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Modulation of Kv1.5 slow inactivation by external cations Kwan, Daniel Cheuk Hang 2006

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MODULATION OF Kvl.5 SLOW INACTIVATION BY EXTERNAL CATIONS by DANIEL CHEUK HANG KWAN B.Sc.(Hon.), The University of  British Columbia, 2000 A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES (Physiology) THE UNIVERSITY OF BRITISH COLUMBIA December 2006 © Daniel Cheuk Hang Kwan, 2006 ABSTRACT Slow inactivation is an intrinsic biophysical property of  voltage-gated potassium (Kv) channels that results in a non-conducting state under physiological conditions. It limits the amount of  current through Kv channels and affects  cellular excitability. However, the molecular basis of slow inactivation is not well understood. In this thesis investigation, the modulation of  slow inactivation in the human Kvl .5 channel by extracellular Zn2+, protons (H+), Ni2+, and other divalent-cations was studied using standard voltage clamp techniques. Zn2+, H+, and Ni2+ accelerated slow inactivation and caused a current inhibition in Kvl.5 expressed in HEK-293 and mouse'M" cells. The current inhibition was hypothesized to result from  the binding of  Zn2+, H+, and Ni2+ to the turret histidine residue (H463) which in turn promoted a slow inactivation process involving the outer pore mouth arginine residue (R487). The current inhibition induced by Zn2+, H+, and Ni2+ was attenuated either by increasing extracellular [K+] or by mutating H463 to glutamine (H463Q) or R487 to valine (R487V). Unitary current analysis revealed H+ and Ni2+ did not change the single channel current at +100 mV or the single channel conductance between 0 and +100 mV, but the number of  blank (null) sweeps recorded with depolarizing pulses lasting up to 1 s was increased. The proportion of null sweeps correlated well with the extent of  inhibition of  macroscopic Kvl .5 current by external H+. A model incorporating two modes of  gating was employed to describe the transitions between the active sweeps (mode A) and the null sweeps (mode U),  and external H+ was proposed to inhibit Kv 1.5 current by promoting mode U  gating. Consistent with this model was the finding  that external K+ antagonized mode U  gating induced by external H+. Channels were observed to switch from mode U  back to mode A during prolonged depolarizations (> 6 s), and the delay in opening (first latency) was correlated with the dwell time in a depolarization-induced slow inactivated state. j Together, the results suggest that Zn2+, H+, and Ni2+ inhibit Kvl .5 current by promoting a slow (P/C-type) inactivation process proceeding from  closed states. TABLE OF CONTENTS ABSTRACT ii TABLE OF CONTENTS iv LIST OF TABLES viii LIST OF FIGURES ix LIST OF ABBREVIATIONS xi ACKNOWLEDGEMENTS xiii DEDICATION . . xiv CO-AUTHORSHIP STATEMENT xv 1. Introduction to slow inactivation and voltage-gated potassium channels 1 1.1 Early Kv channel biophysics . . . 2 1.2 Structure of  Kv channels 6 1.2.1 The pore-forming  domain 9 1.2.1.1 The activation gate 12 1.2.1.2 The selectivity filter  13 1.2.1.3 The outer pore region . 14 1.2.1.4 Permeation and selectivity in K channels 15 1.2.2 Voltage-sensing domain 16 1.2.3 Possible structural model for  Kv channels . . 18 1.2.4 Electro-mechanical coupling in Kv channels 20 1.3 Inactivation in Kv channels 21 1.3.1 N-Type inactivation . 22 1.3.2 U-type inactivation 25 1.3.3 Slow (P/C-type) inactivation .21 1.3.3.1 Role of  the outer pore in slow inactivation 28 1.3.3.2 Role of  the selectivity filter  in slow inactivation 30 1.3.3.3 Role of  S4 in slow inactivation 33 1.4 Kvl .5 and scope of  thesis investigation 36 1.5 References  40 2. Molecular determinants of  the inhibition of  human Kvl.5 potassium currents by external protons and Zn2+ 56 2.1 Introduction 56 2.2-Materials and Methods 59 2.2.1 Cell preparation 59 2.2.2 2.2.3 2.3 Results 2.3.1 2.3.2 2.3.3 K+ 0 relief  of  the effect  of  protons is fitted  by a model of  non-competitive inhibition 68 2.3.4 External Cs+ ions mimic the block-relieving effect  of  K+ 70 2.3.5 Sensitivity to H+ 0 and Zn2+0 inhibition is reduced in MCvl.5 H463Q 71 2.3.6 H+ 0 and Zn2+0 accelerate inactivation 75 2.3.7 Current inhibition by protons and Zn2+0 is reduced in MCvl.5 R487V . . . . 75 2.3.8 Current inhibition by H+ 0 and Zn2+0 is apparently not use-dependent 78 2.3.9 Protons cause a depolarizing shift  of  the Q(V) relationships but do not affect O : . . . 79 2.4 Discussion 83 2.4.1 Evidence against a pore blocking mechanism 85 2.4.2 A connection between current inhibition and an inactivation process . . . . 86 2.4.3 External acidification  and o^-gating charge movement 87 2.4.4 External acidification  and off-gating  charge movement 88 2.4.5 What is the connection between H463 and R487? 89 2.4.6 Inactivation and the influence  of  the charge on and size of  the residue at position 463 91 2.5 References  93 3. The external K+ concentration and mutations in the outer pore mouth affect  the inhibition of  Kvl.5 current by Ni2+ 97 3.1 Introduction 97 3.2 Materials and Methods 100 3.2.1 Cell preparation 100 3.2.2 Recording solutions 101 3.2.3 Signal recording and data analysis 102 3.3 Results 104 3.3.1 Increasing [K+]0 causes a rightward shift  of  the concentration dependence of the Ni2+block 106 3.3.2 The time courses of  the onset and the offset  of  the Ni2+ block are similar ; 107-3.3.3 Ni2+ block is associated with a slight acceleration of  inactivation of  residual currents 109 3.3.4 The K D for  the Ni2+ block is increased in the H463Q and R487V mutants 112 3.3.5 Ni2+ decreases channel availability 112 3.3.6 Ni2+ causes a rightward shift  of  the QON-V  curve but does not affect  QMA X ; 114 Recording solutions 60 Signal recording and data analysis 61 64 Increasing [H+]0 causes a gating shift  and reduces the maximum conductance (g ) 64 Vo  max J Increasing [K+]0 inhibits the reduction of  gmax by extracellular acidification : 65 3.3.7 Co2+ and Cd2+, but not Mn2+, block Kvl.5 116 3.3.8 Co2+ and Zn2+ mimic the effect  of  Ni2+ on Kvl.5 inactivation 118 3.4 Discussion 119 3.5 References  126 4. Single channel analysis reveals different  modes of  Kvl.5 gating behaviour regulated by changes of  external pH 129 4.1 Introduction 129 4.2 Materials and Methods 130 4.2.1 Cell preparation 130 4.2.2 Electrophysiology 131 4.2.3 Data analysis 132 4.3 Results '....• 134 4.3.1 pH affects  channel availability 138 4.3.2 pH does not substantially affect  intraburst behaviour 144 4.4 Discussion 148 4.5 References  154 5. The microscopic changes in Kvl.5 slow inactivation gating caused by external K+ and H+ . 