004; Somodi et al., 2008), there is no rapid component of block of Kv1.5 by 20 mM Ba 2+  (Fig. 2.1C) and the slow block occurs roughly 10 times faster than in ShakerIR. In the light of these differences, we undertook a more detailed analysis of the Ba 2+  block of Kv1.5 in the interest of highlighting mechanistic differences in the Ba 2+  block of these closely-related channels and to validate the use of Ba 2+  as a tool to probe the Kv1.5 pore at low external pH in the second part of this chapter. 53  Figure 2.2. Sequence alignment of hKv1.5 and Shaker from the C-terminal end of S5 to the end of the selectivity filter. The putative regions of Kv1.5 and Shaker forming the turret, pore helix and selectivity filter are shown.  2.3.2 block is not a linear function of [Ba 2+ ] In order to develop a kinetic model for the Ba 2+  block, we examined the relationship between the blocking rate, which was taken as the inverse of τblock following Ba 2+  application in K + -free bath solution, and the concentration of Ba 2+  applied (Fig. 2.1D). As reported for ShakerIR, the blocking rate of Kv1.5 current saturated at a high [Ba 2+ ]o, indicating that a bimolecular blocking reaction was not involved. The data were fit very well to a sequential two- site binding model used to describe the Ba 2+  block of closed ShakerIR channels (see Section 2.2.5 and Hurst et al., 1995). The best fit of the experimental data to Equation 2.1, which is based on Scheme 2.1, is shown as a solid line in Fig. 2.1D and gave an estimate of 0.93 ± 0.15 mM for the apparent equilibrium dissociation constant (KBa,s = k−1/k1) of the superficial Ba 2+  binding site, 0.16 ± 0.01 s −1  for k2 and 0.0025 ± 0.0019 s −1  for k−2. An independent estimate of k−2 was obtained from experiments where Kv1.5 channels were first loaded with Ba 2+  and, after switching back to Ba 2+ -free medium, unblock was measured with test pulses to +50 mV applied at varying frequencies. The underpinning assumption of the experimental approach, which is validated below in Kv1.5 and supported by results in ShakerIR (Harris et al., 1998), is that because depolarization facilitates the unblocking reaction and, since recovery occurs in Ba 2+ -free medium, rebinding of Ba 2+  is therefore unlikely to occur and the relationship between unblock and the pulse frequency allows k−2 to be estimated. Recovery time 54 courses were always well fitted by a single exponential, regardless of the stimulus frequency (e.g., Fig. 2.1C), and unblock was independent of either the extent of the block or the Ba 2+  concentration used in the loading phase of the experiment (not shown). As expected for an unblocking reaction that is facilitated by depolarization, the apparent unblocking rate (1/unblock) is proportional to the stimulus frequency (Fig. 2.1E) when the test pulse duration is fixed and brief. Extrapolation of the fitted line to the y-intercept provides an estimate of k−2 in the absence of pulsing, or, in other words, the value for the Ba 2+  off-rate from the deep binding site at the holding potential of −80 mV and with 0 mM external K+. The best estimate of k−2 derived from Fig. 2.1E is 0.001 ± 0.002 s −1  and buttresses the estimate of 0.0025 ± 0.0019 s −1  from Fig. 2.1D. However, both estimates have a large standard deviation and their accuracy is therefore uncertain. In comparison, the upper limit for the resting state off-rate in ShakerIR channels is 0.0132 s −1  (Harris et al., 1998). 2.3.3 Ba2+ block of Kv1.5 is dependent on stimulation frequency Although an effect of the test pulse frequency on the steady-state level of Ba 2+  block has been noted with ShakerIR and other K +  channels (Armstrong et al., 1982; Harris et al., 1998), to our knowledge this relationship has not been systematically studied. Indeed, an explicit assumption has been that a brief test pulse would have little effect on the slow component of block (Hurst et al., 1995). Especially in view of the greatly enhanced block observed following a 1 min interpulse interval (Fig. 2.1B), a more detailed examination of the frequency-dependence of the Ba 2+  block was warranted. To quantify the relationship between the Ba 2+  block and the test pulse frequency, the steady-state level of block was assessed at two different stimulation frequencies over a range of [Ba 2+ ]. Current amplitude was measured at the end of 10 ms test pulses to +50 mV applied once 55 every 30 s (0.033 Hz) or once every 1.43 s (0.7 Hz). Ba 2+  was applied by a computer-controlled fast perfusion system and the blocking reaction allowed to proceed to the steady-state. Steady- state residual current was calculated by normalizing the peak test current at the steady-state to the control test currents obtained prior to Ba 2+  application. Figure 2.3 shows the concentration- response relationship at 0.033 Hz () and at 0.7 Hz (). Separate fits of the two data sets to the Hill equation (black lines of Fig. 2.3; Equation 2.2) gave estimates for the apparent affinity of the deep Ba 2+  binding site, KBa,d, and the Hill coefficient of 20.3 ± 2.6 M and 1.12, respectively, for the 0.033 Hz pulse frequency, as compared to 361 ± 151 M and 1.16 for responses evoked at 0.7 Hz. This demonstrates that increasing the pulse frequency from 0.033 Hz to 0.7 Hz causes an 18-fold increase of the mid-point of the concentration-response relationship while having no effect on the Hill coefficient, which had a value consistent with deep block arising from the binding of one Ba 2+  ion. Increasing the pulse frequency to 0.7 Hz also caused an upward shift of the foot of the concentration-response relationship. This upward shift reflects the fact that with a saturating concentration of Ba 2+  the degree of block with test pulses of a fixed duration (and a fixed td) is proportional to the time spent at rest (tr). In other words, the decrease of tr as the stimulus frequency increases means there is less time for the re-establishment of block at rest and, as a consequence, the proportion of unblocked channels increases. 56   Figure 2.3. The steady-state level of Ba 2+  block is dependent on the stimulation frequency and the [Ba 2+ ]. Test pulses to +50 mV for 10 ms were applied at either 30 s intervals (0.033 Hz ; ) or 1.43 s intervals (0.7 Hz; ) with 0 K+o and a pH of 7.4. Ba 2+  was applied at various concentrations using a computer-controlled fast perfusion system. The steady-state residual current (1 − proportion blocked (PB,SS)) at the end of the test pulse was normalized to the control test current before Ba 2+  application and plotted against the concentration of Ba 2+  applied (2 M – 20 mM). The black solid lines represent the best fits of the two data sets to the Hill equation. Estimates of the apparent KBa,d and the Hill coefficient are 20.3 ± 2.6 M and 1.12, respectively, for the data collected at 0.033 Hz, while the estimates from the 0.7 Hz data are 361 ± 151 M and 1.16, respectively. Grey dashed lines represent the outcome of a simultaneous fit of the two data sets to Equation 4. From the fit, k2, KBa,s, k−2 , and τd were estimated to be 0.15 s −1 , 0.8 mM, 0.0006 s −1 , and 59 ms, respectively (see text). The dotted grey line represents the solution for the residual normalized current (1 − Pr), where Pr is the proportion of channels blocked after a prolonged rest at the holding potential, i.e. at 0 Hz. A fit of 1 – Pr  to the Hill equation gave a Hill coefficient of 1 and a KBa,d of 4 M, indicating that the apparent KBa,d is profoundly affected by the pulse frequency. Data points are from a total of 29 cells (2 – 8 cells per point) for the 0.7 Hz data; and 45 cells (3 – 9 cells per point) for the 0.033 Hz data. 57 To model the frequency-dependence of the concentration-response relationship, the data obtained at 0.033 Hz and 0.7 Hz were simultaneously fit to Equation 2.4; the outcome of the fit is represented by the grey dashed lines of Fig. 2.3. There are 6 parameters in the fitting equation: td, tr, k2, k−2, KBa,s and d. The value for r in Equation 2.4 arises from, r = (k2 x (1+ KBa,s /[Ba 2+ ]) −1  + k−2)  −1 = (k2 + k−2)  −1 and the value for Pr from, Pr = k2 / (k2 + k−2) where r is the time constant for the Ba 2+  block of the deep pore site at the holding potential, and Pr is the proportion of channels with Ba 2+  occupying the deep site at the steady-state. To estimate td, defined as the average open time during a 10 ms test pulse to +50 mV, the proportion of maximal charge conducted during a pulse was first calculated by dividing the integral of a control pulse current by the integral for the current if activation had been instantaneous (i.e., Imax x 10 ms). The td was then obtained by multiplying the proportion of maximal charge by the maximal open probability (Po) of 0.8 measured from recordings of unitary Kv1.5 currents (Kwan et al., 2006). The mean td calculated with this approach was 6.43 ± 0.25 ms (n = 7 cells). Ba 2+  binding at the deep pore site is assumed to have no effect on td (but see Armstrong et al., 1982). For tr, the time spent at the holding potential during one cycle length, a fixed value of (1/stimulus frequency in Hz − 0.01 s) was used. Consequently, there were four free parameters in the global fitting routine and, of these, the values for k2, k−2 and KBa,s were constrained to be in the range of the mean values ± 1 SD obtained from the fit of the data in Fig. 2.1D. With these conditions, the best fit values for the constrained parameters, k2, KBa,s, k−2 were 0.15  0.08 s −1 , 0.8  0.3 mM and 0.0006  0.0009 s−1, respectively. For d, an unconstrained free parameter, the best estimate obtained from the simultaneous fit to the 0.033 and 0.7 Hz data was 59  19 ms. 58 Apart from providing a good estimate of the shift of the concentration-response relationship at different frequencies, the best fit values derived in Fig. 2.3 can be used to approximate the proportion of unblocked channels at the deep site in the absence of test pulses. This is shown in Fig. 2.3 as the dotted grey line and represents the solution for 1−Pr, where Pr is as defined above. A fit of 1−Pr to the Hill equation gave a Hill coefficient of 1 and a KBa,d of 4 M. In other words, with 4 M Ba2+ the on-rate constant (k2) is equal to the off-rate constant (k−2) and half of the channels have Ba 2+  bound at the deep site. This suggests that even with 10 ms test pulses applied only once every 30 s the KBa,d is overestimated roughly 5-fold, while at 0.7 Hz the KBa,d is overestimated roughly 90-fold, and emphasizes the potential importance of taking the pulse frequency and duration into account when comparing the relative sensitivities of channels to block by Ba 2+ . Thus, reported millimolar values of KBa,d for ShakerIR (Hurst et al., 1995; Harris et al., 1998) describe the apparent affinity of the deep site at a specific voltage and pulse frequency. Equation 2.4 has been shown to fairly faithfully describe the frequency dependence of the concentration-steady state block relationship; however, the analysis of Fig. 2.3 provides no indication of how well the model replicates the time-dependence of either the onset of, or the recovery from block. Experimental data addressing that issue are shown in Fig. 2.4A. For this representative experiment, at the end of several 10 ms control test pulses to +50 mV, a computer- controlled, rapid changeover from zero to 0.2 mM Ba 2+ was made and that concentration was then maintained while test pulses were applied at 0.1 Hz (). Once a steady level of block was attained, the stimulus frequency was decreased to 0.033 Hz () to monitor the progression of the block to a new steady-state. Complete reversal of the enhanced steady-state block observed with 0.033 Hz pulsing was then obtained after reverting to the 0.1 Hz stimulus frequency. The same approach was subsequently repeated with trains of 0.2 (), 0.5 () and 0.7 () Hz pulses. 59 Figure 2.4B shows the output of a simulation of the proportion of unblocked current (1−PB,x; see Equation 2.5) at the end of the x th  test pulse and was derived using the values for k2, KBa,s, k−2, and d obtained from the fit to the data of Fig. 2.3. The values for td, tr and r were calculated as described in connection with Fig. 2.3; Pinitial of Equation 2.5 is 1 minus the proportion of current at the end of the preceding epoch. Figure 2.4B illustrates that the analytical solution reproduces the experimental data quite well not only in terms of the steady-state level of residual normalized current, as expected, but also from the standpoint of the time dependence of the relaxations to a new level of residual current following a change of the test pulse frequency. These results also suggest that the values of k2, KBa,s and k−2 are good estimates of the rate and equilibrium constants for Ba 2+  binding to Kv1.5 channels and provide further support for the two-site binding model. 60   Figure 2.4. A model of the frequency-dependent Ba 2+  binding replicates the time- dependence of block onset and reversal. A, A representative experiment in which a cell was given four 10 ms test pulses to +50 mV in 0 mM Ba 2+  prior to a rapid changeover to 0.2 mM Ba 2+  while test pulses continued to be applied at 0.1 Hz (). After steady-state block was achieved, the pulse frequency was decreased to 0.033 Hz () and the block was allowed to reach a new steady-state. The enhanced steady-state block observed with 0.033 Hz stimulus frequency was reversed upon reverting to 0.1 Hz pulses. The effects of changing the pulse frequency to 0.2 (), 0.5 () and 0.7 () Hz were subsequently examined using the same approach. Data points show current amplitudes normalized to the peak current at the end of the initial control test pulses. B, Plot of simulated proportion of residual normalized current (1 – PB,x) at the end of each test pulse shown in A, calculated using Equation 2.5 and the values for k2, KBa,s,, k−2, and d derived from fits to the data of Figure 2.3. The values for td, tr, and r were those derived from Figure 2.3 (see text), while Pinitial of Equation 2.5 is 1 minus the proportion of current at the end of the preceding pulse train. At each stimulus frequency the numerical simulation provides a good estimate both of the steady-state level of block and the time course of the relaxation to that steady state. 61 2.3.4 Ba2+ causes a weak fast block of open Kv1.5 channels As noted above, the time course of Ba 2+  block onset in Kv1.5 lacks the fast phase seen in ShakerIR channels (Hurst et al., 1995; Harris et al., 1998) and instead exhibits a slow, mono- exponential component (see Fig. 2.1C). However, our analysis of the [Ba 2+ ]-dependence of the blocking rate (1/block; Fig. 2.1D) suggested that, as with ShakerIR, Ba 2+  binds first to a low affinity (KBa,s  1 mM) superficial site and then to a high affinity (KBa,d = 4 M) deep site. Given the low value of KBa,s compared to the concentration of Ba 2+  used in experiments such as those of Fig. 2.1C, it was somewhat surprising that a fast phase of block onset related to Ba 2+  binding to a superficial site was not seen. This motivated experiments where a brief pulse of 20 mM Ba 2+ was applied during a 10 s pulse to +50 mV (Fig. 2.5). At the onset of the Ba 2+  application, the current amplitude rapidly decreased by ~10%, with a time constant that we assume was governed primarily by the kinetics of the solution change. Upon the switch back to Ba 2+ -free solution the current recovered to the same amplitude as that of the control current. These results indicate a fast but comparatively weak block of Kv1.5 by 20 mM Ba 2+ , likely by binding to the superficial pore binding site. If so, the affinity of this site for Ba 2+  in the open state is apparently much lower than that estimated for the superficial site in the closed state (Fig. 2.1D) and appears to account for the lack of a fast phase of block in experiments such as those shown in Fig. 2.1C.  62    Figure 2.5. Fast application of Ba 2+  during a long depolarising pulse reveals very weak open channel block by 20 mM Ba 2+ . Current traces in response to depolarising pulses to +50 mV for 10 s in 0 mM K + , Ba 2+ -free solution (black trace) and with a rapid application and withdrawal of a 0 mM K + , 20 mM Ba 2+  solution during a subsequent pulse (grey trace). Down and up arrows indicate the start and end of the Ba 2+  application, respectively. Exposure to Ba 2+  caused a rapid fall in current amplitude of ~10%. A similar effect was observed in 6 other cells.  63 2.3.5 Ba2+ block of Kv1.5 at rest and open channel dissociation As mentioned in connection to Fig. 2.1B, when Ba 2+  is applied for a prolonged period to resting channels, the first 10 ms pulse shows a slow activation time course mainly reflecting a slow unblock of channels. To characterize this unbinding of Ba 2+  ions from open channels and determine whether it was dependent on factors such as the test voltage or [K + ]o, experiments were performed where Ba 2+  was applied for 2 min to closed Kv1.5 channels at −80 mV in 0.5 mM K + o (see experimental protocol in Fig. 2.6A). Following a 4 min wash to remove all Ba 2+ , a 1.5 s step to +50 mV was used to assess the time course of Ba 2+  unbinding from the open channel. In agreement with findings in other K +  channels (Armstrong et al., 1982; Neyton & Miller, 1988b; Harris et al., 1998) and our estimate of k−2, Ba 2+  remained bound to the channels even after the prolonged wash-out period, such that the first pulse elicited current with a much slower rise time than that seen prior to Ba 2+  application (Fig. 2.6B). The lack of either an initial delay or fast phase, which would be indicative of current through channels that were not blocked at the start of the pulse, together with the single exponential time course of the “post-Ba2+” current rising phase (act,app = 22.56  1.25 ms; n = 11) suggest that with the loading protocol used, all of the channels were Ba 2+  bound prior to the pulse. (With 0 K + o there was, in addition to a slow component with act,app = 15.30  0.99 ms, which we take to reflect Ba 2+  unbinding, a very slow component with  = 1.08  0.14 s (n = 10; data not shown). Investigation of the slower component of current was beyond the scope of this study but we do note that confounding results with 0 K + o solutions and Ba 2+  have also been reported by others (Harris et al., 1998)). The first sweep after Ba 2+  loading showed little or no decay over the course of the pulse, perhaps because of the overlap of inactivation and Ba 2+  unblocking. From the rising phase of the current, the Ba 2+  unbinding rate (1/act,app) was calculated to be 45.79  3.08 s −1 , although this may be an underestimate if substantial inactivation, and hence Ba 2+  trapping, occurs during the rising phase. 64 Additional depolarisations (not shown) evoked currents with activation kinetics similar to that of control, with a small (< 10%) slow component, perhaps associated with residual Ba 2+  that did not unbind during the initial 1.5 s pulse. With subsequent pulses, the contribution of the slow component diminished and current amplitude recovered towards control values. 65  Figure 2.6. Ba 2+  unbinding from the open state is voltage-dependent and inhibited by increasing the [K + ]o. 66 Figure 2.6. Ba 2+  unbinding from the open state is voltage-dependent and inhibited by increasing the [K + ]o. A, Schematic of voltage-pulse and Ba 2+  application protocol. A 1.5 s pulse to +50 mV was applied under 0.5 mM [K + ]o, pH 7.4 conditions. The cell was then exposed to 5 mM Ba 2+  for 2 min, followed by a 4 min wash in control solution, after which the test pulse was repeated. B, Following Ba 2+  exposure (grey trace) the current activation was much slower and well fitted by a single exponential (act, app = 28.14 ms; compare to act = 1.25 ms for the control response (black trace). Inset shows current activation on an expanded time scale. These results are representative of those obtained from 11 cells. C, normalized currents after a 2 min exposure to 5 mM Ba 2+  (with 0.5 mM K + o) recorded at two different test pulse potentials. The activation time course at both voltages was well fitted by a single exponential. At +50 mV, act, app was 22.56  1.25 ms (n = 11), which was significantly faster (P< 0.001) than at +25 mV (act, app = 65.59  7.33 ms; n = 3). Each trace was normalized with respect to its own peak current. D, plot of the Ba 2+  off-rate (1/act, app) derived from experiments such as those described in C, obtained between 0 and +100 mV in 25 mV increments. A fit of the data to the Woodhull model (see Methods) gave an estimate for Ba,off of 0.51  0.04. Data points are from a total of 21 cells; 3-11 cells per point. E, Following Ba 2+  loading (in 0.5 mM K + o) the time course of the current onset in 0.5 mM or 10 mM K + o was examined. The current activated significantly faster with 0.5 mM K + o (act,app = 22.56  1.25 ms; n = 11) than with 10 mM K + o (act, app = 145.40  6.15 ms; n = 5; P < 0.001). Traces were normalized as in C. F, plot of the Ba 2+  off-rate under varying [K + ]o at a test pulse potential of +50 mV. The best fit of the data to the Hill equation (see Methods) gave a Kd for the K + o-inhibition of Ba 2+  unbinding of 1.00  0.08 mM. Data points are from a total of 31 cells, 5-11 cells per point.  67 As reported by others (Neyton & Miller, 1988b; Harris et al., 1998), we found that in Ba 2+ -free medium the dissociation rate of Ba 2+  depended on both the step potential and [K + ]o. Ba 2+  unbinding was faster at more positive potentials (Fig. 2.6C, D), suggestive of outward, voltage-dependent dissociation. Using the Woodhull model (1973), the electrical distance (δBa,off) between the deep-Ba 2+  binding site and the rate-limiting barrier for exit was estimated to be 0.51  0.04. This value is similar to those calculated for ShakerIR (δBa,off = 0.37) and BKCa channels (δBa,off = 0.45) (Neyton & Miller, 1988a; Neyton & Miller, 1988b; Harris et al., 1998). When [K + ]o was changed the properties of control currents were unaffected (not shown), which is consistent with previous reports that, at pH 7.4, Kv1.5 open probability is insensitive to changes in [K + ]o (Jäger & Grissmer, 2001). After a 2 min loading period with 5 mM Ba 2+  (in 0.5 mM K + o) to block all channels (see Fig. 2.6B), the cells were washed with solutions of varying [K + ]o and test pulses to +50 mV were repeated. As shown in Figs. 2.6E and F, Ba 2+  unbinding was slowed by increasing [K + ]o. A fit of the unbinding rates and [K + ]o to the Hill equation (see Methods) gave an apparent Kd of 1.00  0.08 mM [K + ]o for this effect, with a Hill coefficient of 1.02  0.06. We take this to be the dissociation constant at +50 mV for a K+ binding site external to the deep Ba 2+  ion binding site that needs to be vacant for Ba 2+  unbinding, and that would be analogous, if not homologous, to the “lock-in” site described in BK and ShakerIR channels (Neyton & Miller, 1988b; Hurst et al., 1995; Harris et al., 1998). 2.3.6 Ba2+ does not block Kv1.5 at low pH The results presented thus far are consistent with Ba 2+  binding sequentially to two sites in Kv1.5, with the stable block of the channel resulting from Ba 2+  occupancy of the deeper, higher affinity site. Having established that Ba 2+  can access and bind within the pore of Kv1.5 at rest, we proceeded to examine the hypothesis that extracellular acidification results in outer pore constriction and an altered ability of Ba 2+  to bind to the pore. Figure 2.7A illustrates the results 68 of experiments monitoring the current amplitude with 10 ms pulses to +50 mV applied at 1 Hz during the fast application of Ba 2+  at low pH. When the pH was decreased from pH 7.4 to pH 5.5, current amplitude fell quickly to ~6% of the control level. The small residual current is consistent with our previous findings that at low pH an equilibrium exists between the available and unavailable states of the channel (Kwan et al., 2006); it also likely includes a small contribution from endogenous K +  currents in the HEK-293 cell line (Kehl et al., 2002). Subsequent application of 20 mM Ba 2+  at pH 5.5 caused a small additional decrease of the current amplitude. Upon the rapid washout of Ba 2+  and return to pH 7.4, the recovery time course was fast (recov = 4.01  0.85 s; n = 4) and was not significantly different (P > 0.05) from that following pH 5.5 treatment alone ( = 2.60  0.18 s; n = 5; not shown). If lowering pH had no effect on Ba 2+  accessibility, we would have expected to see ~70% block of current under these conditions and, thus, a significant slowing of the recovery time course upon the return to pH 7.4. These results suggest that lowering pH prior to and during Ba 2+  application largely prevents Ba 2+  from accessing its blocking site in most of the channels, allowing them to quickly return to a conducting state after changing back to pH 7.4.  69  Figure 2.7. Low pH inhibits both Ba 2+  binding and unbinding from Kv1.5. A, Diary plot of normalized current amplitudes at the end of 10 ms pulses to +50 mV applied at 1 Hz. Experiments were started in 0 mM K + , pH 7.4 bath solution. Rapid switching to pH 5.5 solution caused a rapid fall of the current amplitude. Subsequent application of 20 mM Ba 2+  (downward arrow), still at pH 5.5, caused a further, small decline. The rapid return (upward arrow) to Ba 2+ - free, pH 7.4 solution resulted in a quick recovery of current amplitude (recov = 3.1 s), which would not be expected had Ba 2+  accessed the deep pore site. B, In the converse of the experiment shown in A, 20 mM Ba 2+  was applied (downward arrow) at pH 7.4, resulting in 71% block of current, and was followed by a rapid wash-off of Ba 2+  (upward arrow) with pH 5.5 solution that caused an almost complete loss of current. Fast change of the bath solution back to pH 7.4 caused a rapid component of current recovery, followed by a large slow component (64% of total recovery) with a time constant of 23.6 s. These results are consistent with the interpretation that the proportion of Ba 2+ -loaded channels is not affected by high frequency pulsing when the pH is low. 70 2.3.7 Ba2+ unbinding is inhibited at low pH To test whether the lack of Ba 2+  block at pH 5.5 is due to H + -induced unbinding of Ba 2+ , which would be indistinguishable from an inability of Ba 2+  to bind in the experiments shown in Fig. 2.7A, experiments were also performed where the pH of the bath solution was decreased following Ba 2+  loading. Fig. 2.7B shows the results from a representative experiment where the current level was allowed to reach a steady-state (72  2% block; n = 6) in 20 mM Ba2+ with pulses applied at 1 Hz, after which the cell was returned to Ba 2+ -free medium first at pH 5.5 and then at pH 7.4. Changing to the pH 5.5 solution caused a further, and almost complete, reduction in current magnitude. In contrast to the situation where Ba 2+  was applied after switching to pH 5.5, the return to pH 7.4 saw current recovery proceed with a fast phase and a slow phase. The amplitude of the fast phase was similar to that of the residual current recorded at the end of the Ba 2+  loading protocol at pH 7.4 (39  4% vs. 28  2% of control amplitude, respectively) and, thus, reflects the relatively rapid recovery of Ba 2+ -free channels from the action of pH 5.5 solution. Had significant Ba 2+  unbinding occurred at the same rate as at pH 7.4 (Fig. 1C) then 63% recovery from the Ba 2+  block would have occurred. Additionally, the time constant of the slow recovery phase (slow = 25.59  5.32 s) was not significantly different (P > 0.05) from that observed for current recovery at pH 7.4 following Ba 2+  loading (Fig. 2.1C). These results are consistent with an inability of Ba 2+  to unbind from Kv1.5 at low pH. Together with the findings from the previous section, this suggests that the conduction pathway between the deep pore binding site and the external solution is changed sufficiently under acidic conditions to restrict free ingress and egress of Ba 2+  to and from its binding sites and is supportive of a low pH- mediated constriction of the outer pore of resting channels. 71 2.4 DISCUSSION 2.4.1 Ba2+ block at pH 7.4 The first series of experiments presented in this chapter show that in Kv1.5, at pH 7.4, external Ba 2+  ions cause a high affinity, slow block of closed channels (Figs. 2.1, 2.3, 2.4) and a low affinity, fast block of open channels (Fig. 2.5). While the time course of the onset of closed channel block (block) followed a single exponential, rather than showing distinct fast and slow phases as in ShakerIR, the dependence of block on [Ba 2+ ]o with 0 mM K + o was still well described by a sequential two-site binding model (Fig. 2.1D) in which the loading of a deep site with an apparent equilibrium dissociation constant (KBa,d) in the low micromolar range depends on the occupancy of a superficial site with an apparent dissociation constant (KBa,s) of approximately 1 mM. Near-saturation of the superficial site with 5 to 20 mM Ba 2+  explains why block was not linearly related to [Ba 2+ ]o (Fig. 2.1D), as would occur for a simple bimolecular reaction. It is unlikely that the slow Ba 2+  block and/or lack of an apparent fast block are due to non-specific effects of charge screening since: i) the block induced by either 1 mM or 20 mM Ba 2+  is qualitatively similar and ii) charge screening would be expected to cause a fast change in current amplitude both when Ba 2+  is applied and when it is removed (see Hurst et al., 1995). Using three different approaches, a very rough estimate of ~0.001 s −1  (with a large standard deviation) was obtained for k−2, the rate constant for unblocking of the deep site from closed channels at −80 mV with nominally zero K+o (Figs. 2.1D, E, 2.3). The apparent off-rate from the deep site was substantially increased either by raising the frequency of 10 ms test steps to +50 mV (Fig. 2.1E), by increasing the test pulse duration (Fig. 2.6B), or by increasing the magnitude of depolarizing pulses (Fig. 2.6D). Indeed, extrapolation of the open channel dissociation rate (Fig. 2.6D) for Ba 2+  to −80 mV, assuming δBa,off = 0.51, gives an estimate that is 72 more than 200 times faster than the measured resting state off rate at −80 mV (Fig. 2.1E). This facilitation of Ba 2+  unbinding by depolarization, which is consistent with results from studies on K +  channels of the squid giant axon (Armstrong et al., 1982), BK channels (Neyton & Miller, 1988b) and ShakerIR (Harris et al., 1998), is likely due to a combination of several factors: i) the deep site is in the pore and therefore “senses” part of the electric field so that depolarization decreases the on-rate and/or increases the off-rate; and, ii) channel opening, which for Kv1.5 is maximal at voltages  +20 mV (Kehl et al., 2002), may cause a conformational change of the binding site and may also permit a knock-off effect by intracellular K +  ions entering the pore. The Ba 2+  unbinding rate from open channels at +50 mV was slowed by increasing [K + ]o (see Fig. 2.6E, F). Interestingly, the apparent Kd (1 mM) for the inhibition of Ba 2+  unbinding by K + o is similar to the Kd (1 mM) for the antagonism by K + o of the low pH-induced loss of channel availability (Kehl et al., 2002), suggesting that the same K + -binding site may be involved in both situations. This site is perhaps homologous to the “lock-in” binding site for the K+-dependent inhibition of Ba 2+  unbinding in ShakerIR and BK channels (Neyton & Miller, 1988b; Hurst et al., 1995; Harris et al., 1998). Indeed, the affinity of the lock-in site for K +  in ShakerIR (Kd = 0.75 mM ; Harris et al., 1998) is quantitatively similar to what we observed in Kv1.5. It should be noted that we measured the Kd of the lock-in site in Kv1.5 with a Ba 2+  ion already bound to the deep site, as was done in ShakerIR. The actual affinity of the lock-in site for K +  may therefore be underestimated, due to the possible coulombic interaction with a bound Ba 2+  ion. 2.4.2 Ba2+ block of Kv1.5 versus ShakerIR The divergence of the KBa,d estimates for Kv1.5 and ShakerIR can probably be accounted for, at least in part, by the test pulse frequency and duration, but there are other significant differences in the Ba 2+  block of Kv1.5 and ShakerIR channels. The fast phase of Ba 2+  block, which is not seen in Kv1.5, arises because Ba 2+  (KBa,s (0 mV) = 19 mM, from  Hurst et al., 1995) 73 causes a rapidly equilibrating (flickery) block of open channels during test pulses used to monitor the development of block. We attribute the absence of a fast phase of Ba 2+  block in Kv1.5 to the comparatively much weaker open channel block. Given the estimate of 1 mM for the KBa,s of the superficial site, this weak open channel block is perhaps unexpected. For example, if this binding site is assumed to be outside the electric field, then 20 mM Ba 2+  would be expected to cause 95% inhibition when applied, as in Fig. 2.5, to open channels. If it is also assumed, for simplicity, that the superficial site that contributes indirectly to the slow block also accounts for the fast open channel block, then possible explanations for the weak open channel block are that the transition to the open conformation changes the site‟s affinity for Ba2+ or that the efflux of K +  through the open pore competitively antagonizes the fast Ba 2+  block. In support of the K + -inhibition of Ba 2+  block, with 3.5 mM K + o only ~40% of channels were blocked at rest by 1 mM Ba 2+ , compared to ~90% with 0 K + o (measured using 10 ms pulses to +50 mV at 0.033 Hz; data not shown). The effect of K +  is likely to be even greater when the channels are open since it has been suggested that, even when the bulk [K + ]o is zero, outward K +  current through ShakerIR channels leads to an accumulation of K +  in the outer pore mouth resulting, in a [K + ] of ~15 mM (Baukrowitz & Yellen, 1995). Although having identical primary sequences from the N-terminus of the pore helix through to the GYGDM sequence of the selectivity filter (see Fig. 2.2), the much weaker open- channel Ba 2+  block of Kv1.5 points to significant structural differences between Kv1.5 and Shaker. The residue at position 449 in the outer pore mouth, which is threonine in wt Shaker, can be an important molecular determinant of Ba 2+  block in ShakerIR (Hurst et al., 1996) and it seems reasonable to propose that the presence in Kv1.5 of arginine (R487) at the position homologous to Shaker T449 contributes to their divergent responses to Ba 2+ . Interestingly, in Kv1.3 the mutation H399K, which is the positional equivalent of Kv1.5 R487, causes a marked 74 decrease of block (Somodi et al., 2004; Somodi et al., 2008), implying, albeit counter-intuitively, that a basic side chain enhances Ba 2+  loading. In the light of the latter observation, the fact that onset of the slow block in Kv1.5 (block ≈ 6 s at −80 mV with 0 K + o and 20 mM Ba 2+ ; Fig. 2.1D) is more than an order of magnitude faster than in ShakerIR (block ≈ 100 s at –90 mV with 2 mM K + o and 20 mM Ba 2+ ; Fig. 9 of (Hurst et al., 1995)) might also be due to the arginine at position 487. 2.4.3 Low pH prevents movement of Ba2+ to and from its deep binding site In support of an inactivation-mediated constriction of the outer pore, the movement of Ba 2+  from its deep pore binding site to the bulk solution is significantly slowed in slow inactivated ShakerIR channels, to the extent that Ba 2+  is effectively trapped within the pore of slow inactivated ShakerIR (Basso et al., 1998; Harris et al., 1998). We have shown here that with external acidification, external Ba 2+  can neither access the deep pore binding site at rest (Fig. 2.7A), nor can Ba 2+ that is already bound at the deep site in the pore leave that binding site (Fig. 2.7B). These results are therefore consistent with the hypothesis that an outer pore constriction, perhaps similar to that proposed to underlie slow inactivation, also underlies the H + - mediated decrease in Ba 2+  access to, or egress from, the pore. 2.4.4 Is the H+-induced pore constriction indicative of resting inactivation? It remains an open question whether the proposed constriction of the outer pore of a closed Kv1.5 channel under resting, acidic conditions is the same conformation the channel enters during slow inactivation, which is typically thought to be strongly coupled to channel activation. Unfortunately, we were unable to assess the Ba 2+  block of slow inactivated channels at pH 7.4 because the inactivation is slow and incomplete, even with long depolarizing pulses (Kehl et al., 2002). While the molecular mechanism for the low pH-induced block of Kv1.5 is 75 still unclear, we (Kehl et al., 2002) and others (Steidl & Yool, 1999; Jäger & Grissmer, 2001) have proposed that the H463 residue in the outer turret acts as the putative pH sensor, since both the decrease in peak current amplitude and the enhanced depolarization-induced current decay are substantially diminished in Kv1.5 H463Q. The ability of the non-inactivating Kv1.5 R487V mutant, which is analogous to Shaker T449V, to mitigate the effects of low pH block are suggestive of an inactivation mechanism involving this residue, but the nature of the coupling mechanism between protonation of H463 and channel inactivation is unknown at this time. Acidosis of cardiac muscle is a common outcome of a number of pathological conditions, such as myocardial ischemia, and can induce arrhythmias (Orchard & Cingolani, 1994). Although the extent of extracellular acidification in native cardiac tissue is seldom likely to be as low as pH 5.5, the pKa of the H + -block of Kv1.5 in 5 mM K + o is ~6.2 (Kehl et al., 2002; Kwan et al., 2006), such that even small decreases in pH can result in a decrease in channel availability and faster macroscopic inactivation. Thus, pathological cardiac acidosis may be associated with reduced availability of Kv1.5 channels due to resting inactivation that, coupled with faster inactivation of the remaining channels and a right-shift of the activation curve (Andalib et al., 2004), results in reduced IKur.  76 3 Kinetic analysis of the effects of H+ or Ni2+ on Kv1.5 current shows that both ions enhance slow inactivation and induce resting inactivation. 3.1 INTRODUCTION As reviewed in Chapter 1, external H +  and Ni 2+  have two major effects on Kv1.5: an acceleration of the current decay during a pulse and a reduction of the peak macroscopic current (Steidl & Yool, 1999; Jäger & Grissmer, 2001; Kehl et al., 2002; Trapani & Korn, 2003; Kwan et al., 2004). We have proposed that these effects represent, respectively, an enhancement of slow inactivation at depolarized potentials and a loss of channel availability at −80 mV, which is defined here as resting inactivation. The finding in Chapter 2 that low pH inhibits the movement of Ba 2+  ions between the external solution and a binding site deep in the pore is suggestive of a H + -induced outer pore constriction occurring at rest and supports the hypothesis that slow inactivation may be occurring at −80 mV. However, there is little information about the kinetics of the onset of and recovery from resting inactivation and how they relate to those of slow inactivation. Presumably, if resting inactivation involves the same outer pore inactivation process as that underlying slow inactivation, the time courses for recovery from both the resting inactivated and slow inactivated states should be similar. In the experiments described here we use fast perfusion changes at resting and depolarized potentials to examine the possibility of a mechanistic relationship between the apparent enhancement of slow inactivation and the resting inactivation following exposure to H +  or Ni 2+  ions, and also assess whether either effect represents a modulation of the slow inactivation observed at pH 7.4. (For a detailed definition of the terms used to describe the inactivation processes under control and low pH or Ni 2+  conditions, please see the Methods, Section 3.2.4.) The approach involves a comparative analysis of the kinetics of the onset of and 77 recovery from the loss of current evoked by H +  or Ni 2+  either at rest (−80 mV) or during a depolarizing pulse (+50 mV), as well as the recovery time course following control (pH 7.4) slow inactivation at +50 mV. We conclude that the H + - or Ni 2+ -induced acceleration of current decay at +50 mV is most simply explained by a concentration-dependent enhancement of slow inactivation, but that H +  biases slow inactivation to a state from which recovery is fast and Ni 2+  biases slow inactivation to a state from which recovery is slow. The H + - or Ni 2+ -induced decrease in peak test current and maximal conductance is due primarily to the relatively slower process of resting inactivation. Recovery from the H + -induced resting inactivation has a time course similar to that for recovery from the H + -enhanced slow inactivation, suggesting that both effects are linked to a similar conformational change. 78 3.2 METHODS 3.2.1 Cell preparation As described in Chapter 2, currents were recorded from HEK-293 cells constitutively expressing wt hKv1.5 channels. Cells were dissociated for passage using trypsin-EDTA and maintained at 37°C in an atmosphere of 5% CO2 in air and in MEM supplemented with 10% foetal bovine serum, 1% (v/v) penicillin-streptomycin, and 0.5 mg ml −1  geneticin. All tissue culture supplies were obtained from Invitrogen (Burlington, Ontario, Canada). 3.2.2 Recording solutions The standard, nominally K + -free bathing solution contained (in mM): 143.5 NaCl, 2 CaCl2, 1 MgCl2, 5 glucose, 10 HEPES and was titrated at room temperature to pH 7.4 with NaOH. Bath solutions containing K +  were prepared by substituting KCl for NaCl. Low pH solutions had the same ionic composition as the standard solution, except for the replacement of HEPES by MES. Ni 2+  solutions were prepared by dilution of a 1 M NiCl2 stock solution with the standard pH 7.4 bath solution. At pH 7.4, the concentration of Ni 2+  that could be used was limited to ≤10 mM by virtue of the solubility product for Ni(OH)2 (~2 x 10 −16 ). The standard patch pipette solution contained (in mM): 130 KCl, 4.75 CaCl2 (pCa 2+  = 7.3), 1.38 MgCl2, 10 EGTA, 10 HEPES and was adjusted to pH 7.4 with KOH. When [K + ]i was decreased to 35 mM, 10 mM NaCl and 109 mM N-methyl-D-glucamine were used to replace KCl. Chemicals were obtained from the Sigma Aldrich Chemical Co. (Mississauga, Ontario, Canada). 3.2.3 Fast solution exchange Rapid changes of the external solution were typically made using a gravity-driven perfusion system in which reservoirs filled with different test solutions were connected to a custom fast application tool (FAT) constructed from polyimide-coated fused silica tubes 79 (0.32 mm ID; Agilent Technologies Canada, Inc., Mississauga, Ontario), which were glued together and fed into a common outlet capillary (0.45 mm ID; see Appendix). While the bath was perfused with control solution (≈2 ml min−1), the FAT outlet was positioned approximately 50 µm from the target cell so that it was exposed to solution flow from the FAT alone. The timing of solution changes was controlled using software-driven transistor-transistor logic (TTL) pulses that switched solenoid valves regulating solution flow between the reservoirs and the FAT. Experiments (not shown, but see Appendix) in which the [K + ]o was changed from 0 to 140 mM during a pulse to 0 mV showed a latency of 50 - 100 ms and a time constant for current decay to the zero current level of 50 – 100 ms. Faster solution changes with shorter latencies were possible but tended to disrupt the whole cell recording. 3.2.4 Definitions of terms Figure 3.1 shows simplified schemes, based on evidence summarized in Chapter 1, of the putative actions of H +  and Ni 2+  ions on Kv1.5 channels at rest (−80 mV; SCHEME 3.1) and at +50 mV (SCHEME 3.1). 80  Figure 3.1. Simplified gating schemes describing the putative actions of H +  or Ni 2+  ions on Kv1.5 channels at resting and depolarized potentials. First order rate constants are shown as kxy, where x and y denote the states (identified by the subscripts 0, 1, 2 or 3) involved in the transition. The concentration of H +  or Ni 2+  ions, also known as the ligand, is shown as [L]; -L denotes a ligand-bound state. SCHEME 3.1: At rest (−80 mV) channels in the upper row are in the available (A or A-L) state and able to pass current during test depolarizations to +50 mV. Channels in the lower row are in the unavailable or resting inactivated state (U or U-L) and remain non-conductive during test depolarizations. Resting inactivation is defined as the transition at −80 mV from the A to U state or from the A-L to U-L state. See Section 3.2.4 for details. SCHEME 3.2: At +50 mV channels are either in the open (O or O-L) and conducting state or in the open-but-slow-inactivated (OI or OI-L) and non-conducting state. Slow inactivation is defined as the transition at +50 mV from the O to OI state, while H + -or Ni 2+ -enhanced slow inactivation is defined as the transition from the O-L to OI-L state. See Section 3.2.4 for details.  81 In SCHEME 3.1 the upper row represents channels in the available (A0 or A-L1) state, meaning that the channel is able to conduct current once the activation gate opens during a test depolarization. The lower row represents channels in the unavailable (U3 or U-L2) or resting inactivated state, meaning that channels remain non-conductive even if the activation gate opens. Resting inactivation is defined here as a loss of channel availability occurring at −80 mV, a potential at which channels are assumed, on the basis of gating current measurements in Kv1.5 (Wang & Fedida, 2001), to be predominantly in the fully deactivated state. Downward and upward transitions in SCHEME 3.1 represent resting inactivation and recovery from resting inactivation, respectively. Binding and unbinding of Ni 2+  or H +  ions, which is also referred to as ligand (-L) binding/unbinding, is represented by the rightward and leftward transitions, respectively, in each row. Under control conditions (pH 7.4, 0 Ni 2+ ), the channels gate primarily in the left column of states (i.e. A0  U3) and the reaction equilibrium is strongly biased to the available state. Conversely, the reaction equilibrium of the ligand-bound or right column of states (i.e. A-L1  U-L2) is strongly biased towards inactivation and becomes more dominant as the ligand concentration  increases. As described in greater detail below, the availability at −80 mV was typically monitored with 20 ms test pulses to +50 mV. SCHEME 3.2 outlines an analogous, simplified relationship for the slow inactivation of channels at +50 mV. We define slow inactivation, sometimes referred to as control slow inactivation, as the current decay observed at +50 mV in control pH 7.4, 0 Ni 2+  solution. In this scheme channels are either open and conducting (O0 or O-L1; upper row) or open-but-slow- inactivated (OI3 or OI-L2; lower row). In both columns the gating reaction is biased towards inactivation, but more strongly so for ligand-bound channels. Inactivation was assessed during a seconds-long step to +50 mV, either in the continuous presence of control solution or during a transient application of ligand (low pH or Ni 2+ ). For consistency with the terminology used 82 previously by us and others (Steidl & Yool, 1999; Kehl et al., 2002; Kwan et al., 2004), we refer to the low pH- or Ni 2+ -induced acceleration of current decay at +50 mV as enhanced slow inactivation, and will return in the Discussion (Section 3.4) to a consideration of whether this terminology is justified. 3.2.5 Signal recording and data analysis In an experiment, a section of glass coverslip to which cells had attached was placed in the recording chamber. Whole-cell currents were recorded at room temperature (20 - 25°C) using an EPC-8 patch-clamp amplifier and PatchMaster software (HEKA Electronik, Lambrecht/Pfalz, Germany), via an ITC-18 digital interface (Instrutech, HEKA Electronik), which also provided the TTL pulses that synchronized the fast solution change with voltage step commands. Patch electrodes were made from thin-walled 1.2 mm (OD) borosilicate glass (World Precision Instruments, Sarasota, FL) and had resistances of 1.0 – 2.5 MΩ, measured in the bath with standard external and internal solutions. The circuitry of the amplifier was used to compensate the membrane capacitance and ≈80% of the series resistance. Where applicable, leak subtraction was performed using the online P/N protocol in PatchMaster, for which the holding potential was −100 mV and the scaling factor was −0.25. Current signals were low-pass filtered at 3 kHz (−3dB, 8 pole Bessel filter) and digitized (18 bit) at a sampling frequency of at least 10 kHz. Voltages were corrected for liquid junction potentials. Holding and test potentials of −80 mV and +50 mV, respectively, were standard. When applicable, current amplitudes were normalized to control measurements made at the beginning of the sweep or experiment. Data from cells showing <80% recovery to control levels after test treatments were discarded. The similarities between the time courses of recovery from slow inactivation and H + - or Ni 2+ -enhanced slow inactivation or resting inactivation were assessed using a simultaneous, or global, fit of data sets to single or double exponential functions. This was performed using the 83 built-in Global Fit function of our analysis software (IGOR Pro, Wavemetrics, Portland, OR), in which the fit was constrained such that the time constants of the fitting function were the same for each data set, but the relative amplitudes of the fast and slow recovery components (Arec,f, Arec,s) and the initial level of availability (A0) were allowed to vary. The chi-square (χ 2 ) statistic was used to test for goodness of fit. For other comparisons, a one-way ANOVA and Tukey test were used to test for statistical significance (P < 0.05). Data are presented as mean ± S.E.M.; n represents the number of cells tested. 3.2.6 Numeric simulation Macroscopic Kv1.5 currents were simulated in IGOR Pro by calculating state occupancies as a function of time, voltage and ligand concentration from the spectral expansion of the Q- matrix (Colquhoun & Hawkes, 1995) generated from state diagrams described in the text and associated figure legends. 84 3.3 RESULTS 3.3.1 External H+ and Ni2+ enhance the current decay and reduce peak current Figure 3.2A displays a current trace obtained using a 5 s pulse from −80 mV to +50 mV in 0 mM K +  solution at pH 7.4. The voltage protocol was then repeated following prolonged exposure to K + -free bath solutions at pH 6.9 and 6.4. As reported by others for Kv1.5 and ShakerIR (Meyer & Heinemann, 1997; Rich & Snyders, 1998; Jäger & Grissmer, 2001), the time course for slow inactivation at pH 7.4 was better fitted to a double exponential. Nonetheless, a single exponential function was used to facilitate a comparison with the current decay at low pH, which was typically mono-exponential. Low pH decreased the peak macroscopic current and the time constant for slow inactivation (τinact; Fig. 3.2A). The small amplitude and non-inactivating time course of the residual current at pH 6.4 point to it being due to endogenous HEK-293 channels (Zhu et al., 1998; Lambert & Oberwinkler, 2005). In 0 K + o the pKa for the decrease in peak current was ≈6.9 (see figure legend), in agreement with our previous report (Kehl et al., 2002). Increasing [K + ]o from 0 to 3.5 mM partially relieved the inhibitory effects of H + , so that substantial current was observed even at pH 5.9 (Fig. 3.2B). Qualitatively similar observations with Ni 2+  in 0 and 3.5 K + o are shown in Fig. 3.2C and D. These results confirm the previous findings that H +  and Ni 2+  decrease the peak macroscopic current and accelerate the current decay during a pulse in a concentration- and [K + ]o-sensitive manner (Steidl & Yool, 1999; Jäger & Grissmer, 2001; Kehl et al., 2002; Kwan et al., 2004). However, the experiment does not distinguish between the reduction in peak current due to enhanced slow inactivation and that due to ligand-induced resting inactivation (see Fig. 3.1). To address this issue, fast solution changes were used to examine the effects of H +  and Ni 2+  on either open or resting Kv1.5 channels. 85  Figure 3.2. Effects of external H +  and Ni 2+  on Kv1.5 current. In all panels, superimposed grey traces represent current recorded during 5 s voltage steps to +50 mV from −80 mV following prolonged exposure (20 – 40 s) to low pH or [Ni2+]. Black lines represent the fits of the current decay to a mono-exponential function with a time constant denoted by τinact. Each panel represents an experiment on a different cell. A, In 0 mM K + o, low pH decreases both peak Kv1.5 current and τinact. Pooled data from 4 cells gave a pKa of 6.9 and a Hill coefficient of 1.4. B, Increasing [K + ]o antagonizes the effect of low pH on peak test current. In 3.5 mM K + o the pKa for the decrease in peak current was 6.2, with a Hill coefficient of 1.8 (n = 4 cells). C, Ni 2+  also causes a concentration-dependent decrease in peak current but this is associated with relatively smaller decreases of the test current decay rate. From 4 cells, the fit of the [Ni 2+ ]-dependence of the mean normalized peak current amplitude to the Hill equation gave Kd and Hill coefficient values of 0.034 mM and 0.85, respectively. D, In 3.5 mM K + o the Kd for the [Ni 2+ ]-dependent decrease of peak current amplitude was 0.52 mM; the Hill coefficient was 1.2 (n = 4 cells). For reasons unclear to us, these Kd values for the Ni 2+  effect are lower than those previously reported (Kwan et al., 2004). 