157 5.1 Introduction 157 5.2 Materials and Methods -158 5.2.1 Cell preparation 158 5.2.2 Electrophysiology 159 5.2.3 Data analysis 160 5.3 Results 161 5.3.1 External K+ antagonizes the H+-induced Kvl.5 current inhibition in ltk~  cells 161 5.3.2 External K+ increases the mean P0 but not the unitary current amplitude : 165 5.3.3 External K+ does not change the intraburst behaviour of  Kvl.5 dramatically 168 5.3.4 External K+ promotes mode A gating 171 5.3.5 External K+ slows the onset but not the recovery from  slow inactivation 175 5.3.6 Kvl.5 shows a slow rising phase at low pH with K+-free  solution 178 5.3.7 External H+ promotes.group L (long first  latency) and N  (null) behaviour 181 5.3.8 The mean long first  latency correlates significantly  with the mean long gap 183 5.4 Discussion 188 5.4.1 External K+ and mode U  gating 189 5.4.2 Mode Uor  not mode U1  190 5.4.3 Mode [/gating and slow inactivation 191 5.4.4 Molecular basis for  the slow rising phase 193 5.5 References  197 6. Slow inactivation in Kvl.5 199 6.1 Mechanism for  the external cation-induced current inhibition 199 6.2 Implications of  slow inactivation gating 203 6.2.1 Mode U  gating versus U-type inactivation 204 6.2.2 Slow inactivation from  closed states 205 6.2.3 Possible mechanism for  closed state inactivation 207 6.2.4 Possible interactions between H463 and the outer pore 208 6.2.5 Gating within bursts and depolarization-induced inactivation 210 6.3 Comparison of  H+-induced effects  in Kvl.5 and in other channels 212 6.3.1 External H+ modulation of  Shaker  212 6.3.2 External H+ modulation of  Kvl.4 213 6.3.3 External H+ modulation of  KCNQ2/KCNQ3 214 6.4 Physiological significance  and future  directions ' 215 6.5 References  219 LIST OF TABLES Table 1.1 Comparison of  properties of  N-type, U-type, and P/C-type inactivation 22 Table 6.1 Comparison of  the effects  of  external Zn2+, Ni2+, and H+ in Kvl .5 200 LIST OF FIGURES Figure 1.1 Gating schemes of  Kv channels 4 Figure 1.2 Structural representations of  Kv channels 8 Figure 1.3 Structural representations of  the pore domain 11 Figure 1.4 Stick model of  the hydrophobic network surrounding the selectivity filter  14 Figure 2.1 Extracellular acidification  decreases the maximum conductance (gmax) and causes a rightward shift  of  the conductance-voltage (g(V))  relationship for  Kvl .5 currents 66 Figure 2.2 Increasing K+0 reduces the inhibition of  MCvl .5 current by protons 67 Figure 2.3 The concentration dependence of  the inhibition of  Kvl.5 currents by protons in zero, 5, and 140 mM K+0 69 Figure 2.4 The concentration dependence of  the antagonism by K+0 and Cs+0 of  the inhibition of /zKvl.5 currents by H+ 0 70 Figure 2.5 The structure of  the S5, S6 and the pore (P) loop of  Kvl .5 inferred  from  the crystal structure of  KcsA 72 Figure 2.6 A point mutation in the turret (S5-P loop), H463Q, reduces the inhibition but not the gating shift  caused by H+ 0 and Zn2+0/ 73 Figure 2.7 In hK\'\  .5 H463G slow inactivation is greatly accelerated and the conductance collapses in zero K+0 at pH 7.4 74 Figure 2.8 A mutation near the pore mouth, R487V, substantially reduces the sensitivity to inhibition by H+ 0 and Zn2+0 77 Figure 2.9 The effect  of  the stimulus frequency  and holding potential on the inhibition of  wt hKvl.5 currents by H+ 0 . 79 Figure 2.10 Extracellular acidification  to pH 5.4 causes a depolarizing shift  of  the QON(V)  and QOF/V)  relationships but does not reduce QMA X  81 Figure 3.1 Ni2+ block of  Kvl.5 currents in 0 mM K+0 105 Figure 3.2 Increasing [K+]0 changes the concentration dependence of  the block of  Kv 1.5 by Ni2+ , 108 Figure 3.3 A comparison of  the time course of  the onset and offset  of  the inhibition of  Kvl .5 by Ni2+ and H+ 0 109 Figure 3.4 Inactivation and recovery kinetics of  Kvl .5 at pH 5.4 or with 2 mM Ni2+ I l l Figure 3.5 Ni2+ sensitivity is reduced in Kvl.5 H463Q and Kvl.5 R487V 113 Figure 3.6 Ni2+ effects  at the single channel level 114 Figure 3.7 Gating charge movement with 1 mM Ni2+ in Kvl.5 116 Figure 3.8 Co2+0 also causes a concentration-dependent block of  Kvl .5 currents but is an order of  magnitude less potent than Ni2+ 117 Figure 3.9 Co2+ and Zn2+ mimic the effect  of  Ni2+ on macroscopic inactivation 119 Figure 4.1 The single channel current amplitude of  Kvl.5 does not change between pH 7.4, 6.4, and 5.9 - • 136 Figure 4.2 Open channel current-voltage relationship showing the single channel conductance does not change with pH 138 Figure 4.3 Current behaviour at the macroscopic level and the ensemble average of  unitary current of  Kvl.5 at pH 7.4, 6.4, and 5.9 are qualitatively similar 139 Figure 4.4 Modal gating of  Kvl.5 at different  pHs 140 Figure 4.5 Decreasing external pH to 6.4 decreases the mean burst length and increases the apparent interburst duration 145 Figure 4.6 Extracellular acidification  does not significantly  affect  gating transitions within bursts . . . 146 Figure 5.1 Effects  of  external K+ on the macroscopic current through Kvl .5 expressed in Itk cells at pH 6.4 163 Figure 5.2 Unitary current traces and the mean open probability with 0, 1, 3.5, and 20 mM external K+ at pH 6.4 , . 167 Figure 5.3 External K+ does not alter gating within bursts at pH 6.4 170 Figure 5.4 External K+ causes parallel changes in both the mean long first  latency and the mean long gap at pH 6.4 173 Figure 5.5 External K+ promotes mode A and short gaps between bursts 175 Figure 5.6 At pH 6.4 increasing [K+]0 increases the burst duration 178 Figure 5.7 Kvl .5 current shows a slow rising phase at low pH with 0 mM external K+ 180 Figure 5.8 Unitary current, mean open probability, and the proportion of  sweeps in groups S, L, and N  at different  pHs 182 Figure 5.9 The mean long first  latency correlates with the mean long gap between bursts . . . 184 Figure 5.10 Ensemble of  idealized unitary current traces with long first  latency at pH 6.4 and 5.9 186 Figure 5.11 Mean burst durations of  Kvl.5 at different  external pH with 0 external K+ 187 LIST OF ABBREVIATIONS Amino Acids 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 ' Gin Q Glycine Gly G Histidine His H Isoleucine lie 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 4-AP - 4-aminopyridine y- single channel conductance t - time constants ANOVA - analysis of  variance BK - Ca2+-activated potassium channel with large conductance Cav - voltage-gated calcium (channel) CHO - Chinese hamster ovary (cell) CNG - cyclic nucleotide-gated (channel) EDTA - ethylenediaminetetraacetic acid EGTA - ethylene-glycol-bis(2-aminoethyl ether)-N,N,N'N'-tetraacetic acid EK  - equilibrium potential of  potassium g - macroscopic conductance gmilx - maximum macroscopic conductance i - microscopic current . I - macroscopic current IK u r - ultra-rapid delayed rectifier  current Im a x - maximum current HEK - human embryonic kidney (cell) HEPES - 4-(2-hydroxyethyl)-l-piperazineethanesulfonic  acid HERG - human ether a-go-go-related gene (channel) K N  - dissociation constant Kir - inwardly-rectifying  potassium (channel) Kv - voltage-gated potassium (channel) MEM - minimum essential medium MES - 2-[N-Morpholino]ethanesulfonic  acid MTS - methanethiosulfonate Nav - voltage-gated sodium (channel) nH  - Hili coefficient NMG+ - N-methyl-D-glucamine NMR - nuclear magnetic resonance p - probability P0 - Open probability PxP - proline-x-proline (motif) QA - quaternary ammonium QMA X  - maximum gating charge Qon - ON gating charge QOFF- OFF gating charge r - correlation coefficient s - second SCAM - substituted cysteine accessibility method SD - standard deviation SEM - standard error of  the mean ShakerlR  - (fast)  /nactivation Removed Shaker  (channel) sr - standard error of  r (correlation coefficient) subscript 'i' - intarcellular subscript 'LG' - long gap subscript 'LL' - long first  latency subscript 'o' - extracellular subscript 'SG' - short gap subscript 'SL' - short