86 3.3.2 Decreasing external pH during a depolarization enhances slow inactivation To isolate the effects of low pH on open channels, we used fast perfusion to rapidly decrease pH during a step to +50 mV (e.g. Fig. 3.3A). With this protocol the reduction of peak current that occurs with prolonged application of low pH solutions (Fig. 3.2) is avoided and the focus is on the effects of H +  on the open state and gating transitions as outlined in SCHEME 3.2 of Fig. 3.1. Figure 3.3A shows superimposed current traces recorded from the same cell in K + -free medium at pH 7.4 (control) and with transient exposures to pH 6.9 and pH 5.9. As in ShakerIR (Starkus et al., 2003), decreasing pH during a depolarizing pulse enhanced Kv1.5 current decay. The mean values for inact measured in this way at pHs ranging from 7.4 to 5.4 in 0 K + o medium are plotted in Fig. 3.3F (); the foot of the concentration-response relationship at 100 - 150 ms may reflect the limiting rate of the solution exchange (see Methods, Section 3.2.3). These results support the hypothesis that low pH enhances slow inactivation of open channels, as outlined in SCHEME 3.2. If it is correct that H +  enhances slow inactivation, then an increase of the rate of slow inactivation at pH 7.4 should be paralleled by an increased rate of current decay at low pH. This was tested by taking advantage of the fact that slow inactivation at pH 7.4 is accelerated roughly two-fold when the [K + ]i is reduced from 130  to 35 mM (compare  and  at pH 7.4; Fig. 3.3F), an effect that has been attributed to a decreased flux through the open channel and a consequent reduction of the occupancy of the outer pore site that controls slow inactivation (Fedida et al., 1999; Ogielska & Aldrich, 1999). As predicted, with 35 mM K + i the current decay observed with rapid switching to pH 6.4 at +50 mV was approximately 2.5-fold faster than with 130 mM ( and  at pH 6.4; Fig. 3.3F). 87  Figure 3.3. The time course of enhanced slow inactivation and resting inactivation is pH dependent. A, Superimposed current traces from the same cell showing the enhancement of current decay by low pH. The 0 mM K+ bathing solution was rapidly and transiently switched from pH 7.4 to low pH during a 20 s step from −80 mV to +50 mV. Dashed lines represent mono-exponential fits of the current decay with τinact values of 4.88 s, 1.11 s and 0.25 s at pH 7.4, 6.4 and 5.9, respectively. B, Current trace showing the onset of resting inactivation induced at pH 5.4 in 0 mM K + o. After a 20 ms control pulse to +50 mV at pH 7.4, the external pH was decreased and a pulse train with increasing interpulse intervals was applied during a single sweep. C, Resting inactivation was also assessed with multiple (superimposed) sweeps. In each sweep a single test pulse was applied at a known interval after the switch to pH 5.4; the interval was increased for each successive sweep. The final, control, trace was obtained without prior exposure to low pH. D, The onset of resting inactivation induced by pH 5.4 in 3.5 mM K + o was measured as described for B. E, Test currents from B - D were normalized to their respective controls and plotted against the cumulative time spent at −80 mV at pH 5.4. Inset, The same plot on a longer timescale shows the steady-state level. The solid and dashed lines represent mono- and bi-exponential fits of the data, respectively; τRI at pH 5.4 in 0 K + o was 171 ms and 122 ms measured with a train and single pulses, respectively. In 3.5 mM K + o at pH 5.4, τRI,fast and τRI,slow were 131 ms and 1.24 s, respectively. F, Time constants for low pH enhanced slow inactivation and resting inactivation derived from experiments such as those in A and B are plotted against pH. All data points represent the mean ± S.E.M. from at least 3 cells. 88 In agreement with previous reports that the time course of slow inactivation of open Kv1.5 channels is insensitive to moderate changes in [K + ]o (Fedida et al., 1999; Jäger & Grissmer, 2001), the τinact at pH 6.4 was unaffected by increasing [K + ]o from 0 to 3.5 mM (Fig. 3.3F; ). Thus, as with slow inactivation at pH 7.4, the H+-enhanced current decay is sensitive to [K + ]i and insensitive to [K + ]o, which is consistent with our earlier conclusion that external H +  acts on open Kv1.5 channels to enhance slow inactivation during a depolarizing pulse. 3.3.3 Current can recover from a transient decrease in pH during a depolarization In contrast to findings with ShakerIR where there is little or no recovery after returning to pH 7.4 (Starkus et al., 2003), in Kv1.5 the transition from low pH back to pH 7.4 during a depolarizing pulse is associated with a substantial recovery current (Fig. 3.3A). Recovery was occasionally to the same level as the inactivating control current, but in many cells (Fig. 3.3A) the recovery current crossed and peaked above the control level. In Fig. 3.3A the rising component of the biphasic recovery current following either the pH 6.9 or the pH 5.9 pulse was well-fitted by a single exponential (τ ≈ 1 s in each case). The declining component of the recovery current typically relaxed back to the control current level provided the pulse was long enough. Importantly, single channel studies with both Kv1.5 (Kwan et al., 2006) and ShakerIR (Claydon et al., 2007) have precluded rapid open channel block as the mechanistic basis for the enhanced decay of current at low pH. As such, the recovery current cannot be ascribed to the reversal of channel block. Instead, the increase of current observed after returning to pH 7.4 solution may reflect channel deprotonation (OI-L2 → OI3 of SCHEME 3.2 of Fig. 3.1) and subsequent recovery from inactivation (OI3 → O0). A further examination of recovery currents at +50 mV is presented below. 89 3.3.4 The onset of low pH-induced resting inactivation is slow The time course of low pH-induced resting inactivation of Kv1.5 at −80 mV was next characterized, on the premise that an analysis of the onset and/or recovery kinetics could help elucidate the mechanistic relationship, if any, between slow inactivation at +50 mV and resting inactivation at −80 mV. The protocol involved a 20 ms control step to +50 mV in pH 7.4 solution applied just prior to the beginning of a 4 - 60 s exposure to low pH solution, during which a train of 20 ms test pulses to +50 mV was applied. For the trace in Fig. 3.3B, K + -free pH 5.4 solution was applied for a 4.3 s duration, during which test pulses were applied. There was a progressive decline of peak test currents as the duration of exposure to pH 5.4 solution increased. A plot of the normalized peak current amplitude against the time spent at low pH at −80 mV (▲; Fig. 3.3E) shows that the onset of H+-induced resting inactivation in K+-free medium is well fitted by a single exponential, with a time constant defined as τRI. Despite the enhanced slow inactivation during test pulses to +50 mV at pH 5.4 (Fig. 3.3A), the brevity of the test pulses in this protocol precluded accumulation of slow inactivation as a contributing factor in the decline of the test currents. This is confirmed in Fig. 3.3C, which shows that the onset of resting inactivation at pH 5.4 is no different when assessed with single test pulses (in multiple, superimposed sweeps) applied after known intervals from the start of the solution change (Fig. 3.3E; compare  and ). The mean values for τRI measured with a train of pulses (e.g. Fig. 3.3B; 161 ± 15 ms; n = 4) and single pulses (e.g. Fig. 3.3C; 177 ± 17 ms; n = 5) are not significantly different (P > 0.05). Figure 3.3F () summarizes the pH dependence of τRI in 0 K + o measured with pulse trains and also indicates that over the pH range tested, the onset of H + - induced resting inactivation was typically slower than for H + -enhanced slow inactivation, except at pH 5.4 where the τRI and τinact converge, possibly due to the limiting rate of the solution change. 90 3.3.5 Low pH-induced resting inactivation is sensitive to [K+]o but not [K + ]i Because extracellular K +  ions antagonize the conductance collapse, or resting inactivation, of Kv1.5 induced by either low pH or Ni 2+  (Fig. 3.2; Jäger & Grissmer, 2001; Kehl et al., 2002; Kwan et al., 2004), it was of interest to determine how the time course of H + - induced resting inactivation was affected by increasing [K + ]o. With 3.5 mM K + o a major kinetic change was that the onset of resting inactivation was bi-exponential (Fig. 3.3D, E). To facilitate a direct comparison of the time course of resting inactivation in either 0 or 3.5 mM K + o, we calculated the time required for the current to relax to 50% (T50) and 90% (T90) of the maximum current decay. At pH 5.4 with 0 mM K + o, T50 and T90 were 0.11 ± 0.01 s and 0.35 ± 0.03 s (n = 6), respectively, and with 3.5 mM K + o the corresponding values increased to 0.19 ± 0.04 s and 1.66 ± 0.33 s (n = 5). The nearly five-fold increase of T90 with 3.5 K + o reflects the substantial contribution of the slower process to resting inactivation and is consistent with the inhibition by K + o of an inactivation process involving the outer pore. In contrast to enhanced slow inactivation measured at +50 mV, the time course of pH 6.4-induced resting inactivation in 0 K + o was not affected by reducing [K + ]i to 35 mM (Fig. 3.3F; ), as would be expected since the closed activation gate, which is located on the cytoplasmic side of the channel, would preclude an effect of [K + ]i on the occupancy of the permeation pathway. 3.3.6 Transient exposure to Ni2+ during a pulse enhances slow inactivation Figure 3.4A shows that, like low pH in Fig. 3.3A, the fast application of Ni 2+  shortly after the start of a pulse to +50 mV results in an enhancement of the rate and extent of current decay. The concentration dependence of this effect at pH 7.4 with 0 mM K + o is summarized in Fig. 3.4E () and indicates that the Ni2+ effect was qualitatively the same as that for low pH. There were additional shared properties of the two ligands. Changing from 0 to 3.5 mM K + o had no effect on τinact measured with 2 mM Ni 2+  (; Fig. 3.4E), and decreasing control inact (in 0 mM Ni 2+ ) by 91 changing [K + ]i from 130 to 35 mM was associated with a 2-fold faster rate of inactivation in 2 mM Ni 2+  (; Fig. 3.4E). Current recovery was also observed following the return to Ni2+-free medium during the pulse, but the recovery time course, although variable, was typically much slower than that following low pH. Thus, like H + , Ni 2+  enhances slow inactivation of Kv1.5 channels at +50 mV. 92  Figure 3.4. The time course of enhanced slow inactivation and resting inactivation is also [Ni 2+ ]-dependent. A, Superimposed current traces from the same cell showing the enhancement of current decay by external Ni2+. During a 20 s step from −80 mV to +50 mV, the control (pH 7.4, 0 mM K+) bath solution was rapidly and transiently switched to one containing Ni2+. Dashed lines represent mono-exponential fits of the current decay with τinact values of 4.08 s, 1.65 s and 0.34 s in 0, 0.1, and 10 mM Ni2+, respectively. B, Current trace showing the onset of Ni2+-induced resting inactivation in 0 mM K+o. After a 20 ms control pulse from −80 mV to +50 mV in 0 mM Ni 2+ , resting inactivation was assessed by switching to 10 mM Ni 2+  solution and applying a pulse train with increasing interpulse intervals. C, The experimental protocol in B was repeated, albeit on a different timescale, with 3.5 mM K + o. D, Test currents from B and C were normalized to their respective controls and plotted against the cumulative time spent at −80 mV in Ni2+. For comparison, results from an experiment using 0.05 mM Ni 2+  are also shown. Solid and dashed lines represent mono- and bi-exponential fits of the data, respectively. E, Time constants for Ni 2+ -enhanced current decay and resting inactivation derived from experiments such as those in A and B are plotted against [Ni 2+ ]. Data points represent the mean ± S.E.M. of 3 – 7 cells. 93 3.3.7 The time course of Ni2+-induced resting inactivation is slow The onset of Ni 2+ -induced resting inactivation in 0 K + o at pH 7.4 was characterized with the same single sweep protocol as for low pH. For the current trace in Fig. 3.4B, 10 mM Ni 2+  was present for the duration indicated by the lower horizontal bar; the peak test pulse current decreased exponentially as the duration of exposure to Ni 2+  at −80 mV increased. A plot of the normalized test current amplitudes for the trace of Fig. 3.4B and a similar experiment with 50 µM Ni 2+  show that the extent and the rate of inactivation are concentration dependent (Fig. 3.4D). At a given [Ni 2+ ], τRI was the same regardless of whether a single sweep comprising a train of test pulses or several sweeps of single pulses (cf. Fig. 3.3C) were used (not shown), confirming that the effect is not due to an accumulation of slow inactivation. Figure 3.4E () summarizes the [Ni 2+ ]-dependence of τRI in 0 K + o. The data mimic the pattern observed with H +  where the value for τRI approaches that for τinact as the [Ni 2+ ] increases. As observed with low pH, the time course of Ni 2+ -induced resting inactivation was bi-exponential when [K + ]o was increased from 0 to 3.5 mM (Fig. 3.4C, D). With 10 mM Ni 2+  the T50 and T90 values of the maximum current decay increased from 0.43 ± 0.01 s and 1.42 ± 0.04 s (n = 7) in 0 mM K + o to 1.28 ± 0.16 s and 9.17 ± 0.24 s (n = 8) in 3.5 mM K + o. Thus, consistent with the properties of an inactivation process involving the pore, the addition of external K +  dramatically slows the onset of Ni 2+ -induced resting inactivation. In 2 mM Ni 2+  there was no significant difference between τRI with 130 versus 35 mM K + i (Fig. 3.4E), which again mirrors the finding with low pH (Fig. 3.3F). Further support for our contention that resting inactivation involves the pore is provided by the results of experiments looking at the effect of Ni 2+  on the Ba 2+  binding to and unbinding from Kv1.5 channels. Figure 3.5A shows experiments where, in a manner similar to that described for Fig. 2.7, the changes in current amplitude in response to applications of 5 mM Ni 2+  and/or 20 mM Ba 2+  were monitored 94 with brief, repetitive pulses. The uppermost diary plots in Fig. 3.5A show the onset of and recovery from decreases in current induced by either Ni 2+  or Ba 2+ . The mono-exponential time course of recovery from Ba 2+  exposure is significantly slower than that of the Ni 2+ -induced loss of current (P <0.05; Fig. 3.5B). The bottom diary plots of Fig. 3.5A show that when Ni 2+  is applied prior to Ba 2+ , current recovery upon wash-out (of both Ba 2+  and Ni 2+ ) occurs with a time constant similar to that for recovery from Ni 2+  exposure alone, consistent with an inability of Ba 2+  to access its deep binding site and slow current recovery. Conversely, if the Kv1.5 channels are exposed to Ni 2+  immediately following Ba 2+  treatment, the time course for recovery upon wash-out was not different from that for recovery from Ba 2+  treatment alone (P > 0.05; Fig. 3.5B), indicating that Ba 2+  did not unbind from the channel during the pulses applied in 0 Ba 2+ , Ni 2+ -containing solution. These results are reminiscent of those observed with low pH and Ba 2+  (Fig. 2.7) and indicate that Ni 2+  can impede the movement of Ba 2+  to and from a binding site within the pore, suggestive of a constriction in the outer pore mouth and consistent with the properties of slow inactivated ShakerIR channels (Basso et al., 1998; Harris et al., 1998).  95  Figure 3.5. Ni 2+  prevents Ba 2+  entry and exit from the pore binding site. A, Diary plots of normalized peak current amplitude measured using 15 ms pulses to +50 mV, applied at 0.5 Hz. Currents are normalized to the average control peak current amplitude at the beginning of each experiment, before test treatments. Each plot represents a separate experiment in a different cell. The durations of exposure to 5 mM Ni 2+  and/or 20 mM Ba 2+  are indicated by the horizontal bars at the top of each plot. B, Comparison of the time constants for recovery from exposure to Ni 2+  only; Ni 2+  followed by Ba 2+ ; Ba 2+  only; and Ba 2+  followed by Ni 2+ . Ni 2+  prevents Ba 2+  from binding to Kv1.5 channels, resulting in a recovery time course from Ni 2+  followed by Ba 2+  that is not significantly different from recovery from Ni 2+  alone. Conversely, when Ba 2+  is applied first, it is trapped within the pore by subsequently applied Ni 2+ . This results in a recovery time course that is not significantly different from the recovery from Ba 2+  alone. Error bars represent S.E.M.; n-values for each data set are shown in parentheses. 96 In summary, the data presented in Figures 3.2 – 3.4 suggest that both external Ni2+ and H +  enhance slow inactivation at +50 mV in a concentration-dependent manner. Like control slow inactivation at +50 mV, neither the H +  nor the Ni 2+  enhancement of slow inactivation is sensitive to [K + ]o but the rate of inactivation with either ligand is augmented by decreasing [K + ]i. In contrast, the ligand-induced resting inactivation is inhibited by increasing [K + ]o and unaffected by decreasing [K + ]i. These properties are consistent with the conclusion that both enhanced slow inactivation at +50 mV and resting inactivation at −80 mV involve an inactivation process involving the pore. However, whether the Ni 2+ - and H + -enhanced slow inactivated and resting- inactivated conformations are mechanistically and conformationally related remains unclear. To begin to address that issue, we next turned to a comparative analysis of recovery kinetics following inactivation at +50 mV and −80 mV. 3.3.8 Recovery from H+-enhanced slow inactivation or resting inactivation is fast In this and the following section the time course of recovery following inactivation at +50 mV or −80 mV with low pH or Ni2+ is compared to that after control slow inactivation at pH 7.4. Comparisons of control and treated responses were made in the same cell, since, in our experience (e.g. Fig. 3.2; see also Rich & Snyders, 1998), there is cell-to-cell variability in the rate and extent of slow inactivation. In Fig. 3.6A single sweeps of a representative experiment show recovery following (from top to bottom): control slow inactivation (5 s at +50 mV at pH 7.4), enhanced slow inactivation (≈5 s at +50 mV at pH 5.4), and resting inactivation (5 s at −80 mV at pH 5.4). In each experimental condition, recovery from inactivation was monitored in 0 K + , pH 7.4 solution with a train of test pulses to +50 mV. 97  Figure 3.6. The kinetics of recovery from H + - or Ni 2+ -enhanced slow inactivation or resting inactivation. A, Current traces recorded in 0 mM K + o from the same cell showing: () recovery at pH 7.4 from control slow inactivation; () recovery at pH 7.4 from slow inactivation enhanced by pH 5.4 and; () recovery at pH 7.4 from resting inactivation induced by pH 5.4. The voltage clamp protocol consisted of a 5 s step to +50 mV, except for the resting inactivation trace, followed by a train of 20 ms test pulses to +50 mV. The dotted horizontal lines indicate the duration of the exposure to pH 5.4 solution. B, Peak test currents from (A), normalized with respect to the peak control current, are plotted against the cumulative recovery time spent at the −80 mV holding potential. The solid lines represent the simultaneous fit to a double exponential function of the time course of recovery from control slow inactivation (), H+-enhanced slow inactivation () and resting inactivation (). The resulting values for τrec,f and τrec,s were 1.48 s and 9.14 s, respectively. C, Normalized peak recovery currents measured with the same protocol as in (A), but with 4 mM Ni 2+  instead of low pH, are plotted against the cumulative recovery time. The recovery from control () and Ni2+-enhanced () slow inactivation were simultaneously fit to a double exponential function with τrec,f = 0.59 s and τrec,s = 10.6 s. Recovery from Ni 2+ -enhanced slow inactivation was entirely via the slow phase. The inset graph shows that recovery from Ni 2+ -induced resting inactivation () was faster than that following enhanced slow inactivation. The solid lines represent separate fits of each data set to a mono- exponential function. τrec was 10.8 s for enhanced slow inactivation and 6.7 s for resting inactivation. 98 For Fig. 3.6B the peak test current amplitudes from Fig. 3.6A were normalized to the peak control current and plotted against the cumulative recovery time at −80 mV. In agreement with previous reports, the recovery from control slow inactivation at pH 7.4 was biphasic, which is indicative of at least two inactivated states (Rich & Snyders, 1998; Perchenet & Clément- Chomienne, 2000). Preliminary independent fits (not shown) suggested a similarity of the time constants for recovery from control slow inactivation with those from enhanced slow inactivation and resting inactivation, both at pH 5.4. This motivated simultaneous or global fitting of the three data sets to a bi-exponential function in which the initial level of availability (A0) and the relative amplitudes of the fast and slow recovery components (Arec,f and Arec,s respectively) were free, while the values for the fast (rec,f) and the slow (rec,s) recovery time constants were constrained to be the same (see Methods, Section 3.2.5). The solid lines of Fig. 3.6B show that the outcome of the global fit was excellent (2 = 0.004). Table 1 summarizes the results from 6 cells and shows that: 1) during the 5 s pulse the extent (1 – A0) of either enhanced slow inactivation or resting inactivation at pH 5.4 is greater than that for control slow inactivation at pH 7.4; 2) after slow inactivation at pH 7.4 channels are roughly evenly distributed between the fast and slow recovery pathways; and, 3) recovery at pH 7.4 from low pH-enhanced slow inactivation or H + - induced resting inactivation is mostly via the fast phase (Arec,f >> Arec,s). Together, these findings imply that two major inactivated states are visited following slow inactivation at pH 7.4, and that the same states are also populated after enhanced slow inactivation or resting inactivation at pH 5.4. However, low pH biases inactivation at −80 mV and at +50 mV towards the inactivated state from which recovery is faster. 99 Table 3.1. Low pH biases inactivation to a state from which recovery is fast.  pH 7.4 pH 5.4  inactivation at +50 mV inactivation at +50 mV inactivation at −80 mV A0 0.53 ± 0.10 0.02 ± 0.01 0.08 ± 0.08 Arec,f ** 0.47 ± 0.19 0.71 ± 0.07 0.89 ± 0.04 τrec,f 1.63 ± 0.1 s *  * Arec,s ** 0.53 ± 0.19 0.29 ± 0.07 0.11 ± 0.04 τrec,s 13.52 ± 2.9 s * * * constrained to be the same as control (pH 7.4) values **  presented as proportion of total recovery current Data are from experiments described in Figs. 3.6A & B and are shown as mean ± S.E.M. for 6 cells. 100 3.3.9 Recovery from Ni2+-enhanced slow inactivation or resting inactivation is slow The time course of recovery following enhanced slow inactivation and resting inactivation in 4 mM Ni 2+  (Fig. 3.6C) was assessed using the same stimulus protocols described for Fig. 3.6A. The main panel of Fig. 3.6C shows the recovery of the normalized peak test currents, obtained from the same cell, following inactivation in control solution and in 4 mM Ni 2+  at −80 mV and +50 mV. In this cell both the extent of control slow inactivation at the end of the 5 s pulse to +50 mV and τrec,f were less than that described in Fig. 3.6B; this underscores the cell-to-cell variability of slow inactivation and the rationale for doing within-cell comparisons. In contrast to the situation following enhanced slow inactivation at +50 mV at pH 5.4 (see Fig. 3.6B), the recovery following Ni 2+ -enhanced slow inactivation was mono-exponential (; Fig. 3.6C). However, the time constant (τrec) for recovery from Ni 2+ -enhanced slow inactivation was not significantly different from rec,s following control slow inactivation (P > 0.05), and the slow component of control recovery and the mono-exponential recovery following Ni 2+  treatment were replicated in a global fit (Fig. 3.6C). In 12 cells, the recovery following control and Ni 2+ -enhanced slow inactivation could be fitted simultaneously (rec,s = τrec = 13.84 ± 0.73 s). In the same cells, recovery from Ni 2+ -induced resting inactivation (see Fig. 3.6C, inset) was also slow and mono-exponential but was consistently faster than recovery following Ni 2+ -enhanced slow inactivation and necessitated separate fits which gave a mean τrec value of 7.12 ± 0.42 s (n = 12). In summary, low pH biases inactivation towards an inactivated state from which recovery is fast. Furthermore, the time constants for recovery both from H + -enhanced slow inactivation and resting inactivation are not significantly different, implying shared recovery pathways and similar mechanistic processes. With Ni 2+ , recovery following inactivation at +50 mV is mono- 101 exponential and proceeds with the same time constant as the slow component of recovery from control inactivation at +50 mV, indicating that Ni 2+  very strongly biases inactivation at +50 mV to the more stable inactivated state visited, albeit to a smaller extent, in control solution. The recovery rates following inactivation in Ni 2+  at +50 mV and −80 mV differ by roughly 2-fold, suggesting that resting inactivation in Ni 2+  is to a state that is not visited in control solution. 3.3.10 Recovery from H+- and Ni2+-induced resting inactivation at depolarized potentials During a prolonged depolarization to +50 mV there is recovery from ligand-enhanced slow inactivation upon the return to control conditions (Figs. 3.3A, 3.4A). The results of Fig. 3.6A and B, which suggest that low pH pushes inactivation towards a state from which recovery is fast, provide a potential explanation for the recovery overshoot shown in Fig. 3.3A. That is to say, the recovery overshoot may reflect a re-equilibration between two different slow inactivated states upon the return to pH 7.4. Figure 3.7 presents results from experiments assessing a related question: whether recovery at +50 mV could also be observed following ligand-induced resting inactivation and, if so, how that recovery compared to that following enhanced slow inactivation. 102  Figure 3.7. Current recovery during a test depolarizing pulse occurs following either H + - or Ni 2+ - enhanced slow inactivation or resting inactivation. The experimental approach was similar to that used in Figure 3.6 except that the test pulse duration was longer and multiple test sweeps were used. An intersweep interval of 60 s allowed for full recovery between sweeps. For the top panels, after a 20 ms control control pulse from −80 mV to +50 mV in pH 7.4, 0 mM K+o solution, resting inactivation at −80 mV was induced by a 10 s application of either pH 6.4 (Ai) or 2 mM Ni 2+  (Bi) solution. To monitor recovery, the test solution was replaced by control solution and the cell held at −80 mV for varying intervals before a 5 s test pulse was applied. For the lower panels, recovery from slow inactivation enhanced by pH 6.4 (Aii) or 2 mM Ni 2+  (Bii) was assessed in the same way, except that low pH and Ni 2+  were applied 100 ms after the beginning of a 5 s pulse from −80 mV to +50 mV. For each column, traces were recorded from the same cell. These results show that with either ligand and following either resting inactivation (top panels) or enhanced slow inactivation (lower panels, and see also Figs. 3.3A, 3.4A) there can be the recovery of current during a depolarizing pulse and that the kinetics of that recovery are qualitatively similar for a given ligand. Dotted lines represent single exponential fits of the amplitude of the fast phase of the test current (). As shown in Figure 3.6, recovery at −80 mV following low pH exposure is faster than that following Ni 2+  exposure. 103 The superimposed traces of Fig. 3.7Ai show the current recovery following resting inactivation at −80 mV induced by a 10 s exposure to pH 6.4 solution. For the first of 5 successive sweeps there was an interval of 500 ms between the return to pH 7.4 solution and the application of a 5 s test pulse to +50 mV; for each successive sweep the interval between the return to pH 7.4 and the test pulse was increased by 2 s. The first sweep shows an initial fast rise of current (peak demarcated by ; τ ≈ 2 ms) reflecting the normal opening of channels that either did not inactivate during the pH 6.4 pulse or that had recovered during the recovery interval at −80 mV. The fast component is followed by a more slowly rising phase of current ( ≈ 553 ms). On successive sweeps the amplitude of the initial rapid component increases as more channels recover during the “interpulse” interval. However, even on the second and third test pulses there is also evidence for current recovery during the pulse, revealed either as a slow initial plateau (sweep 2) or as an inactivation rate (sweep 3) that is slower than that of the fourth and fifth test pulses. Figure 3.7Aii shows six superimposed current traces from the same cell in which slow inactivation was enhanced by switching to pH 6.4 solution 100 ms after the start of a 5 s pulse to +50 mV. Again, test pulses were applied in control solution at varying intervals following the inactivating pulse. The test currents following H + -enhanced slow inactivation are qualitatively indistinguishable from those that follow resting inactivation inasmuch as recovery at +50 mV (τ ≈ 585 ms) is prominent during the first pulse and is present, but less overt, on the second and third pulses. Figure 3.7B shows the results where, using a similar experimental approach and presentation protocol (see figure legend for details), 2 mM Ni 2+  either induced resting inactivation (panel Bi) or enhanced slow inactivation (panel Bii). As with low pH, after resting inactivation and enhanced slow inactivation in Ni 2+  the test pulse current of the first sweep has an initial rapid rise of current (peak demarcated by ; τ ≈ 3 ms). This is followed by a slowly 104 rising phase that can be fitted by a single exponential (τ ≈ 4.7 s and τ ≈ 5.1 s for resting inactivation and enhanced slow inactivation, respectively) and represents current recovery during the pulse. In subsequent sweeps, as with low pH, current recovery at +50 mV from Ni 2+ - enhanced slow inactivation or resting inactivation is apparent in the lower rate of slow inactivation during the test pulses. The results of Fig. 3.7 indicate that recovery from inactivation can occur during a depolarizing test pulse following either enhanced slow inactivation or resting inactivation with either ligand. However, the recovery kinetics following low pH or Ni 2+  treatment differ in two important aspects. As noted in Fig. 3.6 and confirmed in Fig. 3.7 by the open circles and fitted dashed lines, which represent the envelope of the fast phase of current, recovery at −80 mV from either enhanced slow inactivation or resting inactivation is much faster following low pH. Second, the recovery during a pulse at +50 mV, as shown in Figs. 3.3A, 3.4A and 3.7A is also faster following low pH. This again implies that low pH biases inactivation to a state from which recovery, either at −80 mV or +50 mV, is relatively fast, and that Ni2+ biases inactivation to a state from which recovery is slow. 3.3.11 The time course for K+o-facilitated recovery from resting inactivation is the same as that for recovery from enhanced slow inactivation As a further assessment of the relationship between ligand-enhanced slow inactivation and resting inactivation, the effect of increasing [K + ]o on the recovery from resting inactivation was examined. These experiments were also motivated by the finding in Kv1.3 H404N (the positional homologue of T449 in ShakerIR) mutant channels that the time course for recovery from the loss of current induced by removing K + o was much faster than that for recovery from slow inactivation, which was interpreted to mean that the loss of current in 0 K +  solution was not due to slow inactivation (Jäger et al., 1998). For these experiments, unlike those in Figs. 3.6 and 105 3.7 where recovery was monitored in control solution, recovery from the loss of current in 0 K + o was measured in the continued presence of H +  or Ni 2+ . This required the use of cells expressing Kv1.5 channels at a high density so that the residual current, at pH 5.4 or with 2 mM Ni 2+ , was large enough to reliably track recovery kinetics following either enhanced slow inactivation at +50 mV or the induction of a greater level of resting inactivation triggered by removing external K + . Recovery from enhanced slow inactivation at pH 5.4 in 3.5 mM K + o was measured using a single sweep comprising a train of test pulses to +50 mV applied at varying intervals after the inactivating pulse and followed a mono-exponential time course (Fig. 3.8A). For the sweep of Fig. 3.8B, which was recorded from the same cell, the control current in pH 5.4, 3.5 mM K +  solution was recorded prior to inducing a greater level of resting inactivation by switching to 0 mM K + , pH 5.4 solution for 5 s. A train of test pulses was then applied after returning to pH 5.4, 3.5 mM K +  solution. The recovery time course, which reflects the re-establishment of the control level of resting channel availability in 3.5 mM K + , was also mono-exponential and a plot of the normalized peak test currents from panels A and B against the cumulative recovery time at −80 mV (Fig. 3.8C) indicates that the two recovery time courses are quite similar. The mean time constants for recovery at pH 5.4 from enhanced slow inactivation and resting inactivation were 2.38 ± 0.15 s and 2.00 ± 0.10 s, respectively, and were not significantly different (P > 0.05; n = 3). Similar voltage and perfusion protocols were used to compare the recovery in 2 mM Ni 2+  and 3.5 mM K + o (pH 7.4) following enhanced slow inactivation at +50 mV with 3.5 mM K + o or resting inactivation at −80 mV with 0 K+o. Figure 3.8D shows the normalized peak test currents from a representative experiment. The time courses of recovery following slow inactivation and following resting inactivation in 0 K + o were mono-exponential and the mean recovery time constants of 14.85 ± 1.77 s and 17.51 ± 2.28 s, respectively, were not significantly different 106 (P > 0.05; n = 4). The similarity of the time constants for recovery from enhanced slow inactivation and 0 K + o-induced resting inactivation suggest that slow inactivation and [K + ]o- sensitive resting inactivation of Kv1.5 may involve similar kinetic processes. 107  Figure 3.8. The time course for recovery from 0 K + o-induced resting inactivation is the same as that for recovery from enhanced slow inactivation. Using cells expressing Kv1.5 channels at high density, current recordings were made in pH 5.4, 3.5 mM K +  solution. Following a 20 ms control pulse, the recovery of current in 3.5 mM K + o was monitored, in A, after a 500 ms pulse to +50 mV or, in B, after channel availability was decreased by a 5 s exposure to K + -free, pH 5.4 solution. Recovery was monitored using 20 ms test pulses applied at increasing intervals within the same sweep. C, Peak test current amplitudes from (A and B) were normalized with respect to the initial control pulse and plotted against the cumulative recovery time spent at −80 mV. Both data sets were well fit by a mono-exponential function and had similar time constants. Data shown are from the same cell and are representative of 3 experiments. D, Data from another cell (representative of 4 such experiments), where the experiment protocol was analogous to that of panels A and B and was performed at pH 7.4 with 2 mM Ni 2+ . As with low pH, the time courses of recovery from enhanced slow inactivation and resting inactivation were similar. 108 3.4 DISCUSSION The two primary questions addressed in this chapter are, first, how pH and Ni 2+  influence slow inactivation of Kv1.5 and, second, whether the H +  and Ni 2+  induced loss of Kv1.5 channel availability represents slow inactivation occurring at rest. We focus first on the effects of H +  and Ni 2+  at +50 mV before discussing the evidence for, and the ramifications of resting inactivation. Although the enhanced current decay observed with either ligand at +50 mV could in theory arise from fast open channel block, this explanation is ruled out by previous single channel recordings that revealed no change of the single channel conductance with either ligand (Kwan et al., 2004; Kwan et al., 2006). Our results point instead to an enhancement of a complex multistate slow inactivation process as the basis for the accelerated and more complete current decay observed at +50 mV in low pH and Ni 2+  (Figs. 3. 2 - 3.4). As has been reported for ShakerIR channels (Olcese et al., 1997; Klemic et al., 2001; González-Pérez et al., 2008), the time course of the recovery of Kv1.5 channels from control slow inactivation (pH 7.4, 0 mM Ni 2+ ) indicates that there are at least two kinetically distinct slow inactivated states (Fig. 3.6; Rich & Snyders, 1998). Low pH biases slow inactivation to a state from which recovery is fast (Fig. 3.6B; Table 3.1), while Ni 2+  even more strongly biases slow inactivation to a state from which recovery is slow (Fig. 3.6C). While we cannot rule out an alternative explanation that the slow time course of recovery from Ni 2+ -enhanced slow inactivation reflects the time dependence of Ni 2+  unbinding, the similarity of the slower recovery time constant following control and Ni 2+ - enhanced slow inactivation suggests this is unlikely. A seemingly similar biasing of slow inactivation has been reported in Kv1.3, where increasing K +  occupancy of the outer pore by raising [K + ]o increases the amplitude of a fast component of recovery (Levy & Deutsch, 1996a). However, this explanation does not apply in our experiments, since the effects of H +  and Ni 2+  on the recovery time course were observed in 0 mM K + o. 109 In Kv1.5 the H +  or Ni 2+  binding sites are most likely the H463 residues located in the pore turrets (Kehl et al., 2002; Kwan et al., 2004; Eduljee et al., 2007). An important finding in this connection is that substitutions at these positions (e.g., H463G, H463R) can dramatically accelerate slow inactivation (Kehl et al., 2002; Eduljee et al., 2007). In that light, our observation that ligand binding at this site can enhance slow inactivation in wt Kv1.5 bears striking parallels to the action of Cd 2+  to enhance slow inactivation in ShakerIR when the threonine at position 449, a site that also influences slow inactivation kinetics, is mutated to cysteine (Yellen et al., 1994). Like Ni 2+ -enhanced slow inactivation in Kv1.5, in ShakerIR T449C, “ „Cd2+-induced inactivation‟ [at 0 mV] places the channel entirely in the more slowly recovering” of two inactivated states (Yellen et al., 1994). As noted by Yellen et al. (1994), the ligand-induced enhancement of inactivation during a depolarizing pulse cannot be explained solely by an action to make inactivation an absorbing state; it must also involve an increase of the rate constant for the open to inactivated transition. This is illustrated in Fig. 3.9B which shows the outcome of a numerical simulation of a gating scheme incorporating, as required by the experimental data, two open-inactivated states (Fig. 3.9A, see figure legend for details) in which the O → I transitions are unchanged and inactivation is made to be absorbing by setting the rate for one or both of the I → O transitions to zero. In this scenario, inactivation is complete with sufficiently long pulses but the rate of inactivation is limited by the O → I rate constants and cannot achieve the rates observed experimentally. Much faster slow inactivation is achieved when the rate constant for either one of the O → I transitions is increased (Figs. 3.9C, D). The simulation results support the conclusion that ligand binding at H463 in Kv1.5, like Cd 2+  binding at T449C in ShakerIR, increases the rate of the O → I transition. Whether ligand binding does so by causing an allosteric effect, by acting electrostatically to influence the K +  concentration at or near the outer pore mouth, or by some other mechanism, remains an open question. 110  Figure 3.9. Theoretical outcomes of the modulation of slow inactivation by ligand binding. A, The gating model has several closed states in the activation pathway and the open state (O) is coupled to two slow inactivation states, If and Is. For simplicity, the ligand binding/unbinding steps of SCHEME 3.2 (Fig. 3.1) are omitted from this revised gating scheme. Although the inactivation rate constants (O→If, If→O, O→Is and Is→O of 0.5, 2.2, 0.2 and 0.1 s -1 ) have been chosen to give macroscopic current behaviour similar to that in Kv1.5 during a 5 s pulse to +50 mV, the simulation is provided for the purpose of illustration only. Panel B shows the outcome of the model assuming that ligand application, represented by the bar above the superimposed traces, makes one or both of the inactivation states absorbing (I→O = 0 s-1) but affects neither of the O→I transitions. The relatively modest effect of making Is (dotted trace), If (short dashes trace) or both If and Is (long dashes trace) absorbing emphasizes that rapid current decay, such as that observed in Kv1.5 with H + , must involve an increase of the rate constant for the O→I transition. C, the selective enhancement of the O→If transition during a 2 s ligand application produces a rapid, mono-exponential decay of current that is followed by a rapid recovery that “overshoots” the control trace. D, ligand application that selectively enhances the O→Is transition is followed by a slowly recovering current. E, In the model, which does not incorporate inactivation from the resting state, an increase of the O→If rate constant and a decrease of the If→O rate constant, which mimics the time course and extent of the current decay in pH 5.4 solution (Fig. 3.3), causes only a 12% decrease of peak current compared to the control response. This outcome supports the conclusion that the enhancement of slow inactivation makes, at best, a minor contribution to the reduction of peak current observed with Ni 2+  or H + (e.g., Fig. 3.2). The calibration bar represents 1 s for panels B-D and 0.1 s for panel E. 111 3.4.1 Multiple inactivated states can explain the recovery overshoot during a depolarizing pulse A frequent finding with a transient low pH pulse applied during a depolarizing pulse was a recovery current that overshot the superimposed control current trace (Fig. 3.3A). The recovery overshoot can be explained given the existence of at least two inactivation processes at +50 mV (Fig. 3.6). This is illustrated in the numerical simulation of Fig. 3.9C where an overshoot of the recovery current is obtained when the ligand (i.e. H + ) causes a selective enhancement of the O → I transition for the inactivated state from which recovery is fast (If; Fig. 3.9A) and which is not directly connected to a more stable inactivated state (Is). With this constraint, during the ligand application If is populated at the expense of Is and once the ligand is removed there is rapid recovery from If, resulting in the recovery overshoot. When the ligand selectively accelerates inactivation to Is, the recovery kinetics are slow and current does not overshoot the control current level (Fig. 3.9D), as was usually the case following Ni 2+  application. 3.4.2 Low pH and Ni2+ also induce resting inactivation of Kv1.5 channels To account for the concentration dependent decline of the peak test current and maximal conductance (Fig. 3.2) we have proposed that low pH or Ni 2+  must also trigger a resting inactivation process because enhanced slow inactivation, except at pHs much lower than those used in these experiments, is predicted to have a relatively small effect on peak test currents. To illustrate this point, Fig. 3.9E shows the outcome of a numerical simulation of the gating scheme in Fig. 3.9A where the only effect of the ligand is to enhance slow inactivation from the open state by approximately 14-fold, in this case by increasing the rate of the O → If transition and decreasing the rate of the If → O transition. Note that despite an increase of slow inactivation at +50 mV to a rate 2- to 3-fold faster than that observed at pH 5.4, the lowest pH we examined, the effect on the peak current amplitude is much smaller than observed in Figs. 3.2A, B. 112 3.4.3 Modelling the enhancement of resting inactivation by H+ and Ni2+ Experimentally, the onset of ligand-induced resting inactivation followed a single exponential time course that was generally slower than enhanced slow inactivation over the range of [H + ] and [Ni 2+ ] tested (Figs. 3.3F and 3.4E). Figure 3.10 shows the outcome of a numerical simulation of resting inactivation with Ni 2+  (panels A-C) and with low pH (panels D and E) using a four-state model (Figs. 3.1A, 3.10F). The critical feature of the model is that resting inactivation is faster (k12 > k03) and the inactivated state is more stable (k21 < k30) for the liganded states. In the fitting procedure, rates were assigned for the forward and reverse transitions between the A and U states and the rates for the A-L  U-L transitions were adjusted to match as closely as possible, by eye, the experimental steady-state availability relationship (solid lines of Fig. 3.10B, D) and the time-dependence of the availability change (solid lines of Fig. 3.10C, E) over a range of ligand concentrations. In this iterative approach the values for the rate constants for ligand binding (k01, k32) and unbinding (k10, k23) were also adjusted to optimize the fit to the experimental data. We are unsure if the rate constants in the simulation represent unique solutions and, given the absence of direct information about the binding and unbinding kinetics, this analysis is offered only as a preliminary assessment of the model‟s plausibility. 113        Figure 3.10. Numerical simulation of resting inactivation caused by Ni 2+  or low pH. 114 Figure 3.10. Numerical simulation of resting inactivation caused by Ni 2+  or low pH. For the simulation with Ni 2+ , the rate constants, based on the scheme in panel F, were: k01 = 110 mM -1 s -1 ; k10 = 150 s -1 ; k12 = 1.422 s -1 ; k21 = 0.036 s -1 ; k23 = 2.19 s -1 ; k32 = 1100 mM -1 s -1 ; k03 = 0.008 s -1 ; k30 = 0.14 s -1 . A, the time dependence of the Ni 2+ -induced decrease of availability, measured as the sum of the proportions of channels in states A and A-Ni 2+ , was well-fitted by a single exponential. The [Ni 2+ ] for the top sweep was 2.5 M and was doubled for each subsequent sweep. B, the steady-state availability (), measured from the traces of panel A, is plotted against the [Ni 2+ ]. The dashed curve is the fit of the simulated data to the Hill equation (Kd = 36 M; nH = 1.0) and the solid curve is the fit to the experimental data (Kd = 33 M; nH = 1.15). C, the values for τRI for the simulated (, from fits of selected traces in A) and experimental () data are plotted against the [Ni 2+ ]. For the simulation of extracellular pH induced resting inactivation (D, E), protonation was assumed to trigger an inactivation process different from that induced by Ni 2+  binding. For () in D and E, the rate constants were: k01 = 2 x 10 9  M −1 s −1 ; k10 = 9000 s −1 ; k12 = 4.356 s −1 ; k21 =0.176 s −1 ; k23 = 182.3 s −1 ; k32 = 2 x 10 10 M −1 s −1 ; k03 = 0.03 s −1 ; k30 = 0.6 s -1  and the binding was assumed to involve a single site (nH = 1). For () in D and E, the steepness of the experimental availability curve (nH = 1.5) was mimicked by changing k01 and k32 to 5 x 10 12  M −1.5 s −1  and 5 x 10 13  M −1.5 s −1 and  multiplying k01 and k32 by [H + ] 1.5 . The control pH was 7.4 and the test pH ranged from 7.2 to 5.4 in 0.2 pH unit steps. The best fits of the simulated availability data for () and () to the Hill equation are indicated by the dotted line (Kd = 1.8 x10 −7  M; pKa = 6.73, nH = 1.0) and the dashed line (Kd = 1.8 x 10 −7  M, pKa = 6.73, nH = 1.5), respectively. The solid line is the fit of the experimental concentration dependence (Kd 1.6 x 10 −7  M, pKa = 6.8, nH = 1.5) taken from Kehl et al. (2002). In panel E the time constants for the loss of availability for the two simulations described for panel D is compared to the experimental values taken from Fig. 3.3F and represented by . F, the state diagram for resting inactivation at −80 mV. See Figure 3.1A for explanation. 115 The simulation of Ni 2+ -induced resting inactivation shows first, as observed experimentally, a mono-exponential time course for the decrease in channel resting availability, expressed as the sum of the proportion of channels in states A and A-Ni 2+  (Fig. 3.10A), and, second, that the fitted time constants correlate well with the mean experimental values ( and , respectively; Fig. 3.10C). There was also good agreement between the simulated ( and dashed line) and the experimental (solid line) steady-state availability (Fig. 3.10B). Simulated recovery (not shown) in 0 mM Ni 2+  was mono-exponential and the fitted RI of 6.8 s was similar to the experimental value of 7.1 ± 0.4 s. A similar approach was used to simulate the effects of extracellular acidification (Figs. 3.10D, E), however the model explicitly assumed the involvement of a resting inactivation process that differs from Ni 2+ -induced resting inactivation both in its maximum rate (k12) and in the rate of recovery from inactivation (k21). A reasonable fit to the experimental data was obtained when, as with the Ni 2+  simulation, there was a single binding site (; Fig. 3.10D, E). Both the fit of the steepness of the steady-state availability relationship and the fit of the time dependence of inactivation were improved when, as implied from the value of 1.5 for the Hill coefficient fitted to the experimental data (Kehl et al., 2002), highly cooperative binding was assumed (; Fig. 3.10D, E). The time constant for the simulation of recovery at pH 7.4 following a pulse to pH 5.4 was 3.1 s or roughly two-fold slower than the experimentally- measured value of 1.6 s (Table 1). This deviation between the simulated and experimental recovery data may reflect a contribution from protonation of residues elsewhere on the channel. In that connection we have noted previously that the shift of the activation curve is much greater for H +  than for Ni 2+  (Kehl et al., 2002; Kwan et al., 2004). 116 3.4.4 Are the effects of H+ or Ni2+ on open and resting Kv1.5 channels related? It seems well-established that both ligands cause resting inactivation and enhance slow inactivation. But is there a mechanistic connection between these two inactivation processes? We consider this possibility first for low pH. The similarity of the time constants for recovery at −80 mV from H+-induced resting inactivation and control or H+-enhanced slow inactivation (e.g., Fig. 3.6B) is an argument that H + -induced resting inactivation involves the same states visited in H + -enhanced slow inactivation and control slow inactivation. An additional argument for a conformational similarity of the enhanced slow inactivated state and the resting inactivated state is the similarities of their recovery kinetics during a depolarizing pulse (Fig. 3.7A). In other words, at least based on kinetic criteria, H + -induced resting inactivation appears to involve a process akin, if not identical, to slow inactivation. The evidence for conformational similarity between Ni 2+ -induced resting inactivation and enhanced slow inactivation  is less compelling. The finding that recovery from Ni 2+ -enhanced slow inactivation proceeds virtually exclusively with the same slow recovery time course that roughly 50% of the inactivated channels follow after control slow inactivation (Fig. 3.6C) is good evidence that Ni 2+  steers inactivation at +50 mV to the more stable of the two inactivated states. Additionally, recovery at +50 mV following either enhanced slow inactivation or resting inactivation is comparable (Fig. 3.7B), which again implies the same inactivated state is involved in inactivation at −80 mV and +50 mV. Evidence against a mechanistic overlap of resting inactivation and enhanced slow inactivation is that recovery at −80 mV (in 0 Ni2+) from Ni2+- induced resting inactivation was roughly two-fold faster than from Ni 2+ -enhanced slow inactivation (Fig. 3.6C). Both the onset (i.e. τRI) and extent of resting inactivation induced by either ligand were affected by increasing [K + ]o from 0 to 3.5 mM (Figs. 3.2 – 3.4). Exactly why the addition of K + o 117 causes the onset of resting inactivation to become bi-exponential is unclear but we suspect that it is related to complexities of the non-competitive interaction between H + o and K + o (Kehl et al., 2002). The sensitivity of resting inactivation to [K + ]o (Kd = 1 mM; Kehl et al., 2002) contrasts with the comparative [K + ]o insensitivity of slow inactivation of Kv1.5 at +50 mV and is likely due to the absence of the mitigating influence of outward K +  current. This [K + ]o sensitivity might reflect an action of K +  to impede the closing of an inactivation gate in the outer aspect of the selectivity filter. Although the [K + ]o sensitivity of resting inactivation is reminiscent of the [K + ]o sensitivity of slow inactivation in other Kv channels (Baukrowitz & Yellen, 1995; Levy & Deutsch, 1996a; Kiss & Korn, 1998) and offers some support for slow inactivation occurring at rest, it is not an infallible benchmark since even with regard to slow inactivation it is a criterion that is not uniformly satisfied (Yang et al., 1997; Fedida et al., 1999). Nonetheless, support for the possibility that resting inactivation does involve a conformational change of the outer pore mouth comes with our demonstration that, as reported for slow inactivated ShakerIR channels (Basso et al., 1998; Harris et al., 1998), Ba 2+  movement, at −80 mV, to and from a deep binding site in the Kv1.5 pore is prevented at low pH (Fig. 2.7) and by Ni 2+  (Fig. 3.5). 