first  latency TAPS - N-tris[Hydroxymethyl]methyl-3-amine-propanesulfonic  acid TEA+ - tetraethylammonium tcrU  - critical time (burst analysis) TM - transmembrane V 1/ 2 - half-activation  voltage wt - wild-type ACKNOWLEDGEMENTS I would like to express my sincere gratitude to all the people who have contributed their time and effort  to my graduate study experience at UBC. To my graduate supervisor, Dr. Steven J. Kehl, I thank you for  your boundless patience, your support, and all the time you have spent in teaching me what good science is about. I thank my supervisory committee members, Dr. David Fedida, Dr. Eric Accili, and Dr. Edwin Moore, who have been very supportive and have provided helpful  insights into my research projects and into this thesis. It has been a great experience working with other graduate and undergraduate students, and I appreciate all the time we have shared together. I would like to thank all my friends,  especially G.Y., for  all the support and understanding you have given me. Finally, to my wonderful  and loving parents, I am grateful  for your encouragement, guidance, and patience. DEDICATION (^o-  'my, CO-AUTHORSHIP STATEMENT Chapter 2: Kehl, S. J., Eduljee, C., Kwan, D. C., Zhang, S., & Fedida, D. (2002). Molecular determinants of  the inhibition of  human Kvl.5 potassium currents by external protons and Zn2+. J. Physiol. 541:9-24. Daniel Kwan was responsible for  i) performing  the experiments and analysis of  the effects  of  IT on Kvl .5 current at pH 6.4 (Figure 2.1); ii) performing  some of  the experiments and analyses of the K+ attenuation of  the H+-induced current inhibition (Figure 2.2); iii) performing  the experiments and analysis on the R487V mutant; iv) performing  the experiments and analysis of the frequency  dependence of  the H+-induced current inhibition (Figure 2.9); v) preparing and constructing the figures  involved (50% contribution). Cyrus Eduljee performed  the experiments on the H463G, H463Q, and H463Q/R487V mutants. Shetuan Zhang examined the effects  of external H+ on gating current. Steven Kehl conducted the rest of  the experiments and wrote the manuscript. Chapter 3: Kwan, D. C., Eduljee, C., Lee, L., Zhang, S., Fedida, D., & Kehl, S. J. (2004). The external K+ concentration and mutations in the outer pore mouth affect  the inhibition of  Kvl.5 current by Ni2+. Biophys. J. 86:2238-2250. Daniel Kwan was responsible for  i) performing  the experiments and analysis of  the effects  of Ni2+ on Kvl.5 current; ii) performing  the single channel recording and analysis on the effects  of Ni2+ (20% contribution). Cyrus Eduljee performed  the experiments on the H463Q mutants. Shetuan Zhang examined the effects  of  external Ni2+ on gating current. Logan Lee assisted with some of  the experiments. Steven Kehl conducted the rest of  the experiments and wrote the manuscript. Chapter 4: Kwan, D. C., Fedida, D., & Kehl, S. J. (2004). Single channel analysis reveals different  modes of  Kvl.5 gating behaviour regulated by changes of  external pH. Biophys. J. 90:1212-1222. Daniel Kwan was responsible for  i) performing  all the whole-cell experiments and analyses of the effects  of  H+ on Kvl.5; ii) performing  all the single channel recording and analyses; iii) prepare all the figures  in the manuscript; iv) designing all the experiment procedures for  both the single channel and whole cell recordings; v) writing the manuscript except the discussion (85%) contribution). Steven Kehl wrote the discussion for  the manuscript and assisted in editing the rest of  the manuscript. 1. Introduction to slow inactivation and voltage-gated potassium channels Ion channels are indispensable molecular components of  all living cells, from  unicellular prokaryotic bacteria to multicellular eukaryotic organisms. These integral membrane proteins facilitate  and regulate the movement of  ions across the plasma membrane, which is an impermeable barrier for  all charged particles. In non-excitable cells, potassium (K+ or K)-selective channels are involved in basic physiological functions  such as maintaining K+ and electrolyte balance (Hille, 2001). In excitable cells, the flow  of  ions across the plasma membrane gives rise to bioelectric phenomena (Hodgkin and Huxley, 1952), and K channels, in particular the voltage-gated K (Kv) channels, are involved in repolarizing the membrane voltage from  an excited state back to the resting level and in reducing membrane excitability (Hille, 2001). Surprisingly, more than 65 different  genes in the human genome have been identified  to encode various Kv channels (Alexander et ah, 2006). This large repertoire of  Kv channels highlights their important role in fine-tuning  the repolarization step in an action potential and in other functions.  In addition, mutations in various Kv channels can lead to inherited neurologic and cardiac disorders, such as episodic ataxia, epilepsy, long QT syndrome, and sudden cardiac death (Ashcroft,  2000). Therefore,  a detailed understanding of  these proteins, both in terms of  their biophysical properties and pharmacological profiles,  is of  critical importance. In this thesis, the modulation of  slow inactivation in Kvl.5 by external cations is examined, and the purpose of  this chapter is to review some of  the basic biophysical properties of K channels and to lay out the background material for  the reader to understand the results given in the subsequent chapters. An overview of  K channel biophysics will first  be introduced, including the Hodgkin and Huxley formalism  and some early seminal experiments leading to the elucidation of  channel function  and structure. Second, the molecular determinants for  various K channel properties based on a number of  K channel crystal structures (permeation, activation, electro-mechanical coupling, etc.) will be described. Next, the different  types of  inactivation will be described with the focus  on the so-called slow (P/C-type) inactivation. Finally, the scope of this research project will be defined. 1.1 Early Kv channel biophysics There are two main aspects of  Kv channel function:  gating and permeation. The term gating is used to describe how a channel responds to stimuli. For example, Kv channels can populate different  functional  states in response to the membrane voltage, and the transitions between these functional  states are collectively called the gating of  the channel. The two basic functional  states are the closed (non-conducting) state and the open (conducting) state. For most of  the cases, Kv channels are assumed to predominantly reside in a closed state at rest. The probability of  being open increases with depolarization. The gating process by which a channel makes transitions from  a closed state to the open state is called activation, and the reverse process is called deactivation. Gating of  channels can be described using gating models (schemes) in which the transitions between states are governed by voltage-dependent rate constants. Gating models can also be used for  simulating the kinetic, behaviour of  channels to test hypotheses. Probably the best known gating model for  K current was proposed by Hodgkin and Huxley (1952) for  the delayed rectifier  Kv channel of  the squid giant axon. In their model, four