3.4.5 Evidence from other Kv channels for resting inactivation As noted above, there are intriguing similarities between the effects of Ni 2+  on the inactivation of Kv1.5 and the effects of Cd 2+  on the inactivation of ShakerIR T449C channels (Yellen et al., 1994). In both cases, divalent cation binding accelerates slow inactivation and induces resting inactivation. The latter phenomenon was not studied in detail in ShakerIR T449C but it appears, as with Ni 2+  in Kv1.5, that the onset of resting inactivation is much slower than slow inactivation during a strong depolarization. In wild-type Kv1.4 channels (Pardo et al., 1992) and the fast-inactivating T449A, -E and -K ShakerIR mutant channels (López-Barneo et al., 1993) removing K + o causes a loss of current, 118 which in both cases was attributed to a decrease of channel availability at rest. Not only does the same phenomenon occur in Kv1.5 at low pH or in Ni 2+ , as well as in Kv1.5 H463G at pH 7.4 (Kehl et al., 2002), but Fig. 3.8 illustrates that the time course of recovery from the loss of current induced by either removing K + o or by a depolarization is also the same. 3.4.6  Concluding remarks As observed by Lopez-Barneo et al. (1993) in fast-inactivating ShakerIR mutants, in Kv1.5 the enhancement by low pH or Ni 2+  of slow inactivation (depolarization-induced inactivation) is paralleled by the induction of resting (closed) state inactivation. The data also suggest fairly strongly for low pH and less so for Ni 2+ , that resting inactivation, enhanced slow inactivation and slow inactivation can involve similar conformational states. Further study is required, but if resting inactivation truly is a variant of slow inactivation it indicates the potential for variability in the degree of coupling between activation and slow inactivation. Complete uncoupling of the type implied here for Kv1.5 appears also to occur in the pore helix mutant ShakerIR W434F, which has been described as being permanently inactivated at rest (Yang et al., 1997). Resting inactivation in ShakerIR FWFW channels (i.e. with two W434 residues and two W434F residues; Yang et al., 1997) also shares the [K + ]o sensitivity reported here for Kv1.5. As shown for Kv1.5, in the rapidly inactivating FWFW ShakerIR construct the time course of recovery from the resting inactivated, or unavailable, state is similar to that for recovery from depolarization-induced inactivation (Yang et al., 1997). This raises the possibility that the complex, multistate slow inactivation process, which is normally triggered by a tight coupling between outward voltage sensor movement and a subsequent interaction with the pore domain (Loots & Isacoff, 2000), can proceed independently of activation following mutations in the pore helix (ShakerIR W434F) or in Kv1.5 by ligand binding at H463 in the pore turret. 119 4 An examination of the pH or K+o-dependent loss of current through wt ShakerIR or fast-inactivating ShakerIR and Kv1.5 mutant channels 4.1 INTRODUCTION The studies described in Chapters 2 and 3 have shown that, in addition to enhancing depolarization-induced slow inactivation, low pH causes a reduction of the availability of Kv1.5 channels at −80 mV that is likely related to resting inactivation. Low pH has also been shown to enhance slow inactivation of wt ShakerIR at depolarized potentials (Pérez-Cornejo, 1999; Starkus et al., 2003), as well as to induce a decrease of maximal conductance that, based on voltage-clamp fluorimetry measurements, has been suggested to be due to slow inactivation occurring at resting potentials (Claydon et al., 2007). In support of this, single channel recordings showing multiple null sweeps indicate that low pH causes a reduction in the availability of ShakerIR channels to conduct upon depolarization (Claydon et al., 2007). A reduction in peak macroscopic current (at pH 7.4) has also been described for fast-inactivating ShakerIR T449 mutants (-K, -A, and -E) and the Kv1.5 H463G mutant with decreases in [K + ]o and has been proposed to be due to inactivation from the closed state at −80 mV (López-Barneo et al., 1993; Kehl et al., 2002; Zhang et al., 2005). However, the time courses for the onset of and recovery from the putative H + - or [K + ]o-dependent resting inactivation of wt ShakerIR and the fast-inactivating ShakerIR and Kv1.5 mutants has not been assessed. As shown in Chapter 3, a comparison of the kinetics for resting inactivation and slow inactivation can provide information on whether the two processes are mechanistically related. Thus, to examine the possibility that the resting inactivation in either wt ShakerIR or the ShakerIR T449 and Kv1.5 H463G mutants is mechanistically similar to slow inactivation, we have characterized the time- dependence of the onset and offset of the loss of current induced by low pH or low [K + ]o (at pH 120 7.4), either during a depolarization (+50 mV) or at rest (−80 mV). The data show that low pH accelerates the onset of slow inactivation in ShakerIR and that this H + -enhanced slow inactivation accounts for a greater proportion of the decrease in peak current than observed with Kv1.5. It was also found that, in the ShakerIR T449K, T449A and Kv1.5 H463G mutants, the time course of recovery from 0 K + o-induced resting inactivation (at pH 7.4) was the same as that for recovery from slow inactivation. These results support the hypothesis that the loss of wt and mutant ShakerIR or Kv1.5 channel availability induced by low pH or low [K + ]o is a form of slow inactivation occurring at the resting potential. 121 4.2 METHODS 4.2.1 Site-directed mutagenesis and cell preparation We used as the wild-type construct the ShakerIR channel, which has a deletion (6-46) in the amino terminus that completely removes N-type inactivation (Hoshi et al., 1990). Point mutations of wt ShakerIR in the pcDNA3 expression vector were made using the QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA, USA) to convert the threonine at position 449 to a Lys, Ala, or Glu residue. The Kv1.5 H463G mutant was made using similar procedures with the wt Kv1.5 construct in the pcDNA3 vector. Ionic currents were recorded from a tsA201 cell line, which is a transformed HEK-293 cell line that stably expresses an SV40 temperature-sensitive antigen and is able to produce high levels of recombinant proteins. Cells were maintained at 37°C in an atmosphere of 5% CO2 in air and in MEM supplemented with 10% foetal bovine serum, 1% (v/v) penicillin-streptomycin, and 0.5 mg ml −1  geneticin. Two days before an experiment, the cells were washed with MEM, dissociated for passage using trypsin/EDTA and plated on round 25 mm 2  coverslips. Twenty- four hours later, the cells were transiently transfected with 2 µg of either wt or mutant ShakerIR plasmid cDNA using 3 µl of Lipofectamine 2000 (Invitrogen). The cells were also co-transfected with 1 µg of CD8-pcDNA3. Current recordings were performed 24-36 hours after transfection. Before experiments, transfected cells were incubated for ≥30 min with Dynabeads®CD8 (Invitrogen) coated with anti-CD8 monoclonal antibody (1.2 x 109 beads ml−1 in MEM). All tissue culture supplies were obtained from Invitrogen (Burlington, Ontario, Canada). 4.2.2 Recording solutions The standard 3.5 mM K +  bathing solution contained (in mM): 140 NaCl, 3.5 KCl, 2 CaCl2, 1 MgCl2, 5 glucose, 10 HEPES and was titrated at room temperature to pH 7.4 with 122 NaOH. Bath solutions with higher [K + ] were prepared by substituting KCl for NaCl. Solutions with lower [K + ] were prepared by replacing KCl with NaCl. Low pH solutions had the same ionic composition as the standard solution, except for the replacement of HEPES by MES. Due to the endogenous expression of H + -activated outwardly-rectifying chloride channels in HEK- 293 cells (Zhu et al., 1998; Lambert & Oberwinkler, 2005), the anion channel blocker 4,4′- diisothiocyanostilbene-2,2′-disulphonic acid (DIDS) was routinely added (30 - 60 µM) to bath solutions in experiments where a pH ≤5 was used. The addition of DIDS did not have an effect on the properties of K +  currents measured at either pH 7.4 or low pH (data not shown). The standard patch pipette solution was the same as described in Section 2.2.2. Chemicals were obtained from the Sigma Aldrich Chemical Co. (Mississauga, Ontario, Canada). 4.2.3 Fast solution exchange For experiments requiring rapid solution exchange, a Warner Instruments SF-77B Perfusion Fast Step system was used as described in Section 2.2.3. In some experiments, single-walled, triple- barrelled square glass tubing (0.6 mm ID barrels; Warner Instruments) was used instead of theta glass tubing (see Appendix). In these situations, the triple-barrelled tubing was pulled in one step to an inner barrel diameter ≤0.25 mm and then cut into two parts using a ceramic wafer before connection to the solution reservoirs as described previously. Rapid movements of 300 – 600 µm were made to switch between the three barrel positions. The time course of solution exchange, estimated as detailed in Section 2.2.3, had both a latency and time constant of ≈25 ms using this system, regardless of the relative positions of the starting and final barrel. 4.2.4 Definitions of terms The effects of external H +  or K +  ions on wt or mutant ShakerIR and Kv1.5 channels, respectively, are described with respect to their actions at either −80 mV or +50 mV. As such, 123 the term resting inactivation, as defined in Section 3.2.4, is used here to describe the loss of channel availability and, consequently, peak macroscopic current, occurring at −80 mV and induced by low pH or removal of K + o. Similarly, slow inactivation, sometimes called control slow inactivation, refers in this chapter to the current decay observed at +50 mV in control pH 7.4, 3.5 mM K +  solution. For consistency, the accelerated current decay observed at low pH or in 0 mM K + o is referred to as enhanced slow inactivation. Thus, the simplified gating schemes of Fig. 3.1 may also be used here to describe the putative actions of low pH to induce resting inactivation and enhance slow inactivation of wt ShakerIR channels.  Similar schemes may be used to describe the loss of current observed with the fast-inactivating ShakerIR and Kv1.5 mutants upon removing K + o. However, to account for the observation that removal of K +  enhances inactivation, in this case the equilibrium between the K + -bound states (i.e. A-K +   U-K+ in SCHEME 3.1 and O-K+  OI-K+ in SCHEME 3.2) represents the control situation that is biased towards the available and open states, while the K + -unbound inactivated states (i.e. U and OI) represent the resting inactivated and enhanced slow inactivated states populated upon removal of K + o. 4.2.5 Signal recording and data analysis In an experiment, a section of glass coverslip to which cells had attached was placed in the recording chamber. The cells were washed with standard bath solution for 2 min, after which those still bound by Dynabeads ® CD8 were selected for recording. Whole-cell currents were recorded at room temperature (20 - 25°C) as described for Chapters 2 and 3. Where appropriate, the pH or [K + ]o dependence of the peak current amplitude or time course of inactivation was quantified by fitting the data to a generalized form of the Hill equation: 124               (Equation 4.1) where y is the peak current amplitude or time constant; ymax is the maximal value of y; Kd is the apparent equilibrium dissociation constant; [L] is the [H + ] or [K + ]; and nH is the Hill coefficient. For the effects of H + , the value of Kd was converted to a pKa by taking the negative logarithm of the Kd  (which is equivalent to the acid dissociation constant, Ka). Data are presented as mean ± S.E.M.; n represents the number of cells tested. Statistical comparisons were performed using a one-way ANOVA and Tukey test; significance was defined as P < 0.05. 125 4.3 RESULTS 4.3.1 Low external pH enhances slow inactivation and reduces peak current of ShakerIR Figure 4.1A shows a macroscopic current trace (black trace) recorded at pH 7.4 in 3.5 mM K + o from a tsA201 cell expressing wt ShakerIR channels. Also shown is a superimposed current trace recorded at the steady state following a switch to pH 5.0 solution (grey trace). As with Kv1.5, it is clear that slow inactivation is much faster at the lower pH; the time constants for slow inactivation (τinact) at pH 7.4 and pH 5.0 were compared by fitting the respective current decays to a single exponential function (although at pH 7.4 a double exponential may be more appropriate (Meyer & Heinemann, 1997), see Section 3.3.1). The pH-dependence of the τinact measured in similar experiments is summarized in Fig. 4.1B. A fit of the [H + ]-dependence of τinact to the Hill equation gives a pKa value (5.69 ± 0.01; nH = 1.26) that is comparable to that reported by others for ShakerIR channels expressed in Xenopus oocytes (Starkus et al., 1997; Pérez-Cornejo, 1999). 126    Figure 4.1. Decreases in external pH accelerate slow inactivation of ShakerIR and reduce peak current amplitude. A, Superimposed currents recorded from the same cell in 3.5 mM K + o using 10 s depolarizations from −80 mV to +50 mV. The black trace was recorded under control, pH 7.4 conditions. The grey trace was recorded at the steady-state following a switch from pH 7.4 to pH 5.0 solution. Dashed lines represent fits of the current decay to a mono-exponential function with a time constant τinact. B, The time constants measured in experiments such as that in A are plotted against pH and fit to the Hill Equation. C, Peak current amplitudes measured in experiments such as that described in A were normalized to their respective pH 7.4 control and plotted against pH. The solid line represents a fit of the data to the Hill Equation. All data points represent the mean ± S.E.M. from at least 8 cells. 127 The traces of Fig. 4.1A also show that the peak current amplitude is reduced at pH 5.0. This effect is likely distinct from the charge screening effect of the increased [H + ]o, as the test pulse voltage of +50 mV is near the top of the activation-voltage curve, even at very low pH (Starkus et al., 2003; Claydon et al., 2007). The pH-dependence of the peak current amplitude (normalized to that at pH 7.4) is shown in Fig. 4.1C and has a pKa of 4.81 ± 0.30 (nH = 1.00). Together, the results of Fig. 4.1 confirm that extracellular acidification causes: i) an enhancement of both the rate and extent of the current decay at +50 mV; and, ii) a decrease in the peak macroscopic current, as previously reported by others (Pérez-Cornejo, 1999; Starkus et al., 2003; Claydon et al., 2007). In comparison to the pH-dependence of τinact, the lower pKa value of the peak current amplitude suggests that the accelerated slow inactivation may not account for all of the H + -induced decrease in current at lower pH values and, thus, that an additional process occurring at rest may be involved. However, it is difficult from these results to determine the relative contributions of slow inactivation and resting inactivation to the loss of peak current. As such, the next experiments were performed, where the pH of the bathing solution was rapidly changed at either +50 mV or −80 mV to assess the effects of low pH on either slow inactivation or the resting channel availability. 4.3.2 Slow inactivation is enhanced during a depolarization by a transient decrease in pH The superimposed current traces of Figure 4.2 show the effects of rapidly and transiently decreasing pH during a depolarizing pulse. As described in Section 3.3.2 for Kv1.5 and low pH, this protocol avoids the potential effects of H +  on resting channels and allows us to look predominantly at the effect of low pH on ShakerIR slow inactivation from the open state. Similar to our findings with Kv1.5 (see Fig. 3.3A) and those of Starkus et al. (2003) with ShakerIR in Xenopus oocytes, decreasing pH during a depolarizing pulse enhances both the rate and extent of the slow inactivation of open ShakerIR channels. The time constant for slow inactivation 128 measured with transient decreases in pH tended to be larger than those measured at the steady- state (383 ± 27 ms vs. 226 ± 35 ms at pH 5.0, respectively; see Fig. 4.1B). The slower time course may be partly due to the limiting time course for the fast solution change (see Section 4.2.3), such that the pH following the initial switch is greater than 5.0, and resulting in an underestimate of the effect of H +  on τinact. Alternatively, the slightly faster current decay observed at the steady-state (Fig. 4.1) may be due to additional effects of H +  on resting channels that accelerate slow inactivation upon depolarization.   Figure 4.2. Current recovers from a transient decrease in pH applied during a depolarizing pulse. Superimposed current traces recorded from the same cell using 10 s pulses from −80 mV to +50 mV. The black control trace was recorded entirely under 3.5 mM K + o, pH 7.4 conditions. For the grey trace, the pH of the bathing solution was rapidly switched to pH 5.0 for 1 s, starting 500 ms after the beginning of the test pulse. The bars at the top of the figure represent the perfusion protocol. Note that there is a short lag period before the enhancement of slow inactivation at pH 5.0 is observed (see Section 4.2.3). 129 Interestingly, the switch from low pH back to pH 7.4 during a depolarizing pulse was associated with recovery of the ShakerIR current to the same level as that of the inactivating control current at pH 7.4. In that sense, the results of Fig. 4.2 with ShakerIR and low pH are reminiscent of those with Kv1.5 and low pH (see Fig. 3.3A) with the exception that the recovery of ShakerIR current during a depolarizing pulse is relatively slow and does not overshoot the control current. This behavior of ShakerIR channels in the mammalian tsA201 cell line is in apparent contrast to findings with ShakerIR expressed in Xenopus oocytes, which showed no recovery following a transient low pH exposure, although that may have been due to the short duration of the recovery current recording (Starkus et al., 2003). The recovery is unlikely to be due to the reversal of open channel block by H + , as single channel recordings of both Kv1.5 and ShakerIR channels have precluded rapid open channel block as the process underlying the enhanced current decay at low pH (Kwan et al., 2006; Claydon et al., 2007). Open channel block of ShakerIR was also excluded based on the lack of effect of external H +  on instantaneous tail current amplitudes (Starkus et al., 2003). A simple explanation may be that the current recovery reflects de-protonation and a re-establishment of the equilibrium between open and slow inactivated channels at pH 7.4 (see SCHEME 3.2; Fig. 3.1). 4.3.3 The time course of H+-induced resting inactivation It has previously been suggested that the enhanced rate of slow inactivation is the main cause of the reduction of ShakerIR peak current at low pH (Starkus et al., 2003). However, the recent voltage-clamp fluorimetry studies of Claydon et al. (2007) showed that, at rest at low pH, the outer pore of ShakerIR undergoes conformational changes associated with slow inactivation, suggestive of a H + -induced resting inactivation process. Additionally, numerical simulations (not shown) like those described in Fig. 3.9E, whereby the rate of the O → I transition is increased so that the simulated time constant for slow inactivation matches that observed experimentally at 130 low pH, result in a decrease in peak current amplitude that is at least 50% smaller than that observed experimentally. Thus, it is unlikely that H + -enhanced slow inactivation accounts for all of the loss of peak current at low pH. In the following experiment we tested the hypothesis that the low pH-induced decline of peak current is due in part to a process occurring at the resting potential. As with Kv1.5 in Chapter 3, the following experiments were performed on the premise that an analysis of onset and/or recovery kinetics might shed light on any mechanistic relationship between slow inactivation at +50 mV and resting inactivation at −80 mV. Figure 4.3A shows superimposed current traces recorded in multiple sweeps with test pulses to +50 mV applied after variable durations of exposure to pH 4.5 solution at −80 mV. As the length of time at pH 4.5 increased, the peak test current amplitude declined in a mono-exponential manner, with a time constant defined as τRI. The contribution of cumulative slow inactivation to the total current loss observed in the last sweep is precluded by the 60 s intersweep interval, during which channels fully recovered (not shown). Based on the numerical simulations described above, the very fast H + -enhanced slow inactivation at +50 mV during the test pulses likely accounts for at least some of the reduction of peak current observed in each sweep. The measurement of the onset of resting inactivation is also confounded by the frequently observed ≈2-fold decrease in the value of τinact between the first and last test currents in low pH solution, which we assume is due to the limiting kinetics of the solution (and pH) change during the short exposure times of the first few sweeps and the very fast and highly pH-dependent time course of slow inactivation. Thus, although this experimental protocol was successfully used to discern the extent and time course of H + -induced resting inactivation of Kv1.5, the results with ShakerIR are not as definitive. Nevertheless, the time constants (τRI) for the loss of ShakerIR peak current amplitude over a range of pHs are summarized in Fig. 4.3B and were, in general, greater than 131 τinact, except at pH 5.5. Additionally, the pH dependence of τRI appears to be left-shifted compared to τinact. This is consistent with the findings in Fig. 4.1B and C, which showed that the peak current amplitude had a lower pKa than the time constant of slow inactivation. Interestingly the loss of current at pH values as low as 4 (τRI ≈ 200 ms) occurs at least an order of magnitude faster than the previously reported pH-dependent changes in fluorescence emission from a fluorescent tag at S424C in the Shaker outer pore (Claydon et al., 2007). Given the limitations of the present data and the fact that the fluorescence changes do not necessarily directly report on the conducting state of the channel, it is difficult to conclusively say how accurately either measurement reflects on the time course of H + -induced resting inactivation.  132   Figure 4.3. The time course of resting inactivation is time- and pH-dependent. A, Superimposed ShakerIR current traces from the same cell showing the time course of the loss of peak current amplitude induced at pH 4.5 in 3.5 mM K + o. For each trace, a single 800 ms test pulse from −80 mV to +50 mV was applied at a known interval following the switch from pH 7.4 to pH 4.5 solution; the interval was increased for each successive sweep. Also shown is a control current trace recorded at pH 7.4. The time constant for the onset of resting inactivation (τRI) was derived by fitting the peak current amplitudes of the test pulses to a single exponential function. B, τRI values measured in experiments like that displayed in A are plotted against pH. The time constants for enhanced slow inactivation at low pH (from Fig. 4.1) are also shown for comparison. The plot shows that the pH dependence of resting inactivation is right-shifted to that of τinact, consistent with the results shown in Fig. 4.1C. 133 4.3.4 Recovery from H+-enhanced slow inactivation is fast To further examine the nature of the H + -enhanced slow inactivated state, the time course for recovery from enhanced slow inactivation at +50 mV was compared to that for recovery from slow inactivation under control, pH 7.4 conditions. Given that our ability to separate the H + - induced resting inactivated state from the H + -enhanced slow inactivated state was limited by the speed of the solution change, the recovery kinetics from resting inactivation were not assessed. Pair-wise comparisons were made with measurements from the same cell to control for any variability in the rate and extent of control slow inactivation. Figure 4.4 displays single sweeps from a representative experiment monitoring recovery following: A, control slow inactivation induced by 10 s at +50 mV at pH 7.4 and, B, enhanced slow inactivation induced by ≈5 s at pH 4 at +50 mV. Recovery from each condition in standard 3.5 mM K + , pH 7.4 solution was monitored with a train of test pulses to +50 mV.  134     Figure 4.4. The recovery of ShakerIR channels from H + -enhanced slow inactivation is fast. A, Current trace recorded in a single sweep showing the recovery following a 10 s conditioning pulse from −80 mV to +50 mV to induce slow inactivation in 3.5 mM K+o at pH 7.4. The recovery was monitored with a train of 20 ms test pulses to +50 mV. B, Following a brief control pulse at pH 7.4, enhanced slow inactivation was induced by rapidly switching from pH 7.4 to pH 4.0 solution 50 ms after the start of a 5 s conditioning pulse to +50 mV. The solution was switched back to pH 7.4 at the end of the conditioning pulse, after which recovery was monitored as in A. C, The peak test current amplitudes from panels A and B were normalized to the control current at the beginning of each sweep and plotted against the cumulative recovery time at −80 mV. The solid and dashed lines represent, respectively, the results of a global fit of the control and enhanced slow inactivation time courses to a double exponential function. Arec,f and Arec,s represent the relative proportions of the fast and slow components of current recovery from control slow inactivation. τrec,f and τrec,s are the time constants for the fast and slow recovery components. 135 The peak test current amplitudes for the experiments in Figs. 4.4A, B were normalized to the control current amplitude at the beginning of each respective sweep and plotted against the cumulative recovery time at −80 mV (Fig. 4.4C). Perhaps indicative of the inherent variability of inactivation between different preparations, the time course for the recovery of ShakerIR channels from control slow inactivation at pH 7.4 has been variously described as mono- exponential (Loots & Isacoff, 1998; Ray & Deutsch, 2006) or bi-exponential (Yellen et al., 1994; Olcese et al., 1997; González-Pérez et al., 2008). In our hands, recovery from control slow inactivation was better described by a double-exponential function (Fig. 4.4C) in which roughly 60% of the channels recovered via the slower pathway. In contrast, the time course of ShakerIR recovery from low pH-enhanced slow inactivation could be well fit by a mono-exponential function with a time constant (τrec = 2.13 ± 0.23 s; n = 5 cells) that was of the same order of magnitude as that of the fast component of recovery from control slow inactivation (τrec,f = 3.14 ± 0.29 s in the same 5 cells). As with our previous findings with Kv1.5 and low pH (see Section 3.3.8), the recovery from control slow inactivation and H + -enhanced slow inactivation could be simultaneously fit to a double exponential function. The global fit parameters for recovery from control and H + -enhanced slow inactivation in 5 cells are summarized in Table 4.1 and show that: 1) after a 5 s exposure at pH 4.0 the extent of enhanced slow inactivation (1 – A0; where A0 is the normalized residual current) at pH 4.0 is greater than that achieved after a 10 s inactivating pulse at pH 7.4; 2) following slow inactivation at pH 7.4, channels are roughly evenly distributed between the fast and slow recovery pathways (Arec,f ≈ Arec,s), and; 3) recovery from H + -enhanced slow inactivation is entirely via the fast pathway. Together, these results suggest that ShakerIR channels, like Kv1.5, can visit at least two slow inactivated states at pH 7.4 and that low pH promotes inactivation at +50 mV to the state from which recovery is fast. 136 Table 4.1. Low pH biases ShakerIR inactivation to a state from which recovery is fast.  pH 7.4 pH 4.0  inactivation at +50 mV inactivation at +50 mV A0 0.17 ± 0.03 0.06 ± 0.02 Arec,f 0.41 ± 0.06 1.00 τrec,f 2.33 ± 0.18 s * Arec,s 0.59 ± 0.06 0 τrec,s 18.29 ± 3.94 s * * constrained to be the same as control (pH 7.4) values A0 represents the residual current at the end of a 5-10 s pulse to +50 mV Arec,f and Arec,s represent the proportion of the fast and slow recovery components, respectively. Data are from experiments similar to those described in Figure 4.4 and are shown as mean ± S.E.M. for 5 cells.   