closed states and one open state were linked in series as shown in Figure 1.1 A. Each transition was assigned a voltage-dependent rate constant. The forward  rate constants (toward the open state) became larger with depolarization, whereas the backward rate constants had the opposite voltage dependence. To account for  the delay in current onset and the sigmoidal kinetics of activation, the Kv channel was proposed to start primarily at CI  at rest {i.e.,  at negative holding potentials) and, upon depolarization, progressed through the other three closed states {C2  to C4) before  finally  reaching the open state {O). To account for  this behaviour, Hodgkin & Huxley (1952) envisioned that a Kv channel contained four  "particles," which could be in either a non-permissive or in a permissive state, and that once all four  "particles" were in the permissive state the channel opened. Conversely, as soon as a single "particle" became non-permissive, the channel reverted to a closed, non-conducting state. This model was consistent with the decay of ionic current upon repolarization being well-fit  by a single exponential function.  In the wake of the the Hodgkin-Huxley model, more sophisticated and complex models for  other Kv channels have been developed (Hoshi et al., 1994; Schoppa and Sigworth, 1998a; Schoppa and Sigworth, 1998b; Schoppa and Sigworth, 1998c; Zagotta et al., 1994b; Zagotta et al., 1994a). One common modification  is the inclusion of  another closed state {C5-  in Figure 1.1 B) to account for a concerted opening step (Ledwell and Aldrich, 1999; Pathak et al., 2005; Smith-Maxwell et al., 1998). In this state, all the gating particles are activated (in the permissive state), but the channel is not yet open. In their seminal 1952 paper, Hodgkin & Huxley also proposed that the voltage-dependent steps resulted from  charged elements moving within the membrane voltage field  (Hodgkin and Huxley, 1952), an hypothesis that accurately predicted the existence of  "gating currents" in voltage-gated channels. However, it was about twenty years before  these hypothetical gating currents could be measured. The first  recording of  gating current was made by Armstrong & Bezanilla (1974) from  voltage-gated sodium (Nav) channels in a preparation where all the A • P(V) 2P(V) 3p(V) 4P(V) C t — » C2 — „ C3 * » C4  "  »T O 4a(V) 3a(V) 2a(V) a(V) P(V) 2p(V) 3p(V) _4P(V) 5(V) C1  " ^ C2 * » C3  "  C5— • O 4a(V) 3a(V) 2a(V) a(V) y(V) Figure 1.1 Gating schemes of  Kv channels. A. A gating scheme for  generic Kv channels based on the Hodgkin-Huxley model (Hodgkin and Huxley, 1952). A channel can either be in one of  the closed, non-conducting states (CI  to C4) or in the open state(O). Transitions between states are all voltage-dependent. B. In more recent models of  activation, another closed state (C5) is included before  the open state to account for  the concerted opening step observed. In C5, all the gating particles are assumed to have activated. No inactivated state is included. permeant ions were removed. Less than a decade later, Bezanilla et al. (1982) reported the first gating current from  the Kv channel of  the squid giant axon. The gating currents recorded in both studies were very small compared to the ionic current and were capacitive (transient) in nature. Integrating the gating currents with respect to time gave an estimate of  the amount of  gating charges mobilized. The studies on gating currents have provided important information regarding transitions between closed states that are otherwise electrically silent. In the absence of  structural information  in the early days, the activated voltage sensor was hypothesized to open a "gate" in the Kv channel that otherwise prevented the flow  of  K+. Interestingly, internally applied quaternary ammonium compounds, such as tetraethylammonium (TEA+), were found  to block the Kv channel in squid giant axon and in the frog  node of  Ranvier only when the activation gate was open (Armstrong, 1966; Armstrong and Hille, 1972). In addition, bound TEA+ and other small quaternary ammonium compounds could become trapped in the Kv channel if  the activation gate was closed by repolarization, which suggested that the activation gate acted as a physical barrier and was located between the cytoplasmic face  of  the channel and the receptor site for  quaternary amines (Armstrong, 1966; Holmgren et al., 1997). These data provided additional insights into Kv channel structure, and, as we shall see later, these structural interpretations were elegantly confirmed  in the crystal structures of  several K channels. Once the activation gate is open, K+ ions start to move down their electrochemical gradient, a process known as permeation. Hodgkin & Keynes (1955) were the first  to propose that the Kv channel of  the cuttlefish  giant axon was a multi-ion pore, meaning that multiple ions occupied the conducting pathway during a single-file  permeation process. However, at that time the structural basis for  the high selectivity of  Kv channels for  K+ over Na+ was unknown. Under normal physiological conditions, the selectivity sequence for  most Kv channels is Tl+ > K+ > Rb+ > NH4+ > Cs+ » Na+ = Li+ (Blatz and Magleby, 1984; Heginbotham and MacKinnon, 1993; Hille, 1973; Latorre and Miller, 1983; Shapiro and DeCoursey, 1991), and selectivity in K channels is clearly not based solely on the ionic crystal radius since that of  Na+ is less than that of K+. In a series of  studies, Mullins (1959) proposed that the selectivity of  Kv channels was based on the energy required for  dehydration (reviewed in Hille, 2001). Roughly a decade later, Bezanilla & Armstrong (1972) proposed that the Kv channel pore contained "coordinating cages" that replaced the hydration shell for  K+, in effect  acting as surrogate water molecules. It took another 25 years later before  the chemical basis for  these coordinating cages was revealed in the very first  crystal structure of  a K channel (Doyle et al., 1998). With a sustained depolarizing pulse, most ion channels do not stay in the open state(s) indefinitely  but instead undergo a process called inactivation that results in a non-conducting state that is distinct from  the closed states described above. Hodgkin & Huxley (1952) proposed that the transient Na+ conductance of  the squid giant axon was due to the closing of  a separate, inactivation gate which functioned  independently from  the activation gate. This view of inactivation gating was adopted for  the transient K currents (Connor and Stevens, 1971). A number of  different  types of  inactivation, with distinct molecular mechanisms, have been discovered. Inactivation is discussed in greater detail later in this chapter. To summarize, a number of  ingenious experiments between the 1950's and 1980's provided functional  data from  which structural insights for  ion channels were drawn, but the overall structure of  Kv channels was just starting to emerge. It became clear that the different functional  states of  Kv channels arose from  different  conformations,  and starting in the 1980's, a growing number of  studies began to focus  mainly on the structural aspect of  ion channels (Noda et al., 1984; Papazian et al., 1987; Steinbach, 1989; Tanabe et al.,1987) as the ion channel field entered the era of  molecular cloning. Various ion channels were cloned in the last twenty years, and a movement towards studying cloned channels in heterogenous systems was evident. The cloning and structural studies on ion channels have revolutionized the field  of  channel biophysics by providing a physical framework  for  the functional  data. 1.2 Structure of  Kv channels In 1987, the Jan group published the primary sequence of  the first  Kv channel, the Shaker channel, from  the fruit  fly  Drosophila melanogaster  (Papazian et al., 1987). At that time, several voltage-gated sodium (Nav) and calcium (Cav) channels had already been cloned (Noda et al., 1984; Tanabe et al., 1987); therefore,  comparisons were quickly drawn between the cloned voltage-gated channels. A major difference  was that the size of  Shaker  was about a quarter of  the cloned Nav and Cav channels. The cloned Shaker  gene was found  to encode a channel-forming a-subunit of  a Kv channel, which was proposed and later confirmed  to contain six transmembrane helices with intracellular N- and C-termini, as shown in Figure 1.2 A (Doyle et al., 1998; Jiang et al., 2003; Long et al., 2005a). It was shown that a Kv a-subunit is analogous to a single domain of  a Nav or Cav channel, and a Kv channel was proposed to be a homotetrameric construct with a four-fold  rotational symmetry, as shown in Figure 1.2 B and C (Doyle et al, 1998; Jiang et al., 2003a; Long et al., 2005a; MacKinnon, 1991). Following the cloning of  the Shaker  channel, three other homologues (Shab, Shaw,  and Shal)  appeared in relatively rapid succession (Butler et al., 1989) and were also confirmed  to encode functional  Kv channels (Wei et al., 1990). The topology of  all the cloned Kv channels was later found  to be conserved; therefore,  they were grouped into the so-called 6TM-1P channel family  (Alexander et al., 2006; Hille, 2001). As the list of  cloned K channels grew, the need for  a systematic nomenclature for  all K channels became clear. Originally, channels were named based on the species and tissue of origin. However, without knowing the corresponding gene, a channel might, and often  did, acquire more than one name. In 1991, a naming system based on homology and evolutionary relationship between the different  K channels was proposed and implemented shortly afterward (Chandy, 1991). Each Kv channel protein is now identified  as Kvx.y in which x and y are numbers representing a specific  subfamily  and the order of  discovery in that particular subfamily, respectively (Hille, 2001). In addition, the Kv channel genes are labelled as KCNMV in which M is a letter assigned to a specific  subgroup, and N  is a number assigned to the particular channel. The four  Drosophila  Kv channels (Shaker,  Shab, Shaw,  and Shal)  are now known as the prototypes of  the Kvl to Kv4 subfamily  members, and their gene families  have been named as KCNA to KCND, respectively. By comparing their primary sequences, Kv channels are seen to differ  mainly in the 100A 10QA Figure 1.2 Structural representations of  Kv channels. A. Schematic drawing of  the pore-forming  a-subunit of  Kv channels. B. Side view of  the ribbon representation of  the Kvl.2 crystal structure showing the transmembrane domains (TM), the T1 domain (Tl), and an associated P-subunit (P) (Long et al., 2005a). Each of  the 4 subunits is represented by a different  colour. C. Stereo view of  the Kvl.2 crystal structure from  the extracellular side directly above the pore (Long et al., 2005a). The transmembrane segments for  the subunit in red are labelled from  SI to S6. The pore domain (S5 and S6) of  one subunit is adjacent to voltage-sensing domain from another subunit. composition of  the intracellularly-located N- and C-termini, whereas the transmembrane segments and their linkers are, in the main, very similar. Likewise, the different  members within the same subfamily  often  have subtly different  biophysical and pharmacological properties that arise from  the slight differences  in their primary sequences. Based on structural similarities, a number of  functional  domains have been identified  in all Kv channels. For example, the Tl domain in the cytoplasmic N-terminus of  each a-subunit is responsible for  tetramerization, in which only subunits from  the same subfamily  can form  homo- or heterotetramers (Bixby et al, 1999; Pfaffinger  and DeRubeis, 1995; Shen et al, 1993; Shen and Pfaffinger,  1995; Xu et al, 1995; Zerangue et al, 2000). The Tl domains are thought to associate with each other during protein translation before  individual subunits are folded  properly (Robinson and Deutsch, 2005; Strang et al, 2001). Tl domains also serve as the docking sites for  the N-type inactivation (see below) particle and auxiliary P-subunits (Gulbis et al, 2000). The P-subunits, which are shown with the Kvl.2 crystal structure in Figure 1.2 B (Long et al, 2005a), can modulate channel function  and also act as chaperone proteins to increase the. surface  expression of  Kv channels (Wible et al, 1998). The N- and C-termini may contain additional motifs  and/or domains which affect  channel function  (Jerng and Covarrubias, 1997; Ju et al, 2003; Nishida and MacKinnon, 2002; Wray, 2004) but lie outside the focus  of  this study. In addition to the Tl domain, a Kv channel has two other functional  domains: the pore-forming  (pore) domain and the voltage-sensing domain. The pore domain constitutes S5, the P-loop, S6, and the associated linkers; the voltage-sensing domain is formed  by SI, S2, S3, S4 and the associated linkers, with S4 being the main voltage sensor. 1.2.1 The pore-forming  domain The pore domain is a basic functional  unit of  K channels. Even though K channels from different  families  may have substantial differences  in their primary sequences that give rise to their specific  conduction and gating properties (De Biasi et al, 1993; Harris and Isacoff,  1996; Kirsch et al, 1992), the general pore architecture and basic permeation process are thought to be very similar among K channels (Doyle et al, 1998; Heginbotham et al, 1992; MacKinnon el al, 1998; MacKinnon and Doyle, 1997). Figure 1.3 A shows an alignment of  the outer pore regions (from  the end of  S5 to the start of  S6) from  a number of  K channels including Shaker.  From the alignment, it is clear that the two linkers (S5-P and P-S6) are the least conserved, whereas the P-loop is highly conserved within a subfamily  and differs  only slightly between subfamilies.  