In summary, the data of Figs. 4.1 – 4.2 show that external H+ enhances the rate and extent of the current decay at +50 mV. The similarity of the time constant for recovery from H + - enhanced slow inactivation to that for the fast phase of recovery from control slow inactivation at pH 7.4 (Fig. 4.4) suggests that low pH biases inactivation to the less stable of two states visited following slow inactivation in control solution. An additional H + - and time-dependent resting inactivation process at −80 mV appears to contribute to the loss of conductance (Fig. 4.3); however, the very fast time course of slow inactivation at low pH and the comparatively slow time course of our perfusion system prevents a precise characterization of the kinetics of resting inactivation.  137 4.3.5 The open probability of fast-inactivating T449 mutants is [K+]o-dependent The ability of low pH to enhance slow inactivation and decrease the macroscopic peak current of wt ShakerIR channels has striking similarities to the previously reported effects (at pH 7.4) of decreasing [K + ]o on fast-inactivating ShakerIR T449K, -A and -E mutants expressed in Xenopus ooctyes (López-Barneo et al., 1993). Figure 4.5 shows the effects of varying [K + ]o on ShakerIR T449K and T449A mutants, respectively, and confirms the findings of López-Barneo et al. (1993). For unknown reasons, T449E channels did not express in tsA201 or HEK-293 cells. Panels A and B of Fig. 4.5 show superimposed current traces recorded from cells expressing the T449K and T449A mutants, respectively, in solutions of varying [K + ]o. In standard pH 7.4 solution with 3.5 mM K + o the rate of slow inactivation was considerably enhanced in the T449K (τinact = 39.0 ± 0.7 ms; n = 5) and T449A (τinact = 187 ± 11 ms; n = 5) mutants, compared to wt ShakerIR (τinact = 1850 ± 400 ms; n = 40). Elevating [K + ]o very modestly increased the time constant of slow inactivation for both mutants; in 140 mM K + o, τinact ≈ 100 ms for T449K while τinact ≈ 200 ms for T449A in 10 mM K + o (the pulse length was too short to measure τinact in 140 mM K + o), consistent with earlier reports (López-Barneo et al., 1993). Far more striking is the dependence of the peak current amplitude on [K + ]o. Despite the large reduction in the outward driving force for K +  movement, maximal (or near-maximal) current in either mutant was observed in 140mM K + o. Conversely, the T449K mutant showed a nearly complete loss of current when the [K + ]o was reduced to 0 mM; the current reduction was ≈50% for the T449A mutant in 0 K+o. In contrast, there is little effect of removing K + o on the peak amplitude of wt ShakerIR currents (data not shown; see also López-Barneo et al., 1993; Baukrowitz & Yellen, 1995). 138     Figure 4.5. The effect of external [K + ]o on the amplitude of macroscopic currents through fast-inactivating ShakerIR mutants. A and B, Superimposed current traces recorded from cells expressing either ShakerIR T449K (A) or T449A (B) channels. Each trace was recorded with a test pulse from −80 mV to +50 mV in pH 7.4 solutions with the indicated concentrations of K+o. C, The peak current amplitudes from experiments such as those in (A and B) were normalized to the maximum peak current amplitude (at 10 mM K + o for T449A and 140 mM K + o for T449K) and plotted against [K + ]o. Each data point represents the mean ± S.E.M. of measurements from at least 3 cells. Solid lines represent fits of the data to the Hill Equation, parameters for which are shown on the plot. Dashed lines represent the normalized peak current amplitude observed with test pulses applied in nominally K + -free bath solutions. In the fits of the data to the Hill Equation, the maximum normalized current amplitude was a free parameter; that is, ymax in Equation 4.1 was not constrained to be equal to 1.  139 Figure 4.5C summarizes the [K + ]o-dependence of the normalized peak current amplitude through ShakerIR T449K and T449A. The values from the fits of the data for the T449K mutant (Kd = 4.8 mM) and the T449A mutant (Kd = 2.7 mM) to the Hill equation are slightly different from those previously reported (Kd = 8.3 mM and 0.8 mM, respectively; López-Barneo et al., 1993). Importantly, the qualitative effect of a [K + ]o-dependent loss of current was preserved, although in our hands the T449A mutant showed less than 50% current loss when switching from 10 mM to 0 mM K + o. It is unlikely that the loss of current induced by decreasing [K + ]o from 10 mM to 0 mM K + o is due to the relatively modest increase in the rate of slow inactivation. We next investigated the hypothesis, first proposed by López-Barneo et al. (1993), that the loss of current through fast-inactivating ShakerIR T449 mutants reflects a loss of channel availability, (perhaps due to a slow inactivation process) occurring at the resting potential. 4.3.6 ShakerIR T449 mutants undergo time-dependent resting inactivation in 0 mM K+o The time course for the resting inactivation of the T449K and T449A mutants was assessed as described for wt ShakerIR channels at low pH. For the superimposed sweeps of Fig. 4.6 a test pulse to +50 mV was applied at increasing intervals after the switch from a 10 mM K +  to a 0 mM K +  solution at pH 7.4. Shown for comparison is a control sweep recorded at the beginning of the experiment. Both the T449K and T449A mutants exhibited a mono-exponential decline in the peak test current amplitude as the time spent in 0 mM K +  increased. For T449K channels, resting inactivation was nearly complete (84 ± 2%) and τRI at −80 mV was 203 ± 11 ms (n = 7). In agreement with Fig. 4.5, in the T449A mutant the time course (τRI = 8.81 ± 3.54 s) of the current loss was slower and the extent (35 ± 11%; n = 4) of resting inactivation was less. It should be noted that for both mutants the rate of slow inactivation in 0 mM K + o did not change between sweeps (e.g. first sweep-τinact ≈ last sweep-τinact ≈ 40 ms for T449K and 160 ms for T449A) and was similar to that observed in 10 mM K + o (see above). Thus, enhanced slow 140 inactivation during the test pulse cannot account for the progressive decline in peak current amplitude. The results therefore confirm that removing K + o induces a time-dependent decrease in peak current consistent with a decrease in channel availability occurring at the resting potential.   Figure 4.6. Removing external K +  induces a time dependent decline in current amplitude in fast-inactivating ShakerIR mutants. Each panel shows superimposed current traces recorded from cells expressing ShakerIR T449K (A) or T449A (B) channels. In each sweep a 500 ms test pulse from −80 mV to +50 mV was applied following a known interval after the switch from the control 10 mM K + , pH 7.4 solution to a 0 mM K + , pH 7.4 solution. Also shown are control current traces recorded in 10 mM K +  at the beginning of each experiment. As the duration of exposure to 0 mM K +  solution increased, the peak current amplitude of the test pulse decreased. Dashed lines represent fits of the peak test current amplitudes () to a mono-exponential function of time.  141 4.3.7 Recovery of T449 mutants from 0 K+o-induced resting inactivation has the same time course as recovery from slow inactivation To assess whether the resting inactivation process underlying the loss of peak current in 0 mM K + o  might be mechanistically related to slow inactivation, we next compared the recovery time courses for the two processes. The recovery of the T449K (Fig. 4.7A) and T449A (Fig. 4.7D) mutants from slow inactivation was measured in multiple sweeps using a conventional double pulse protocol, with test pulses applied at increasing intervals following a conditioning pulse to +50 mV in 10 mM K + o. Then, in the same cell, following a control pulse in 10 mM K + o, resting inactivation was induced by switching to 0 mM K +  solution for 2 s (T449K; Fig. 4.7B) or 30 s (T449A; Fig. 4.7E) and recovery was followed by applying a test pulse at increasing intervals upon the return to 10 mM K + o. For each experiment, the peak test current amplitude in each sweep was then normalized to its control current and plotted against the interpulse interval (Figs. 4.7C and F). From fits of the data to a mono-exponential function, the time constant for the recovery of T449K from slow inactivation was 2.85 ± 0.28 s for T449K (n = 8) and the time constant for recovery from resting inactivation was 2.25 ± 0.25 s, which was not significantly different (P > 0.05). The time constants for the recovery of the T449A mutant from slow inactivation (12.55 ± 0.57 s; n = 5) and resting inactivation (13.37 ± 3.34 s) were also not significantly different (P > 0.05). In both mutants, the similarity of the recovery time courses following slow inactivation at +50 mV and resting inactivation induced by 0 K + o at −80 mV suggest that removal of external K +  promotes resting inactivation of T449K and T449A channels to a state that is also visited following slow inactivation and, consequently, that the two inactivation processes are related. 142  Figure 4.7. The recovery of T449K and T449A ShakerIR mutants from slow inactivation and resting inactivation induced by 0 K + o follows the same time course. A and D, Superimposed current traces recorded from cells expressing ShakerIR T449K (A) and T449A (D) show the recovery from slow inactivation in 10 mM K + o, at pH 7.4. For each trace, a conditioning pulse to +50 mV was applied to induce slow inactivation. Recovery was then monitored with a test pulse to +50 mV; the interval between the conditioning and test pulse increased with each successive sweep. B, Recovery of T449K channels from resting inactivation was monitored with multiple sweeps in a similar manner. Following a control pulse, resting inactivation was induced by switching from 10 mM to 0 mM K +  solution for 2 s. Test pulses to monitor recovery were applied upon the return to 10 mM K +  solution; the interpulse interval increased with each successive sweep. C, The peak test currents from panels A and B were normalized to their respective peak control currents and plotted against the recovery time at −80 mV. The solid and dashed lines represent fits of the data to mono-exponential functions. E, Resting inactivation of T449A channels was induced by a 30 s exposure to 0 mM K +  solution. Test pulses to monitor the recovery were applied following the return to 10 mM K + o; the time interval between the return to 10 mM K +  and the application of the test pulse increased with each sweep. Control currents were not recorded at the beginning of each sweep, due to the slow recovery time course of this particular mutant. F, The peak test currents from D were normalized as described for C and plotted against the recovery time. Peak test currents from E were normalized to the maximal test current and also plotted against recovery time. The dashed and solid lines represent mono-exponential fits of the data. The current traces shown in A and B were recorded from the same cell, as were the traces of panels D and E. 143 4.3.8 Recovery from slow inactivation and resting inactivation follow the same time course in the Kv1.5 H463G mutant Recapitulating the findings with the fast-inactivating ShakerIR T449 mutants in low [K + ]o (López-Barneo et al., 1993), in the fast-inactivating Kv1.5 H463G mutant the switch to K + - free solution at pH 7.4 causes a loss of current, which we have also referred to previously as a conductance collapse (Jäger & Grissmer, 2001; Kehl et al., 2002; Zhang et al., 2005). Although not yet investigated at the single channel level, it seems likely that this conductance collapse involves resting inactivation, and the [K + ]o-dependence of the channel availability in this mutant provides an opportunity to further explore the relative time courses of recovery from slow inactivation and resting inactivation in Kv1.5 independent of changes of pH or [Ni 2+ ] (see Chapter 3). The time constant, measured as described in Fig. 4.6, for the near complete loss of peak current associated with decreasing [K + ]o from 3.5 mM to 0 mM was 0.92 ± 0.09 s (n = 7). Importantly, decreasing [K + ]o from 3.5 mM to 0 mM has relatively small effects on the time constant of slow inactivation for the H463G mutant (τinact = 75.9 ± 1.7 ms, n = 12 at +50 mV in 3.5 mM K + o; τinact = 63.1 ± 3.0 ms, n = 4 in 0 mM K + o), such that the decrease in peak current cannot be ascribed to the acceleration of slow inactivation. Because the current decay is rapid at pH 7.4 in the H463G mutant (τinact ≈ 65 ms at +50 mV; Fig. 4.8A), which is believed to be due to an acceleration of slow inactivation (Zhang et al., 2005; Eduljee et al., 2007), it was necessary in the characterization of the recovery kinetics to use a paired pulse protocol to avoid cumulative inactivation that occurred with trains of 20 ms test pulses (not shown). The superimposed traces of Fig.4.8A were recorded at pH 7.4 in 3.5 mM K + o and a 500 ms conditioning pre-pulse from −80 mV to +50 mV caused more than 90% inactivation. In the first sweep, this conditioning pulse was followed 50 ms later by a 200 ms test pulse to +50 mV; for each successive sweep the interpulse interval was doubled out to a 144 maximum interval of 6.4 s. For the traces of Fig. 4.8B, which were obtained from the same cell, a 20 ms pre-pulse to +50 mV was applied in K + -free solution at pH 7.4, after which [K + ]o was increased to 3.5 mM and a 20 ms test pulse to +50 mV was applied at an interval after the start of the K +  pulse that was varied between sweeps from 0.05 to 6.4 s. At the end of each sweep the external solution was changed back to K + -free solution and the intersweep interval of 10 s was sufficient to ensure that the steady-state level of resting inactivation was re-established before the next sweep started. The graph of Fig. 4.8C shows the normalized peak test currents of the traces of panels A and B plotted against the interpulse interval, as well as the lines representing the best fit of the two data sets to a mono-exponential function. It indicates that the time course of recovery from slow inactivation () and resting inactivation () is virtually the same. In six such experiments, the mean  values of 1.46 ± 0.09 s (n = 6) and 1.43 ± 0.17 s for recovery from slow inactivation and resting inactivation, respectively, were not significantly different (P > 0.05) and support the conclusion that the recovery pathways for slow inactivation and the 0 K + o- induced loss of current are similar in Kv1.5 H463G. 145    Figure 4.8. Recovery at pH 7.4 of the Kv1.5 H463G mutant from slow inactivation and resting inactivation follow similar time courses. A, To monitor recovery of H463G mutant channels from slow inactivation in 3.5 mM K + o, a two pulse protocol was used. Briefly, slow inactivation was induced with a 500 ms pulse to +50 mV, followed by a 200 ms test pulse to the same potential, applied at increasing intervals in subsequent sweeps (8 superimposed sweeps are shown). B, In the same cell, to measure recovery from resting inactivation induced by 0 mM K + o, a 20 ms control pulse was applied in 0 mM K + o, followed immediately by a switch to 3.5 mM K +  and a subsequent test pulse to monitor recovery. The interval between the solution change and the test pulse was increased in each of 7 subsequent sweeps. C, The peak test currents for recovery from slow inactivation and resting inactivation, normalized to the peak control current at the beginning of each sweep are plotted against the interpulse interval at −80 mV. Both data sets were well fit by a single exponential function and had similar time constants. A simultaneous fit of the data to a single exponential gave a time constant of 1.48 s (not shown). The findings are representative of 6 experiments. 146 4.4 DISCUSSION We have characterized, for the first time, the kinetics of the onset of and recovery from the loss of current induced by low pH or removal of [K + ]o on wt or T449 fast-inactivating mutant ShakerIR channels, respectively. These experiments were designed to address two questions: whether the low pH or [K + ]o-induced loss of peak current through wt ShakerIR or T449 mutants, respectively, can be attributed to an inactivation process occurring at rest (−80 mV) and, if so, whether this resting inactivation represents slow inactivation involving the outer pore that has become uncoupled from channel activation. While further study is required, the results suggest that ShakerIR channels may undergo a resting inactivation process involving the outer pore mouth that is modulated by external H +  and K +  ions. A similar resting inactivation process is also suggested for the [K + ]o-dependent loss of Kv1.5 H463G channel availability. 4.4.1 Decreasing pH or [K+]o enhances slow inactivation of wt or mutant ShakerIR channels, respectively The results of Figs. 4.1 and 4.2 confirm the earlier reports that external acidification enhances both the time course and extent of ShakerIR current decay and support the view that this effect reflects an enhancement of the rate of slow inactivation (Pérez-Cornejo, 1999; Starkus et al., 2003; Claydon et al., 2007). Similar to results with Kv1.5, recovery from H + -enhanced slow inactivation occurred following a transient decrease in pH during a prolonged depolarization to +50 mV (Fig. 4.2). In contrast to our findings with Kv1.5 (Fig. 3.3A), the recovery current gradually returned to the control level without any overshoot. Thus, the model used to describe the enhancement of slow inactivation of Kv1.5 (Fig. 3.9), whereby acidification promotes entry to the faster recovering of two slow inactivated states cannot be applied here. An alternate, and perhaps simpler, explanation for this recovery in ShakerIR is provided by 147 revisiting SCHEME 3.2 of Fig. 3.1, where H +  ions cause a concentration-dependent shift of the O  OI equilibrium towards the slow inactivated (OI) state; consequently, the recovery observed upon the return to pH 7.4 reflects a return to an equilibrium that is less strongly biased towards the slow inactivated state. Interestingly, the overall time course of recovery (at −80 mV) from H+-enhanced slow inactivation was faster than that from control slow inactivation at pH 7.4 (Fig. 4.4). However, the results of the global fits showed that the time constant for the recovery from slow inactivation at pH 4.0 matched that for the fast component of recovery from control slow inactivation, suggesting that this difference may be attributed to a H + -induced biasing of slow inactivation towards the faster recovering of two slow inactivated states. It is unclear why this is not reflected as a recovery overshoot following a transient exposure to low pH at depolarized potentials (see above; also Fig. 4.2), in contrast to what we have seen with Kv1.5 (Fig. 3.3). Nevertheless, the similarities of the recovery time constants following control and enhanced slow inactivation are consistent with previous reports showing that the low pH effect is mitigated by the T449V mutation (Starkus et al., 2003; Claydon et al., 2007) and suggestive of the involvement of slow inactivation, while the lack of an effect of H +  on the single channel conductance (Claydon et al., 2007) or instantaneous tail currents (Starkus et al., 2003) indicates that the enhanced current decay is not due to H + -mediated pore block. 4.4.2 Low pH may induce resting inactivation of wt ShakerIR channels It is unlikely that the loss of ShakerIR macroscopic peak current at low pH is due entirely to the very fast slow inactivation rate, especially since single channel recordings showing multiple null sweeps imply that the loss of current is due to channels entering an unavailable state at rest (Claydon et al., 2007). We have attempted to characterize the time course for the H + - induced loss of peak current amplitude at −80 mV (Fig. 4.3) and found this resting inactivation 148 to be generally slower than H + -enhanced slow inactivation at +50 mV. However, it remains to be seen whether the results accurately reflect the time course and extent of resting inactivation, given that: i) the rapid onset of slow inactivation during the test pulses may significantly contribute to the reduction in peak current; and, ii) the kinetics of the solution exchange were rate limiting such that the time constant and, therefore, relative contribution of enhanced slow inactivation to the current loss was not constant throughout the experiment. 4.4.3 Comparison with the H+-dependent current loss of Kv1.5 For reasons outlined above, the difficulty in separating the H + -induced resting inactivation of ShakerIR from H + -enhanced slow inactivation is in contrast to the effects of low pH on Kv1.5 and suggests that H + -induced resting inactivation might contribute less to the loss of peak current in ShakerIR compared to Kv1.5. Correspondingly, the pH-dependence of the effect on ShakerIR peak current (pKa = 4.81 in 3.5 mM K + o; Fig. 4.1C) is lower than that of Kv1.5 (pKa = 6.2 in 5 mM K + o; from Kehl et al., 2002), likely due to the absence of a histidine residue present in the Kv1.5 turret that has been suggested to be the H + -binding site (Steidl & Yool, 1999; Kehl et al., 2002). This is supported by the observation that the pH sensitivity of the ShakerIR F425H mutant (analogous to Kv1.5 H463) is right-shifted (pKa = 5.8 in 2mM K + o; Pérez-Cornejo, 1999). Based on the pKa for the pH effect in wt ShakerIR, the H + -binding site has been suggested to be the aspartate residue in the K +  channel signature sequence, GYGD (Pérez- Cornejo, 1999; Starkus et al., 2003). The H + -induced resting inactivation of Kv1.5 is antagonized by small (3.5 mM) elevations in [K + ]o (Fig. 3.3; Kehl et al., 2002). In contrast, preliminary experiments with moderate (≤10 mM) increases of [K+]o have shown little effect on the loss of peak ShakerIR current induced by low pH (data not shown). However, increasing [K + ]o from 3 mM to 99 mM completely inhibited the H + -induced loss of fluorescence from a fluorophore tagged at S424C in 149 the outer pore of ShakerIR (Claydon et al., 2007). This latter finding suggests that high concentrations of external K +  ions can inhibit the H + -induced loss of channel availability at rest, such that channels may still undergo outer pore rearrangements associated with slow inactivation (and reported by the S424C fluorophore) upon depolarization. To further address whether H + - induced resting inactivation involves the outer pore, a possibility for future experiments would be to assess whether decreasing the pH at resting potential (−80 mV) has the same effects as depolarization-induced slow inactivation on Ba 2+  binding to ShakerIR channels. That is, does H + -induced resting inactivation impede the entry and exit of Ba 2+  to and from a deep pore binding site in the same way that  slow inactivation does (Basso et al., 1998; Harris et al., 1998)? 4.4.4 Enhanced slow inactivation in the ShakerIR T449 and Kv1.5 H463G mutants The rate of slow inactivation in the ShakerIR T449K and T449A mutant channels was, as reported earlier (López-Barneo et al., 1993), considerably faster than that of the wild-type channel in 3.5 mM K + o at pH 7.4 (Fig. 4.5). The effect of changing [K + ]o on the slow inactivation time courses of the T449K and T449A mutants was, compared to the effect of pH on wt ShakerIR, relatively moderate and, as such, unlikely to account for the large reductions in peak current amplitude that accompanied reductions in [K + ]o from 10 mM to 0 mM. Similarly, mutations of the histidine residue at position 463 in the Kv1.5 turret have also been shown to modulate the slow inactivation rate and confer sensitivity to changes in [K + ]o; while we have looked exclusively at the H463G mutant here, the H463R, -K, -A and -E mutants also show significantly accelerated slow inactivation and a loss of peak current amplitude in 0 K + o (Kehl et al., 2002; Zhang et al., 2005; Eduljee et al., 2007). As with the ShakerIR T449 mutants, an acceleration of slow inactivation (see section 4.3.8) in the H463G mutant is unlikely to account for the near complete loss of current observed when switching from 3.5 mM to 0 mM K + o. We did not examine further the modulatory effects of [K + ]o on slow inactivation at depolarized 150 potentials, which are thought to be due to the ability of K +  to impede, by a foot-in-the-door effect, the constriction of the selectivity filter (Baukrowitz & Yellen, 1995; Baukrowitz & Yellen, 1996; Kiss & Korn, 1998). 4.4.5 K+o-dependent resting inactivation of fast-inactivating ShakerIR and Kv1.5 mutant channels Although it had previously been suggested that the loss of current exhibited by ShakerIR T449 fast-inactivating mutants in low [K + ]o may be due to the entry of channels into a resting (closed) inactivated state (López-Barneo et al., 1993), this possibility had not been systematically studied. As with low pH and wt ShakerIR, we have characterized the time dependent decrease in the peak current amplitude of ShakerIR T449K and T449A mutants with removal of [K + ]o  (Fig. 4.6). It is unclear why, in contrast to the previous report (López-Barneo et al., 1993) where the peak current was reduced by ≈90%, the loss of current through ShakerIR T449A channels was incomplete (≈50%) in 0 mM K+o. Nevertheless, the effect of 0 K + o on the peak current amplitude of the T449K and T449A mutants cannot be attributed to an acceleration or accumulation of slow inactivation, since the effect of decreasing [K + ]o on slow inactivation is small and significant current loss is observed on the first pulse following prolonged periods in 0 K + o solution at −80 mV. The recovery of the T449K and T449A mutants from 0 K + o-induced resting inactivation was measured in 10 mM K + o and found to follow the same time course as recovery from control slow inactivation in 10 mM K + o (Fig. 4.7). Consistent with our hypothesis, these results suggest that the T449 mutant channels undergo a K + o-dependent resting inactivation process that may be akin to slow inactivation occurring at rest. In a similar fashion, the time-dependent decrease in the peak current amplitude of Kv1.5 H463G channels upon removal of K + o was characterized. We propose that this loss of channel availability in 0 K + o is due to slow inactivation occurring at rest, based on the finding that the 151 time courses for the recovery from slow inactivation and the putative 0 K + o-induced resting inactivation are the same, which implies that the resting inactivated state visited in 0 K + o at −80 mV is analogous to the slow inactivated state at +50 mV. These results are consistent with the previous conclusions that the Kv1.5 H463G mutant is permanently inactivated in K + -free solutions, based on the observation of robust Na +  tail currents similar to those observed through slow inactivated wt Kv1.5 channels or permanently inactivated Kv1.5 W472F mutant channels (Zhang et al., 2005). 