The P-loop contains the signature sequence TxxTxGY/FGD which forms  the selectivity filter  (Doyle et al., 1998; Heginbotham et al., 1992; Heginbotham et al., 1994). Much of  the theoretical work on permeation and selectivity has used the KcsA channel pore as a structural model. The KcsA channel is an intracellular-proton activated K channel from  the bacterium o Streptomyces  lividans,  and it was the first  K channel for  which high resolution (2-3 A) crystal structures were obtained (Doyle et al., 1998; Zhou et al., 2001b). The sequence of  the pore for KcsA is included in Figure 1.3 A for  comparison. KcsA is homologous to the pore domain of  Kv channels or the inward rectifier  K (Kir) channel in the 2TM-1P family  (Caprini et al., 2005; Lu et al., 2001). Indeed, KcsA can be substituted into the pore domain of  Shaker  to form  a functional Kv channel (Caprini et al., 2005; Lu et al., 2001; Lu et al., 2002). The a-subunit of  KcsA consists of  two transmembrane helices: the outer helix, which is homologous to S5 in Kv channels or Ml in Kir, and the inner helix, which is homologous to S6 in Kv and M2 in Kir channels. These helices line the hydrophilic pore and form  the interface  between the pore lumen and the hydrophobic lipid environment of  the membrane (Aiyar et al., 1994; Lu and Miller, 1995; Shieh and Kirsch, 1994; Yellen et al., 1991). In addition, the KcsA structure reveals two constrictions along the central axis of  the pore: the cytoplasmic bundle crossing formed  by the convergence of  the inner helices and the selectivity filter  formed  by the signature sequence (Figure 1.3 5 grey-blue sticks). The bundle crossing is thought to be the activation gate for  K channels, whereas the selectivity filter  is a major regulatory site for  permeation and selectivity as discussed above. A Shaker 418 hKvl . 1 348 rKvl .2 350 hKvl . 3 368 rKvl .4 502 hKvl .5 456 IS5-P l i n k e r | P - l o o p I P-S6 I EAGSENS FFKSIPDAFWWAVVTMTTVGYGDMTPVGVWGKIVGS —EEAE-H-S Y—TIG —DERD-Q-P V-TTIG — DDPT-G-S H—TIG —DEPTTH-Q K-ITVG — DNQGTH-S R-ITVG rKv2 . 1 349 —KDEDDTK-K AS Tl IY-K-LL G KcsA 51 -RGAPGAQLITY-R-L—S-E-A LY—TL—RL-AV Cavity Figure 1.3 Structural representations of  the pore domain. A. Sequence alignments of  the outer pore region from  the indicated K channel compared to that of  Shaker.  The letters represent the standard single letter code for  each amino acid. The amino acid sequence of  Shaker  is listed in full,  while the dash (-) in the sequence of  other channels represents a residue identical to that in Shaker.  The number in front  of  each sequence show the position of  a highly conserved glutamate (E) residue. B. Side view of  ribbon representation of  the KcsA crystal structure with the front and back subunits removed for  clarity (Roux, 2005). The S5 helices (green), pore helices (orange), and S6 helices (blue) are colour coded, whereas the turret residues are shown as a continuous grey line. The selectivity filter  is shown as grey-blue sticks with the carbonyl oxygen atoms shown in red. The green spheres represent K+ at the coordination sites as labelled. C. Electron density map of  the selectivity filter  (Zhou et al., 2001b). The green spheres represent K+ ions, whereas the red dots represent water molecules. The selectivity filter  orientation is similar to that shown in B. Seven coordination sites are shown, with S, and S4 labelled. K ions are shown to occupy the two external binding sites (S0 and Sext) external to S,, with the K+ ion occupying Se x t being surrounded by 4 water molecule. The K+ ion at the central cavity site is surrounded by 8 water molecules, i.e., fully  hydrated. 1.2.1.1 The  activation gate As noted.previously, access of  K+ to the conducting pathway is regulated by the activation gate. From the KcsA structure, the location of  the bundle crossing on the cytoplasmic side of  the channel, and the large central cavity behind it, agrees well with the description for  the activation gate proposed previously (Armstrong, 1966; Holmgren et al., 1997). As shown in Figure 1.3 B, this bundle crossing arises from  the convergence of  the inner helices (blue) to form  an "inverted teepee." The dimensions of  the bundle crossing are such that the passage of  K+ is prevented; therefore,  the KcsA structure is considered to represent the closed conformation  of  K channels (Doyle et al., 1998). On the other hand, in the crystal structure from  MthK, a prokaryotic Ca2+-activated K channel isolated from  the bacterium Methanobacterium  thermoautotrophicum,  the inner helices were shown to be displaced laterally from  the central pore thus creating an opening sufficient  for  K+ to gain access to the central cavity and the selectivity filter  (Jiang et al., 2002a; Jiang et al., 2002b). From a comparison of  the two structures, a conserved glycine residue in the inner helix was proposed to act as a hinge point in opening of  the activation gate (Ding et al., 2005; Jiang et al., 2002b; Magidovich and Yifrach,  2004). In eukaryotic K channels, a proline-x-proline (PxP) motif  is located below this conserved glycine residue, and this motif  is seen in the Kvl .2 structure to form  a kink in S6 and constitutes part of  the activation gate (del Camino et al., 2000; del Camino et al., 2005; del Camino and Yellen, 2001; Long et al., 2005a; Long et al., 2005b; Webster et al., 2004). Between the activation gate and the selectivity filter,  an internal cavity is clearly evident both in the KcsA and the Kvl .2 structure. The size of  this central cavity permits the accommodation of  quaternary amines such as TEA+. In addition, the inner cavity also contains the binding sites for  the N-terminal inactivation peptides and other compounds (Armstrong and Loboda, 2001; Chen and Fedida, 1998; Choi et al., 1993; Fedida, 1997; Zhou et al., 2001a). 1.2.1.2 The  selectivity  filter External to the bundle crossing, the selectivity filter  forms  a functional  unit which, as its name implies, selects K+ over Na+ and other physiological cations during the permeation process. The selectivity filter  is formed  by the K channel signature sequence located in the C-terminal half of  the P-loop (Heginbotham et al., 1994; Heginbotham and MacKinnon, 1993), whereas the N-terminal half  of  the P-loop forms  the pore helix, which has its long axis pointing towards the central cavity (Doyle et al., 1998). The pore helix was originally proposed to have a dipole that could stabilize the ion in the central cavity (Doyle et al., 1998). However, this dipole of  the pore helix has been shown not to contribute significantly  to cation binding in the central cavity, at least in Kir channels (Chatelain et al., 2005). Residues in the pore helix form  an extensive y hydrophobic and hydrogen bond network with the side chains of  the valine and tyrosine residues in the V-G-Y-G sequence of  the selectivity filter,  as shown in Figure 1.4 (Doyle et al., 1998). The V-G-Y-G sequence is folded  such that the side chains are all facing  the pore helix while the carbonyl oxygen atoms are pointing into the conduction pathway, an arrangement that agrees well with previous models for  ion selection in K channels (Bezanilla and Armstrong, 1972; Hille, 1973; Mullins, 1959). The hydrophobic network surrounding the selectivity filter  is thought to act like a spring that holds the selectivity filter  open. In addition, the glutamate residue E71 at the internal end of  the pore helix has been proposed to interact with the aspartate residue in the G-Y-G-D sequence (Cordero-Morales et al., 2006b), and these interactions may. underlie a gating function  of  the selectivity filter  (Berneche and Roux, 2005; Cordero-Morales et al, 2006a; Cordero-Morales et al., 2006b). The interaction between the selectivity filter  and permeant ions, and the role of  the selectivity filter  in inactivation gating, is discussed in more detail later in this chapter. W68 i amm 11 m . &Bm W67 gSS, I SB."' Figure 1.4 Stick model of  the hydrophobic network surrounding the selectivity filter. The figure  is oriented parallel to the plasma membrane with the "pore" at the centre. Residues are as labelled. Y78 is the tyrosine residue in the GYGD signature sequence. Hydrogen bonds are shown as blue sticks. [Doyle et al. (1998); reprint with permission.] 