4.4.6 Comparisons with [K+]o-dependent current loss in other channels The [K + ]o-dependent resting inactivation of the ShakerIR T449K mutant might be considered analogous to the 0 K + o-induced loss of availability observed with Kv1.4 channels, which have a lysine residue at the positional equivalent of T449, suggesting that this channel may also undergo resting inactivation (Pardo et al., 1992; Jäger & Grissmer, 2001). Indeed, the amount of gating charge displaced by Kv1.4 channels is invariant in solutions of different [K + ]o, indicating that channels gate normally in low [K + ]o but are unable to conduct current, consistent with a putative resting inactivation process (Pardo et al., 1992). Similarly, elevating [K + ]o also allows for (partial) recovery from the resting inactivation of wt Kv1.5 channels at low pH or in Ni 2+  (Chapter 3; Fig. 3.8). Intriguingly, the Kd for the K + o-dependent relief of the H + -induced resting inactivation (≈1 mM) is similar to that for the K+o-dependent rescue of the Kv1.5 H463G conductance (Kehl et al., 2002; Zhang et al., 2005), suggesting that perhaps a similar K + -binding site and/or resting inactivation process may be involved in both effects. Since the residues G380 and H404 in mouse (m)Kv1.3 are the positional homologues of H463 and R487, respectively, in Kv1.5, the mKv1.3 H404R mutant can be considered analogous to Kv1.5 H463G. Indeed, similar to the H463G mutant, rapid inactivation and a conductance collapse upon removal of K + o is observed in both mKv1.3 H404R and H404N mutant channels (Jäger et al., 1998). However, 152 given that the time course of mKv1.3 H404N current recovery upon re-addition of K + o was much faster than that following slow inactivation, this effect was deemed to be due to a different, unidentified process (Jäger et al., 1998). While it is clear that the [K + ]o-sensitive loss of channel availability depends on the residue at position 449 in Shaker (or its positional equivalent in other channels), the exact mechanism for this effect remains unclear. It has been suggested that charged residues, as with the T449K mutation, may repel K +  ions and decrease their ability to act as a “foot-in-the-door” to inhibit slow inactivation; consequently, the reduction of [K + ]o would be expected to have a greater ability to promote slow inactivation due to the vacancy of the selectivity filter (Molina et al., 1997). Support for this hypothesis is provided by the finding that in hKv1.3 channels, protonation of the H533 (homologous to Shaker T449) at low pH actually inhibits slow inactivation, purportedly due to the repulsive electrostatic forces of the protonated histidine that decrease the ability of the K +  ions to leave the pore (Somodi et al., 2004; Somodi et al., 2008). However, such electrostatic interactions cannot explain why similar effects on inactivation and [K + ]o-sensitivity are observed in the T449A, -S, and -Q mutants (López-Barneo et al., 1993; Schlief et al., 1996). Like the ShakerIR T449 mutants, the ability of mutations at position 463 in Kv1.5 to increase the rate of slow inactivation or influence the [K + ]o-dependence of current amplitude show little correlation to the properties of the substituted residues (Eduljee et al., 2007). In summary, the results presented here support the hypothesis that the low pH or [K + ]o induced loss of current through wt or mutant ShakerIR and Kv1.5 channels, respectively, involves a resting inactivation process. The similarity of the time courses for their recovery points to a similar underlying process for slow inactivation at depolarized potentials and resting 153 inactivation; however, further assessment is necessary to understand how external H +  and K +  ions are modulating outer pore inactivation at rest.  154 5 Discussion 5.1 SUMMARY OF RESULTS The properties of the slow inactivation of Kv channels have been studied extensively over the last twenty years. However, our understanding of this complex gating process is still, relative to channel activation and fast inactivation, quite limited. While there is now a general consensus that slow inactivation involves a localized constriction of the selectivity filter, several questions remain, such as how slow inactivation is regulated and how the voltage sensors are involved. The study of slow inactivation is complicated by its variability, even amongst members of the same Kv channel subfamily. Perhaps underlying the diverse phenotypic characteristics of slow inactivation is the considerable variation in the primary sequence of the outer pore regions of various Kv channels, particularly in the turret (S5-P linker; see Fig. 1.2A). Another example of structural heterogeneity is the low conservation, even amongst Kv1 sub-family members, of the residue at the position homologous to T449 in Shaker (Fig. 1.2), which has been shown to have a significant role in slow inactivation. Finally, investigations of slow inactivation typically involve assessments of changes in the conducting nature of the channel upon depolarization; examination of inactivation occurring from the resting state is further complicated by the non- conducting nature of the channel at all potentials. In this dissertation I have examined the hypothesis that external H +  and, in the case of Kv1.5, Ni 2+  ions cause a reduction in the peak current amplitude through wt Kv1.5 and ShakerIR channels by inducing a resting inactivation process that is mechanistically related to slow inactivation. The possibility that the low [K + ]o-induced reduction of peak current amplitude through fast-inactivating ShakerIR T449 and Kv1.5 H463G mutants is due to resting inactivation 155 was also assessed. The results show that at resting potential (−80 mV) and at low pH or in Ni2+, the outer pore of Kv1.5 channels undergoes a conformational change that inhibits Ba 2+  ion entry into as well as its exit from a binding site deep in the selectivity filter (Chapters 2 and 3). This finding, which is suggestive of a slow inactivation process involving the outer pore occurring at rest, is supported by the results of Chapter 3, in which the first characterization of the kinetics for the onset of and recovery from the effects of low pH and Ni 2+  applied at depolarized and resting potentials is presented. The results confirm that H +  and Ni 2+  enhance slow inactivation at +50 mV, and demonstrate that low pH and Ni 2+  also induce a resting inactivation process at −80 mV. A comparative analysis of the recovery time courses has shown that external H+ and Ni 2+  ions modulate inactivation (both at +50 mV and −80 mV) differently; H+ promotes inactivation to a less stable, quickly recovering state, while Ni 2+  biases inactivation to a more stable, slowly recovering state. However, the similarities of the time constants for recovery from enhanced slow inactivation and resting inactivation, particularly at low pH, suggest that resting inactivation is mechanistically similar to slow inactivation. Finally, a comparative analysis of the kinetics of the loss and recovery of current through wt ShakerIR induced by low pH or through fast-inactivating ShakerIR and Kv1.5 mutants induced by removing K + o, was performed (Chapter 4). As with wt Kv1.5, the results imply that the reduction in peak current induced by these manoeuvres is due, at least in part, to a process occurring at rest. Together, the findings presented here strongly suggest that external cations such as H + , Ni 2+  and K +  ions can regulate the availability of Kv1.5 and ShakerIR channels at rest, and that this regulation may involve an uncoupling of slow inactivation from channel activation, such that inactivation can occur at rest. 156 5.2 IN LOW pH OR Ni2+ THE OUTER PORE OF KV1.5 IS CONSTRICTED AT REST To my knowledge Chapter 2 describes the first assessment of the effect of externally applied Ba 2+  on Kv1.5 channels. As with ShakerIR (Hurst et al., 1995), the results indicate that in Kv1.5 there are two sequential binding sites, likely in the selectivity filter, although the affinity of these sites for Ba 2+  differs between the two channel types. It was also shown that there is a strong effect of test pulse frequency on the apparent affinity of the deep binding site for Ba 2+  (Fig. 2.3). Importantly, although it is not often considered, this shows that the effects of pulse frequency must be taken into account when comparing the dissociation constants for the superficial and deep Ba 2+  binding sites from different studies. A major finding from the Ba 2+  accessibility studies was that Ba 2+  binding and unbinding from Kv1.5 channels at −80 mV was impeded by external acidification (Fig. 2.7) or by the presence of external Ni 2+  (Fig. 3.5). Unfortunately, due to the very slow time course and incomplete nature of Kv1.5 slow inactivation, we were unable to assess Ba 2+  binding to or unbinding from slow inactivated Kv1.5 channels at pH 7.4. Thus, an important question remains whether the decreased accessibility of the deep Ba 2+  binding site induced by H +  and Ni 2+  at rest is similar to what might occur when Kv1.5 channels undergo slow inactivation in response to depolarization at pH 7.4. Nevertheless, the ability of a two-site sequential binding model developed for ShakerIR to describe Ba 2+  binding to Kv1.5 and the ability of K + o to impede Ba 2+  unbinding from Kv1.5, similar to a K + -sensitive “lock-in” site in ShakerIR and BK channels (Neyton & Miller, 1988b; Hurst et al., 1995; Harris et al., 1998), suggests that the overall properties of Ba 2+  binding are the same in ShakerIR and Kv1.5. Thus, the decrease in Ba 2+  accessibility at low pH or Ni 2+  has been taken to mean that a localized constriction of the selectivity filter has occurred at rest to decrease the continuity between the external solution and the deep Ba 2+  binding site, consistent with the properties of slow inactivation in ShakerIR at 157 depolarized potentials (Basso et al., 1998; Harris et al., 1998), and supportive of the hypothesis that H +  and Ni 2+  ions induce resting inactivation. Although it was not done here, it may be informative in the future to examine the Ba 2+  accessibility of wt ShakerIR at low pH, and of the ShakerIR T449K and Kv1.5 H463G mutants in 0 and 3.5 mM K + o, to determine whether the selectivity filter is also constricted under these conditions. In contrast to wt Kv1.5, the Kv1.5 H463G and ShakerIR T449K mutants inactivate rapidly and completely with depolarization at pH 7.4 in 3.5 mM K + o. In this way, comparisons between the Ba 2+  binding to or unbinding from resting inactivated and slow inactivated channels can be made directly. Ba 2+  accessibility studies with the ShakerIR T449A mutant in 0 K + o would likely encounter complications similar to those we saw with wt Kv1.5 at pH 7.4; that is, the incomplete loss of current in 0 K + o would make interpretation of changes in Ba 2+  accessibility due to resting inactivation difficult. 158 5.3 THE H+- OR Ni2+-INDUCED RESTING INACTIVATION OF KV1.5 OR ShakerIR Although reductions in Kv1.5 and ShakerIR peak current induced by external H +  or Ni 2+  ions had been reported before and suggested to be due to a resting inactivation process (Kehl et al., 2002; Kwan et al., 2004; Kwan et al., 2006; Claydon et al., 2007), to date a systematic examination of the development of this process at negative potentials had not been performed. Importantly, such kinetic information may provide clues to the mechanistic basis for resting inactivation when, for example, compared to the kinetics of depolarization-induced slow inactivation. It should be noted that the time course of changes in fluorescence purported to be associated with H + -induced rearrangements of the ShakerIR outer pore at −80 mV were reported in voltage-clamp fluorimetry experiments (Claydon et al., 2007). However, it was unclear whether these slow (over tens of seconds) rearrangements accurately reflect the time course of the H + -induced loss of conductance, or if they reflect the time course of the solution change in that recording system. By using fast perfusion changes to distinguish the effects of low pH or Ni 2+  on Kv1.5 at either depolarized or resting potentials, we have shown that the enhancement of slow inactivation alone cannot account for the loss in peak current amplitude. Instead, the channels undergo a time- and concentration-dependent resting inactivation process at −80 mV, which leads to a reduction in peak macroscopic current. These findings are consistent with single channel recordings of Kv1.5 showing that, in the presence of H +  or Ni 2+ , channels are not available to conduct current upon depolarization (Kwan et al., 2004; Kwan et al., 2006). The conclusion that H +  and Ni 2+  induce a Kv1.5 resting inactivation process similar to slow inactivation is also in agreement with our finding that Ba 2+  movement between its deep pore binding site and the external solution is impeded by high [H + ]o or [Ni 2+ ]o. In ShakerIR channels, although several factors are suggestive of a H + -induced resting inactivation, including the lower pKa for the decrease in peak current relative to that for the time constant of slow inactivation 159 (Chapter 4), as well as single channel and voltage-clamp fluorimetry measurements at −80 mV (Claydon et al., 2007), we have found that characterizing the time course of the decline of peak current due to this putative resting inactivation process is confounded by the greatly accelerated slow inactivation of ShakerIR associated with low pH values and the comparatively slow speed of the perfusion system. Thus, although a time and pH-dependent decline of ShakerIR current at −80 mV can be observed, it is difficult to discern precisely how much of that decline is due to enhanced slow inactivation and whether it accurately depicts the time course of resting inactivation. 5.3.1 A comparison of the kinetics of slow inactivation and resting inactivation An interesting observation was that the onset of resting inactivation of Kv1.5 at −80 mV was generally slower than that for slow inactivation at +50 mV and was accelerated as the [H + ]o or [Ni 2+ ]o increased. Based on the current data, it is difficult to come to a definitive, generalized conclusion regarding the kinetics of resting inactivation onset. However, under the assumption that resting inactivation is mechanistically similar to control slow inactivation at pH 7.4, the generally slower onset of resting inactivation, compared to slow inactivation, may be due to the absence, at −80 mV, of any allosteric contributions of activated voltage sensors. This arises from the observation that, under control pH 7.4 conditions, slow inactivation is considered to be strongly coupled to activation, such that it occurs primarily from the open state during depolarization (Hoshi et al., 1990; Rasmusson et al., 1995; Olcese et al., 1997). In this respect, the energy barrier for resting inactivation of Kv1.5 under control conditions is likely quite high. However, as the [H + ]o or [Ni 2+ ]o increases, the energy barrier for resting inactivation may decrease such that, at very low pH (pH 5.4) or high [Ni 2+ ]o, it progresses at a rate approaching that of slow inactivation. One would expect that similar arguments could be made for the H + - induced resting inactivation of ShakerIR. The lower pH values required to induce significant 160 resting inactivation of ShakerIR may be reflective of a greater energy barrier that needs to be surmounted and/or smaller decreases in the energy barrier in response to protonation. In support of the latter possibility, recall that ShakerIR lacks a histidine residue in the turret, which in Kv1.5 (H463), has been proposed to be the H +  and Ni 2+  binding site (Steidl & Yool, 1999; Kehl et al., 2002; Kwan et al., 2004). Mutating H463 in Kv1.5 to a glutamine (Q) diminishes the effect of low pH and Ni 2+  on peak current (Kehl et al., 2002; Kwan et al., 2004), while substituting a histidine residue for the homologous residue in ShakerIR (F425H) confers greater pH sensitivity of the peak current amplitude (Pérez-Cornejo, 1999). In contrast to the onset, the overall time courses for recovery from resting inactivation in Kv1.5 tended to be slightly faster than that from enhanced slow inactivation (see Fig. 3.6). Specifically, recovery from Ni 2+ -induced resting inactivation was faster than from Ni 2+ -enhanced slow inactivation. A simple explanation would be that this is also related to the lack of a modulating effect from the voltage sensors; however, one must keep in mind that, for H + -induced resting inactivation of Kv1.5, the time constants for the fast and slow components of recovery did not differ from those for recovery from H + -enhanced slow inactivation and the overall faster recovery is related to the stronger biasing, by H + , of inactivation to the faster recovering of two inactivated states. Two related questions arise from these findings: how are the fast and slowly recovering slow inactivated states of Kv1.5 different at the mechanistic and structural levels; and, how do H +  and Ni 2+ , both of which we presume are acting at H463 in the Kv1.5 turret, differentially bias the entry into these inactivated states? One question that has not been assessed is whether the effects of H +  or Ni 2+  involve a cooperative process; indeed, the Hill coefficient values for the low pH and Ni 2+ -induced reduction in peak current are >1, suggesting that cooperativity may be involved. It may therefore be informative to examine the H + - and Ni 2+ - induced resting inactivation in concatameric channels where the H463 residue of one or more 161 linked subunits is replaced by a glutamine, in a  manner analogous to that used by Yang et al. (1997) in their study of the ShakerIR W434F mutant. In this way it might be possible to determine whether the protonation of or binding of Ni 2+  to one H463 residue in the Kv1.5 channel is sufficient to induce resting inactivation. 5.3.2 Molecular mechanisms of H+-induced resting inactivation in Kv1.5 and ShakerIR 5.3.2.1 Modulation of the turret The greater sensitivity of Kv1.5 current to decreases in pH (pKa = 6.2 in 5 mM K + o; Kehl et al., 2002) compared to ShakerIR (pKa = 4.8 in 3.5 mM K + o; Chapter 4 ), shown here and previously by others (Pérez-Cornejo, 1999; Starkus et al., 2003; Claydon et al., 2007), suggests that the H + -binding sites underlying the pH-sensitive current loss are different in Kv1.5 and ShakerIR channels. As mentioned above, the likely H + -binding site in Kv1.5 is the H463 residue in the turret (Steidl & Yool, 1999; Kehl et al., 2002; Eduljee et al., 2007). ShakerIR lacks a histidine at this position and the H +  binding site has been suggested, based on the pKa value, to be an aspartate residue in the selectivity filter GYGD sequence (Pérez-Cornejo, 1999; Starkus et al., 2003). The importance of H463 in the turret in the resting inactivation of Kv1.5 is emphasized by the ability of external Ni 2+  and Zn 2+ , known histidine ligands, to induce a loss of channel availability with properties similar to those of H + -induced resting inactivation (Chapter 3; Zhang et al., 2001a; Zhang et al., 2001b; Kehl et al., 2002; Kwan et al., 2004). Additionally, the fast-inactivating Kv1.5 H463G mutant also shows a loss of channel availability in 0 K + o solutions that is very similar to the behaviour of the fast-inactivating ShakerIR T449 mutants and likely to be associated with resting inactivation (Chapter 4; Kehl et al., 2002; Zhang et al., 2005). Unfortunately, it is not yet clear how protonation of H463 or substitution of H463 by glycine leads to both enhanced slow inactivation and resting inactivation at the level of the selectivity 162 filter, nor how H +  and Ni 2+  bias inactivation differently. An indirect interaction with the R487 residue between the top of the selectivity filter and the P-S6 linker has been suggested (Jäger & Grissmer, 2001; Kehl et al., 2002; Trapani & Korn, 2003). A similar involvement of the analogous turret and pore residues in ferret (f)Kv1.4 (H508 and K532, respectively) has also been implicated in the H + -enhanced slow inactivation of this channel, which has been proposed to underlie the concomitant reductions of channel availability (Claydon et al., 2000; Claydon et al., 2002; Li et al., 2003). 5.3.2.2 Modulation of the selectivity filter As noted above, ShakerIR channels lack a protonatable histidine residue in the outer turret and thus, the H + -binding site mediating the enhanced slow inactivation observed at low pH may instead be the conserved aspartate residue (D447) at the top of the selectivity filter. This is the position in KcsA (D80) which has been proposed to interact with the W67 residues of the “aromatic cuff” (ShakerIR W434) to stabilize the selectivity filter in the conducting conformation (Doyle et al., 1998). It has also recently been suggested that inactivation in KcsA is inhibited by the E71A mutation, which breaks the inactivation-promoting D80-E71 interaction, and allows for a reorientation of D80 into a “flipped” conformation such that it points upwards towards the turret (Cordero-Morales et al., 2006; Cordero-Morales et al., 2007). Protonation of D80 has also been shown to disrupt the conformation of the external end of the selectivity filter, and this was suggested to be correlated with the inactivated state of the pore (Miloshevsky & Jordan, 2008). Although eukaryotic Kv channels do not possess a D80-E71 interaction, neutralization of D447 in ShakerIR, by mutation to asparagine, results in a non- conducting channel that exhibits similar properties (i.e. gating charge-voltage (Q-V) relationship, “rescue” of current by T449Y) to those of the permanently slow inactivated W434F mutant (Seoh et al., 1996b; Yang et al., 1997). Thus, the D447N mutant might also be a permanently 163 slow inactivated channel, supporting the hypothesis that protonation and, therefore, neutralization of the D447 charge is the basis for the H + -enhanced slow inactivation of ShakerIR shown here and by others (Starkus et al., 2003; Claydon et al., 2007). It may therefore be informative to test whether the D447N mutant shows a voltage-clamp fluorimetry profile similar to that of wt ShakerIR at low pH and the W434F mutant. That is, does a fluorophore at S424C (in the turret) in the background of the D447N mutant provide rapid reports on the movement of the S4 voltage sensor upon depolarization in a manner characteristic of that seen in slow inactivated channels? Furthermore, as suggested by Eduljee et al. (2007) an ability of the aspartate at this position in Kv1.5 (D485) to project towards the external milieu may provide another potential interaction site near the selectivity filter for the H463 residue in the turret. Although protonation of the highly conserved D447 in ShakerIR has been suggested to underlie the enhanced slow inactivation observed at low pH, and may also be involved in any H + -induced resting inactivation, it is interesting to note that in hKv1.3 the protonation of a nearby H399 (≈ShakerIR T449) residue at low pH actually mildly inhibits slow inactivation (Deutsch & Lee, 1989; Somodi et al., 2004; Somodi et al., 2008). In this Kv1 isoform it was suggested that, in physiological solutions, the resulting electrostatic barrier formed by the protonated H399 residues impede K +  exit from the pore, thereby inhibiting emptying of the selectivity filter and slowing the onset of slow inactivation; conversely, with inwardly directed currents, the protonated H399 residues impede K +  entry to the selectivity filter, facilitating its emptying and resulting in accelerated slow inactivation (Somodi et al., 2004; Somodi et al., 2008). However, a similar effect is not observed with ShakerIR T449H, which shows accelerated slow inactivation at low pH in both low and high [K + ]o (López-Barneo et al., 1993). Additionally, although the rate of Kv1.3 slow inactivation is decreased at low pH, a reduction in peak current amplitude perhaps related to a resting inactivation process was also noted at low pH 164 (with 5 mM K + o ; Somodi et al., 2004; Somodi et al., 2008). Intriguingly, it seems that this proposed resting inactivation of Kv1.3 at low pH can occur in low [K + ]o even though slow inactivation is inhibited by the delayed K +  exit rate due to H399 protonation; although it should be noted that the involvement of a slow inactivation process in the H + -induced current loss is suggested by the restoration of peak current amplitude with elevated [K + ]o (Somodi et al., 2008). Together, the results indicate that, while modification of the region near the outer mouth of the selectivity filter at low pH can contribute to the inactivation characteristics of Kv channels, it is also unlikely to be the sole determinant of slow inactivation, as suggested by the involvement of the outer turret in Kv1.5 proposed above. 165 5.4 RESTING INACTIVATION IN FAST-INACTIVATING MUTANTS IS HIGHLY SENSITIVE TO [K+]O The first indications that Kv channels may undergo a loss of availability due to inactivation from resting, closed states came from early studies of the [K + ]o-dependent decreases of the peak current amplitude of Kv1.4 channels (Pardo et al., 1992) and fast-inactivating ShakerIR mutants (López-Barneo et al., 1993). To date, most investigations have focused on the actions of K + o to inhibit slow inactivation from the open state at depolarized potentials. In contrast, the experiments presented in this dissertation have looked explicitly at the ability of K + o-removal to enhance the resting inactivation of fast-inactivating ShakerIR and Kv1.5 mutants at −80 mV. The results of Chapter 4 confirm that the ShakerIR T449A and -K mutants, as well as Kv1.5 H463G, undergo a time-dependent reduction in peak current amplitude and, thus, channel availability, in 0 K + o solution at −80 mV. The similarity between this 0 K + o-induced resting inactivated state and the slow inactivated state was inferred from the parallel time courses for recovery from resting inactivation (upon the return of K + o) and the recovery from slow inactivation. An analogous ability of external K +  to promote recovery of Kv1.5 from H + - or Ni 2+ - induced inactivation was observed (Fig. 3.8) and this recovery also has the same time course as that from slow inactivation. In contrast, the recovery of mKv1.3 H404N mutant channels from the current loss in 0 K + o was much faster than from slow inactivation, leading to the conclusion that 0 K + o conditions induced a transition into a closed, non-conducting state by an unidentified process distinct, at least by kinetic criteria, from slow inactivation (Jäger et al., 1998). The K + - sensitive resting inactivation described in this dissertation is also distinct from the “defunct” state that Shaker channels may enter in symmetrical K + -free solutions (Gómez-Lagunas, 1997; Melishchuk et al., 1998; Loboda et al., 2001), given that resting inactivation does not require long depolarization in 0 K +  and readily recovers at −80 mV with the return of K+o. 166 Returning to the question of resting inactivation in wt Kv1.4 channels, it is informative that Kv1.4 has a lysine residue at the position homologous to T449 in ShakerIR (see Fig. 1.2), which may account for its relatively rapid rate (amongst Kv1 family members) of slow inactivation and the similarly [K + ]o-dependent resting inactivation of the Kv1.4 and ShakerIR T449K channels. Conversely, Kv1.1 and Kv1.2 channels have, respectively, tyrosine and valine residues at this position and, similar to the “non-inactivating” T449Y and T449V mutants, inactivate very slowly upon depolarization and do not exhibit [K + ]o-dependent decreases in conductance (López-Barneo et al., 1993; Jäger et al., 1998). Interestingly, Kv4 channels are typically considered to undergo a preferential closed-state inactivation process distinct from slow inactivation (see Chapter 1.3.4), whereby increases in [K + ]o enhance inactivation (Jerng & Covarrubias, 1997; Kirichok et al., 1998). However, it has also been reported that in the absence of K + o, Kv4.3 exhibits accelerated inactivation and a decrease in peak macroscopic current with repetitive stimulation, which were attributed to a collapse/constriction of the selectivity filter in a manner reminiscent of slow inactivation (Eghbali et al., 2002). It has since been proposed that elevating [K + ]o may enhance preferential closed-state inactivation by inhibiting a “vestigial” slow inactivation process in Kv4 channels, thereby increasing the population of an unstable open state and the probability that channels inactivate from a pre-open closed state (Kaulin et al., 2008). It is unclear whether a 0 K + o-induced decrease in Kv4 peak current amplitude and channel availability also occurs at −80 mV in the absence of stimulation, similar to the proposed resting inactivation observed with Kv1.4 and the fast-inactivating ShakerIR and Kv1.5 mutants. 5.4.1 How does K+o modulate resting inactivation? The ability of external K +  to antagonize slow inactivation has been well described, and involves a “foot-in-the-door” mechanism whereby K+ occupancy of a site in the outer pore inhibits the constriction of the selectivity filter (Baukrowitz & Yellen, 1995; 1996; Kiss & Korn, 167 1998; Kiss et al., 1999). A similar effect likely accounts for the K + o-dependent slowing of the onset of the resting inactivation of Kv1.5 induced by external H +  or Ni 2+  ions and perhaps prevents resting inactivation of Kv1.4 channels. While a reduced occupancy of a selectivity filter K +  binding site is the simplest explanation for the enhanced slow inactivation (in low [K + ]o) and resting inactivation (in 0 K + o), an unanswered question remains: how does the return of K +  to the external milieu facilitate recovery from resting inactivation? This is particularly perplexing given the underpinning assumption that in the inactivated channel the selectivity filter is constricted and, consequently, that the accessibility of external K +  and Ba 2+  to binding sites within the selectivity filter is markedly reduced (Basso et al., 1998; Harris et al., 1998; Kiss et al., 1999). The voltage-dependent effects of K + o on the recovery of slow inactivated Kv1.3 channels has been attributed to the slow, voltage-dependent binding of K +  ions to a modulatory site that eventually results in a destabilization of the slow inactivated state (Levy & Deutsch, 1996a; Levy & Deutsch, 1996b). Although it appears that this modulatory site has an effective membrane electrical field distance of ≈0.3 and lies external to the constriction of the selectivity filter, its exact identity and location remains unknown (Levy & Deutsch, 1996a). Given that the binding of K +  at hyperpolarized potentials to the modulatory site in Kv1.3 tended to be slower than recovery from slow inactivation, it seems unlikely that the same process can be attributed to the K + o- facilitated recovery at −80 mV from resting inactivation described here and which tracks recovery from slow inactivation (e.g. Kv1.5 H463G or wt Kv1.5 at low pH), assuming that the similar recovery time courses from slow inactivation and resting inactivation are not coincidental. Because K +  facilitated recovery from resting inactivation can occur over wide a range of time courses (e.g. 2 s versus 20 s for Kv1.5 at low pH or in Ni 2+ , respectively), it is unlikely that the binding of K +  to inactivated channels is the rate-limiting step. However, K +  bound at a site 168 close to the slow inactivation gate may, upon recovery from inactivation, rapidly equilibrate with binding sites deeper within the selectivity filter and help stabilize the selectivity filter in a conducting conformation. Intriguingly, in Kv1.5 the Kd (≈1 mM) for the antagonism by K + o of the loss of availability of the H463G mutant (Zhang et al., 2005) or the wt channel at low pH (Kehl et al., 2002) is very similar to the apparent Kd (1 mM) for the inhibition of Ba 2+  unbinding by K + o (Chapter 2). This site may be homologous to the “lock-in” binding site for the K + - dependent Ba 2+  unbinding in ShakerIR (Kd = 0.75 mM) and BK channels (Kd = 0.3 mM) (Neyton & Miller, 1988b; Hurst et al., 1995; Harris et al., 1998) and/or an external low affinity K +  binding site (Kd ≈ 2 mM) in the selectivity filter that has been shown to indirectly modulate slow inactivation by “trapping” K+ ions within the filter (Kiss & Korn, 1998; Kiss et al., 1999; Ogielska & Aldrich, 1999). In the case of the ShakerIR T449 mutants, the wide range of Kd values for K + o-dependent resting inactivation (Chapter 4; López-Barneo et al., 1993; Schlief et al., 1996) may be due to the close proximity of position 449 to the external K +  binding site and a consequent effect of the residue at position 449 on K +  accessibility (see below). Given the potential location of the K +  binding site mediating recovery within the membrane electrical field, it may be of interest in the future to assess whether the K + -dependent recovery from resting inactivation is voltage-dependent. It is not clear how mutations at ShakerIR T449 or Kv1.5 H463 so drastically increase the sensitivity of these channels to K + o removal, as well as their susceptibility to inactivation, nor how an endogenous Lys residue (at the position equivalent to Shaker T449) imparts Kv1.4 channels with an intrinsic sensitivity to removal of K + o  at pH 7.4. In the case of the ShakerIR T449K and -E mutants, strong electrostatic effects on K +  have been suggested to cause a reduced K +  occupancy of binding sites in the outer selectivity filter (López-Barneo et al., 1993; Molina et al., 1997). However, charge effects clearly cannot account for the K + -sensitive inactivation 169 observed with the ShakerIR T449A, -S and -Q mutants (López-Barneo et al., 1993; Schlief et al., 1996) or with Kv1.5 H463G. A more generalized view may be that the residue at the position equivalent to Shaker 449 can affect the overall accessibility of external K +  to binding sites in the selectivity filter, perhaps by electrostatic and/or steric alterations. This is supported by the finding that substitution of T449 with -A, -V or -Y decreases the ability of externally applied Ba 2+  ions to reach the deep binding site in ShakerIR, while leaving the Ba 2+  exit rate from this site unchanged (Hurst et al., 1996). Consequently, it may be inferred that in the fast-inactivating mutants, the equilibrium for K +  occupancy of binding sites deep within the selectivity filter is biased towards those sites being vacant, such that inactivation (from both the open and resting states) is enhanced. However, one must be cautious in making direct correlations between the accessibility of the deep Ba 2+  binding site and the propensity for inactivation, given that Ba 2+  binding is inhibited in the T449V and -Y mutations, which also inhibit slow inactivation of ShakerIR (Hurst et al., 1996). Intriguingly, preliminary investigations in Kv1.5 have found that Ba 2+  binding to the superficial and deep sites is also drastically altered in the R487Y and R487V mutants (analogous to T449Y and T449V in ShakerIR; SJ Kehl, C Tong and YM Cheng, unpublished observations). Specifically, in contrast to wt Kv1.5 and more akin to the corresponding ShakerIR mutants, Ba 2+  binding to the superficial site is enhanced in the R487Y and -V mutants, while Ba 2+  binding to the deep site is slowed. Conversely, unlike ShakerIR, the R487V mutation has little effect on slow inactivation at pH 7.4, but does inhibit H +  or Ni 2+ - induced resting inactivation (Kehl et al., 2002; Kwan et al., 2004). Clearly, the residue at positions homologous to T449 in Shaker has important effects on both the accessibility of ion binding sites in the selectivity filter and on the rate of inactivation; however, there are still unanswered questions as to exactly how these effects come about. 170 5.5 IMPLICATIONS OF RESTING INACTIVATION Building upon the previous evidence, detailed in Chapter 1, several of the findings presented here imply that the resting inactivation observed at −80 mV may involve a process similar to slow inactivation occurring at +50 mV. In Kv1.5, these include the outer pore constriction induced at rest by low pH and Ni 2+ , the similarities in the time courses for recovery from slow and resting inactivation and the sensitivity of resting inactivation to [K + ]o. Although the examination of a H + -induced resting inactivation of wt ShakerIR was hampered by the concurrent acceleration of slow inactivation and by the limiting rate of solution change with our perfusion system, we were also able to show that the K + o-dependent recovery from resting inactivation of fast-inactivating ShakerIR T449 and Kv1.5 H463G mutants likely follows the same recovery pathway as slow inactivation. Additionally, that resting inactivation can occur at −80 mV, from the fully closed state, distinguishes it from U-type inactivation and preferential closed-state inactivation (see Chapter 1), which tend to require at least partial activation of the voltage sensors. The following sections address two major implications of the conclusion that slow inactivation can occur from the resting state: i) the common assumption that inactivation is strongly coupled to activation may no longer hold (at least under the conditions studied here), and; ii) the role of charge immobilization, and that of the voltage sensors in general, in slow inactivation is questioned. 5.5.1 Coupling between activation and slow inactivation The coupling between slow inactivation and activation is such that it occurs primarily from the open state during depolarization (Hoshi et al., 1990; Rasmusson et al., 1995; Olcese et al., 1997). Additionally, closure of the slow inactivation gate in the outer pore has been shown to result in conformational changes of the inner cavity of ShakerIR channels such that the open state of the intracellular activation gate is stabilized (Panyi & Deutsch, 2006). Additionally, the 171 accessibility of internally applied TEA or cysteine modifying agents (e.g. Cd 2+ , MTSET, MTSEA) to engineered binding sites in the inner pore and internal cavity is reduced when the channel is in the slow inactivated state (Panyi & Deutsch, 2006; Panyi & Deutsch, 2007). The assumption with resting inactivation occurring at −80 mV is that the conformational changes in the outer pore underlying the loss of channel availability are occurring while the inner activation gate is closed. However, it remains unknown whether the resting inactivation of Kv1.5 channels at low pH or in Ni 2+ , and of the fast-inactivating ShakerIR and Kv1.5 mutants in 0 K + o, is causing allosteric effects in the inner pore region similar to those seen with slow inactivation at depolarized potentials. Gating current measurements from Kv1.5 at low pH and in Ni 2+  show that there is little change in the total amount of charge moved during channel activation, which suggests that the activation pathway is relatively normal (Kehl et al., 2002; Kwan et al., 2004). Conservation of gating charge is also seen with wt ShakerIR at low pH (Claydon et al., 2007) and in the permanently inactivated ShakerIR W434F mutant (Perozo et al., 1993). I did not study and, to my knowledge, there is no information about the effects of resting inactivation on channel opening. Although a modulatory effect of resting inactivation on the conformation or gating properties of the activation gate and inner cavity would not be expected to affect the conducting state of the channel (given that the channel would still be in the non-conducting resting inactivated conformation), it would still be interesting to see if changes in the inner pore region, such as those described with slow inactivation, can occur. Not only would this allow further comparison between the resting and slow inactivated states, but it would also provide information regarding whether the conformational changes associated with resting inactivation are restricted to the selectivity filter and/or outer pore region alone. 172 5.5.2 Involvement of the voltage sensor in slow inactivation The long-standing association between slow inactivation and the S4 voltage sensors originated largely from observations of gating charge immobilization following prolonged depolarization and slow inactivation (Olcese et al., 1997; Chen et al., 1997). Voltage clamp fluorimetry measurements suggestive of a close approximation between the top of S4 and the outer pore during inactivation also support this view (Loots & Isacoff, 1998; Loots & Isacoff, 2000), as does the crystal structure for Kv1.2 showing the proximity of the top of S4 and the outer pore region of a neighbouring subunit (Long et al., 2005a). Consequently, slow inactivation is often considered to progress in at least two steps: an outer pore constriction and a stabilization of the voltage sensors in an activated conformation. However, our finding that slow inactivation may become uncoupled from channel activation argues against an obligatory involvement of the voltage sensors in inactivation, and provides further support to an emerging view that charge immobilization may represent an intrinsic property of the voltage sensors at depolarized potentials (Villalba-Galea et al., 2008). Although the resting inactivation of Kv1.5 and ShakerIR described here was dependent on external acidification, exposure to Ni 2+ , removal of [K + ]o or mutations at the outer pore mouth or turret, the potential [K + ]o-dependent resting inactivation of wt Kv1.4 channels in physiological solutions (Pardo et al., 1992) speaks to the possibility of a broader role of resting inactivation in the regulation of Kv channel activity. A question that arises is why, if voltage sensor movement and slow inactivation are independent processes, the recovery time courses from both processes track each other so closely (in ShakerIR) (Olcese et al., 1997; Loots & Isacoff, 2000)? One possibility is that this correlation is coincidental. This is supported by the finding, in Kv1.5, that the return of gating charge following inactivation is much faster than the recovery of the pore from slow inactivation (Wang & Fedida, 2002), which implies an uncoupling of the two processes and, perhaps, independent 173 recovery pathways. An alternative explanation, albeit one requiring further exploration, is that perhaps the parallel time courses for recovery is related to the coupling of both slow inactivation and the immobilized voltage sensors to the status of the intracellular activation gate. As discussed above, slow inactivation has been shown to stabilize the open state of the intracellular activation gate (Panyi & Deutsch, 2006; Panyi & Deutsch, 2007). It has also recently been proposed that charge immobilization is dependent on an inter-subunit interaction between the S4- S5 linker and S6 that stabilizes both the open state of the activation gate and contributes to the immobilization of the voltage sensors (Batulan et al., 2010). Thus, it is possible that the recovery from both slow inactivation and charge immobilization is dependent on the closing of the activation gate. Intriguingly, this may explain why the overall time course for recovery from resting inactivation tends to be faster than that for recovery from slow inactivation, since the activation gate is (assumed to be) closed in the resting inactivated state. 174 5.6 FINAL REMARKS Although the concept of resting inactivation has previously been invoked to account for the loss of channel availability associated with the removal of [K + ]o (Pardo et al., 1992; López- Barneo et al., 1993; Jäger et al., 1998; Zhang et al., 2005); with the irreversible loss of ionic current following the W434F mutation in the ShakerIR pore helix (Yang et al., 1997); and with the reversible effects of low pH or Ni 2+  (Kehl et al., 2002; Kwan et al., 2004; Kwan et al., 2006; Claydon et al., 2007), the mechanistic relationship between resting and slow inactivation was still unclear. The studies described in this dissertation examined, for the first time, the kinetics for the onset of and recovery from the resting inactivation of wt and mutant Kv1.5 and ShakerIR channels induced by manoeuvres including low external pH, application of external Ni 2+ , and removal of K + o. Together, the findings support the hypothesis that slow inactivation of Kv1.5 and ShakerIR channels may, under certain conditions, uncouple from activation, such that it occurs at rest. Further study is required to elucidate the molecular movements underlying resting inactivation, which appears to involve interactions within the outer turret and the selectivity filter, as well as whether resting inactivation affects the channel in areas other than the outer pore. The finding that inactivation can occur from the resting state raises questions about the “dogma” of coupling between slow inactivation and activation, as well as any role of the voltage sensors. Intriguingly, the K + o-and pH-dependence of resting inactivation may present an avenue for the regulation of Kv channel availability that is use-independent, in contrast to slow inactivation, which requires channel activation. With respect to the role of Kv1.5 in cardiac electrophysiology, given that significant resting inactivation and enhanced slow inactivation are observed at pH 6.9 in 0 K + o, a slight decrease in extracellular pH may result in physiologically relevant decreases in IKur, which contributes to the repolarization of the atrial action potential 175 (Fedida et al., 1993; Feng et al., 1997). This may be important during episodes of cardiac acidosis, perhaps related to ischemia, during which resting inactivation of Kv1.5 could potentially result in a decrease of the total repolarizing current and a prolongation of the action potential, perhaps decreasing the risk of atrial fibrillation. However, during ischemic heart attack, accumulation of K +  in the extracellular fluid would be expected to antagonize the H + -induced loss of Kv1.5 current. As suggested by Trapani and Korn (2003), small decreases in pH may also reduce Kv1.5 current due to the H + -induced depolarizing shift of the conductance-voltage relationship. Of course, in the presence of other channels and transporters in native cardiac tissue, the functional outcome of acidosis is likely to be more complicated. Nevertheless, the prolongation of the atrial action potential is a well-established principal in the treatment and prevention of atrial fibrillation (for reviews, see Brendel & Peukert, 2003; Ehrlich et al., 2008; Ford & Milnes, 2008; Ehrlich & Nattel, 2009) and Kv1.5 presents an attractive target for pharmacological intervention, based on its selective expression in human atrial, but not ventricular, tissue (Wang et al., 1993; Li et al., 1996). 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Two different systems were used: in Chapters 2 and 4, the commercially available Warner Instruments SF-77B Perfusion Fast Step system was optimized for use; while in Chapter 3 a custom fast application tool (FAT) was made by Dr. Steven J Kehl. The following sections detail the construction, optimization and characterization of each system. A.1.1 The Warner Instruments SF-77B Perfusion Fast Step Figure A.1 shows a photograph of the SF-77B Perfusion Fast Step system used in the experiments of Chapters 2 and 4. The major components of the apparatus include the stepper motor that holds and switches the position of the perfusion pipette, and the control box. As described in Sections 2.2.3 and 4.2.3, the system was optimized to provide rapid solution changes for whole-cell patch clamp experiments by pulling theta (Chapter 2) or triple-barrelled square glass tubing (Chapter 4) obtained from Warner Instruments to a smaller inner diameter (Figure A.2). The smaller inner diameter allows for the use of shorter movements between the barrel positions and decreases the time course of the solution exchange. The method of heating and pulling theta or triple-barrelled glass tubing has previously been described (Jonas, 1995; Hinkle et al., 2003). As described in detail in the excellent review by Jonas (1995), the pipette is not pulled apart in the electrode puller, but, rather, to a narrower diameter before being manually cut apart with a serrated ceramic wafer. It is critical in this cutting step to ensure a clean (rather 194 than jagged) edge as close as perpendicular to the axis of the pipette as possible, as tip irregularities can disrupt the laminar flow of solution (Jonas, 1995).   Figure A.1. The Warner Instruments SF-77B Fast Step Perfusion system. Shown on the left is the stepper motor, which holds the perfusion pipette (made either from theta tubing or triple- barrelled square tubing). The photograph shows the solution manifolds that connect the solution reservoirs to the perfusion pipette. In our experiments, two-pronged manifolds were sometimes used. Shown on the left is the control box for the perfusion system, which allowed for adjustment of the step size (e.g. 200 – 700 µm movement). The control box also served to connect the stepper motor to the acquisition software, through which the position of the perfusion pipette could be controlled via TTL pulses. The position of the pipette could also be switched manually using the dial on the front of the control pulse. Image courtesy of Warner Instruments (http://www.warneronline.com/product_info.cfm?id=27).  195   Figure A.2. Optimization of perfusion pipettes for use in the SF-77B Fast Step system. A, The upper panel shows a schematic of the triple-barrelled square perfusion pipette before heating and pulling. The lower panels depict the cross-sections of triple-barrelled square and theta tubing and show their respective dimensions. B, the triple-barrelled pipettes were pulled to an internal diameter of ≈0.25 mm using a vertical electrode puller. Following heating and pulling, the pipettes were cut into two parts using a ceramic wafer (along the cut line). The bottom panels depict the end of the cut pipette, with the new, smaller openings. C, Schematic showing the orientation of the perfusion pipette relative to a cell during an experiment. The pipette is positioned in the bath such that the cell is exposed to only one of the three test solutions (two in the case of experiments using theta tubing). Lateral steps of the perfusion pipette, triggered from the control box, rapidly change the solution that the cell is exposed to. 196 A.1.2 The custom fast application tool The experiments in Chapter 3 were performed using a custom made FAT to rapidly change the test solution. Figure A.3 shows a schematic of the FAT positioned close to a patch- clamped cell in the bath. Unlike the SF-77B Fast Step system, the FAT did not move to change solutions. Instead, the software-driven TTL pulses opened and closed the valves regulating the flow of solution from the reservoirs, such that only one solution was flowing through the FAT at a time. This method has previously been described by Dittert et al. (1998).  Figure A.3. The custom fast application tool. The FAT was constructed by gluing polyimide- coated fused silica tubes together and allowing them to feed out of a common outlet capillary, which was placed in the bath close to the target cell. The feeder tubes were connected to the solution reservoirs. The flow of solution was regulated by software-driven TTL pulses such that only one solution was flowing through the FAT at a time. 197 A.1.3 Characterizing the time course of the fast solution change. The same method was used to measure the kinetics of solution changes made using either the SF-77B Fast Step system or the custom FAT. While the bath was constantly perfused with control solution, the assembled perfusion pipette or FAT was positioned as close as possible to the patch-clamped cell. At the beginning of the experiment, the cell was exposed to low (0 or 0.5 mM) K +  solution from one barrel of the perfusion pipette or the FAT. Outward K +  current was elicited by a pulse to 0 mV, during which the solution was rapidly and transiently switched from a low K +  solution to a 140 mM K +  solution (Fig. A.4). The time course of the solution exchange was estimated from a fit of the resulting current decay to a single exponential function. For the SF-77B Fast Step system, the time constant of the current decay was typically ≈25 ms, with a latency of ≈25 ms. Thus, exchange of the solution could be expected to be complete in ≈150 ms. For the custom FAT, the time constant and latency for the current decay were typically between 50 – 100 ms.  198  Figure A.4. Characterization of the perfusion kinetics. The grey trace represents a current trace recorded from a cell upon a 1 s depolarization from −80 mV to 0 mV (see voltage protocol at the top of the panel). Using the SF-77B Fast Step system outfitted with a perfusion pipette constructed from theta tubing (see text), the solution that the cell was exposed to was rapidly changed during the test pulse from a 0 mM K +  bathing solution to one with 140 mM K +  (see perfusion protocol at the top of the panel). The kinetics of the solution change are observed as a rapid decrease in the current due to the change in the equilibrium potential for K + . The latency for the exchange, t, was measured by eye, while the time constant of the solution change, τ, was obtained by fitting the current decay to a single exponential function. The experimental protocol described here was used to measure the kinetics of solution changes with both the SF-77B Fast Step and the custom FAT. 

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