1.2.1.3 The  outer pore region The outer pore region, which is formed  by the S5-P and P-S6 linkers, is the least conserved region of  the pore domain in the Kv channel family  (Figure 1.3 A). Within this region, however, a glutamate residue at the start of  the S5-P linker (E418 in Shaker)  is absolutely conserved among all Kv channels, but its functional  role in normal gating is not well understood (Larsson and Elinder, 2000; Ortega-Saenz et al., 2000). Mutating this residue to an alanine (E418A) or a cysteine (E418C) accelerates slow inactivation (see below) (Larsson and Elinder, 2000; Yifrach  and MacKinnon, 2002), whereas the E418D mutation slows the rate of inactivation (Ortega-Saenz et al., 2000). In both the KcsA and Kvl.2 crystal structures, the S5-P linker is folded  as a random coil at the outer most part of  the channel and forms  the so-called turret region (Doyle et al., 1998; Long et al., 2005a). The turret region varies in length, with the human ether-a-go-go  gene-related channel (HERG) having one of  the longest (38 residues compared to 12 residues in Shaker).  The turret regions of  Shaker  and HERG have been proposed to form  an ordered helical structure (Clarke et al, 2006; Elinder and Arhem, 1999; Jiang et al., 2005; Torres et al., 2003). The role of  the turret in the regulation of  channel function  is less well known, but mutations in this region have been shown to affect  the gating of  Kv channels (Steidl and Yool, 1999). On the other end of  the P-loop, the P-S6 linker is relatively short but functionally important. The residue T449 in Shaker,  or the homologous residue in other Kv channels, has been shown to affect  the kinetics of  slow inactivation (Lopez-Barneo et al, 1993; Ogielska et al, 1995; Schlief  et al, 1996) as well as the binding of  TEA+ in Kv channels (Heginbotham and MacKinnon, 1992; MacKinnon and Yellen, 1990). However, the molecular mechanism by which slow inactivation is modulated by this site is still largely unknown. 1.2.1.4 Permeation and  selectivity  in K  channels The main determinant of  permeation and selectivity in K channels is the selectivity filter. Based on the high resolution KcsA structure, there are up to seven K+ binding sites along the conducting pathway as shown in Figure 1.3 C (Roux, 2005; Zhou et al, 2001b). At the most external site (Sext), K+ is proposed to be fully  hydrated, whereas at the adjacent binding site (S0), the K+ ion is seen to be partially hydrated by three to four  water molecules with the rest of  the hydration shell filled  by the carbonyl oxygen from  each of  the four  glycine residues (G79) in the G-Y-G-D sequence (Berneche and Roux, 2001; Zhou et al, 2001b). Based on electrostatic calculations, occupancy of  Se x t and S0 is mutually exclusive. In the selectivity filter,  a fully dehydrated K+ may bind to any of  the four  different  sites (S, to S4) coordinated by the backbone carbonyl oxygen of  Y78-G77 (S,), G77-V76 (S2), V76-T75 (S3) or T75-V74 (S4) (Berneche and Roux, 2001), but at any given time, only 2 K+ ions may occupy the selectivity filter  (Aqvist and Luzhkov, 2000; Berneche and Roux, 2001; Morais-Cabral et al, 2001; Zhou et al, 2001b). The two K' ions may either be in the S|-S3 or S2-S4 configuration,  and the other two sites are occupied by water molecules. A fully  hydrated K+ can bind to the central cavity site positioned between S4 and the bundle crossing; however, other ions (Na+, Tl\ Rb+, and Cs+) may have to lose part of their hydration shell before  binding to this site, which may contribute to some degree of selectivity (Berneche and Roux, 2001; Zhou et al, 2001b). In addition, S2 shows differential affinity  towards different  ions, and this selectivity for  permeant ions over non-permeant ions may underlie the different  permeability ratio for  the permeant ions (Aqvist and Luzhkov, 2000; Berneche and Roux, 2001; Noskov et al., 2004; Zhou and MacKinnon, 2004). These results are consistent with a model in which selectivity in a Kv channel is based on competitive binding between ions (Immke and Korn, 2000; Kiss et al, 1998; Korn and Ikeda, 1995). Furthermore, K+ and other permeant cations binding to these sites may modulate channel function  and gating, especially in slow inactivation of  Kv channels. In summary, the pore domain is critical for  all K channels as it forms  a highly selective conduit for  K+ movement across the plasma membrane. However, permeation of  ions does not start until the activation gate is opened, a process that is coupled to the activation of  the voltage sensor. 1.2.2 Voltage-sensing domain In Kv channels, although SI, S2, and S3 contribute to voltage-sensing, the S4 segment is considered to be the main voltage sensor. Each S4 segment contains basic residues (arginine (R)) or lysine (K)) at every third or fourth  position (reviewed in Hille, 2001; Wei et al., 1990). The total number of  equivalent gating charges per channel has been estimated to be 13 e0 for Shaker  (Noceti et al., 1996; Schoppa et al., 1992); that is, each S4 contributes between 3-4 gating charges during voltage-sensing. This is consistent with the finding  that in Shaker,  neutralizing each of  the four  outer most arginine residues (R362, R365, R368, R371) decreases the total number of  gating charges in the mutant channel by 4 (Aggarvval and MacKinnon, 1996; Seoh et al., 1996). Using the substituted cysteine accessibility method (SCAM), the four  outermost arginine residues were shown to be accessible to extracellular methanethiosulfonate  (MTS) reagents at depolarization, but only the second to fourth  arginine residues were accessible to intracellular MTS reagents at hyperpolarization (Larsson et al, 1996). These results strongly support the role of  these four  residues as the primary gating charges in Shaker  that move across the voltage field  during voltage-sensing. Besides the basic arginine residues, some acidic residues in S2 (Shaker  E283, E293) and S3 (Shaker  D316) are also involved in voltage sensing (Aggarwal and MacKinnon, 1996; Seoh et al., 1996), and they may form  salt bridges with the gating charges in S4 and stabilize them (Papazian et al., 1995; Sato et al., 2003). In Nav and Cav channels, the S4 in each of  the homologous domains also contains positive charges (Noceti et al., 1996; Yang et al., 1996; Yang and Horn, 1995), however, the four  S4 segments may contribute differently  in voltage-sensing and other processes (Cha et al., 1999a; Chanda et al, 2004; Kuhn and Greeff,  2003; Lam et al., 2005). Even though S4 has generally been accepted as the major carrier of  the gating charges, the precise molecular motion of  S4 is still not fully  understood. A number of  models have been proposed over the years (reviewed by Hille (2001) and Horn (2004)), and two crystal structures of  Kv channels, the KvAP channel from  the bacterium Aeropyrum pernix and the mammalian (r)Kvl.2 from  rat brain, have been solved in the search for  possible mechanisms for  voltage sensing (Jiang et al., 2003; Long et al., 2005a). The two crystal structures show different orientations of  some of  the transmembrane helices, and from  the KvAP structure, an unconventional view described as the voltage-paddle model was proposed. However, the voltage-paddle model was found  to be inconsistent with a number of  functional  studies that preceded and followed  its publication (Gandhi et al., 2003; Gonzalez et al., 2005; see Laine et al., 2004). On the other hand, the Kvl .2 structure agreed with most of  the predictions deduced from  the various functional  studies. Based on the Kvl.2 crystal structure, a possible model for voltage sensing is described below. 1.2.3 Possible structural model for  Kv channels The crystal structure of  Kvl .2 may be the best representation of  native eukaryotic Kv channels to date. The six transmembrane segments are oriented more-or-less perpendicular to the plane of  the membrane, with the S1-S2 linker, S3-S4 linker, the S5-P linker, and P-S6 linker being extracellular. This is consistent with the S1-S2 linker being glycosylated (Gandhi et al., 2003; Shen et al:, 1993). However, S4 is seen in the crystal structure to be partially exposed to the lipid environment without being completely protected by SI, S2, and S3 (Long et al., 2005a), which differs  from  the conventional view in which the hydrophilic S4 segment is thought to be shielded completely from  the lipid environment by SI, S2, and S3 to minimize the thermodynamically unfavourable  charge-lipid interaction. A low resolution KvAP model obtained from  electron microscopy in a more native state also suggested that S4 was exposed t.o the lipid environment (Jiang et al., 2004). The counter-charges in SI, S2, and S3 may be sufficient  to stabilize the gating charge within the lipid core. Given the above structural model, how does S4 move during voltage sensing? Unfortunately,  the Kvl .2 crystal structure did not give much insight into the movement of  S4 with respect to the membrane. However, a number of  functional  studies have suggested S4 is surrounded by a small canaliculus, and that the voltage field  drops across a very small region around S4 (Baker et al., 1998; Goldstein, 1996; Larsson et al., 1996; Starace and Bezanilla, 2004). For example, when the outermost arginine residue in S4 of  Shaker  was mutated to histidine (R362H), a proton current could be recorded upon hyperpolarization in the presence of a proton gradient. It appeared that the imidazole ring of  the histidine residue could bind a proton on the more acidic side and transfer  it to the more basic side with very small movement (Starace and Bezanilla, 2004). The simplest explanation for  this result is that the external and cytoplasmic solutions are separated by a very small space, and that the electric field  is focussed onto a small region. In most models, S4 moves as a rigid structure. Upon depolarization, the electrostatic force  drives the gating charges outward. However, the movement of  S4 is probably more a rotational than an outward (translational) movement (Cha et al., 1999b). S4 may also become tilted, and both the rotational and tilting motion of  S4 may occur over a small region as the four outermost gating charges are shuttled to the external milieu (Ahern and Horn, 2005; Cha et al., 1999b; Glauner et al., 1999). This model is proposed on the basis of  a fluorescence  study which shows limited S4 movement upon depolarization (Chanda et al., 2005). In addition, the structural requirements for  this model are consistent with the Kvl.2 crystal structure (Long et al., 2005a). However, the actual "canaliculi" around S4 have not yet been mapped onto the Kvl .2 crystal structure, and the relative movements of  the different  transmembrane segments has also not been measured. The above model can account for  most functional  data available and seems to be the most probable model thus far. 1.2.4 Electro-mechanical coupling in Kv channels One of  the most interesting questions in studying Kv channel activation is how the conformational  changes in S4 lead to the opening of  the activation gate formed  by S6. As discussed above, the opening of  the activation gate appeared to involve only the inner helices (S6) with the positions of  both the selectivity filter  and the outer helices (S5) remaining essentially unchanged. Opening of  the activation gate involved a rotation of  S6 at a conserved glycine (Ding etal.,  2005; Lee, 2005; Magidovich and Yifrach,  2004; Seebohm et al., 2005; Wang et al., 2005; Zhao et al., 2004) or at the PxP motif  (Bright et al, 2002; Labro et al, 2003; Rich et al, 2002; Webster et al, 2004) which may act as a hinge. But the question remains: how is the movement of  S4 coupled to the conformational  changes of  S6? This question has been partly answered by studies of  a chimeric channel. As mentioned, the pore domains from  different  K channels are structurally very similar; therefore,  it is possible to "swap" the pore domains between two different  K channels. Moreover, the machinery necessary for  the coupling between the voltage sensor and the pore domain may be revealed by progressively substituting the pore domain of  a non-voltage-gated K channel into that of  a Kv channel. Such an experiment has been performed,  and when the KcsA pore domain was substituted into Shaker,  two phenylalanine residues (F401 and F404) in the S4-S5 linker were required to confer  voltage-dependent activation (Caprini et al, 2005; Lu et al, 2002). This intimated that the conformational  changes in S4 were coupled to the movement of  the S4-S5 linker. That is, S4 may be indirectly coupled to S6 via the S4-S5 linker. The coupling between the S4-S5 linker and S6 is further  suggested by the Kvl.2 crystal structure (Long et al, 2005a). In the crystal structure, the pore domain of  each subunit has its S4-S.5 linker associated with its own C-terminal half  of  S6 below the PxP motif,  as shown in Figure 1.2 C (Long et al., 2005b). It is proposed that an "outward" movement of  S4 during depolarization would pull on S6 via the S4-S5 linker, and that this pulling motion results in a rotation, either at the conserved glycine residue or at the PxP motif,  that opens the intracellular gate (Long et al., 2005b). This simple model seems to have merit for  eukaryotic Kv channels; however, it is not clear whether a similar interaction exists in other voltage-gated channels. For example, the human Nav1.5 does not have the PxP motif  in any of  the S6 segments,'"but it does contain a glycine residue in each of  the S6 in domains I, II, and III (Wang et al., 2005). In summary, the crystal structures from  a number of  K channels (KcsA, MthK, KvAP, Kvl .2, etc.) have contributed significantly  in our understanding of  the molecular mechanisms for activation, permeation, and selectivity. The crystal structures have provided a snapshot of  the channels. However, channels are dynamic proteins, and the detailed conformational  changes rely more on the functional  studies. 1.3 Inactivation in Kv channels Inactivation is an intrinsic property of  Kv channels (Bezanilla, 2000). In the presence of a continuous depolarization, Kv channels enter a stable non-conducting inactivated state, and this process is manifested  as a time-dependent current decay under voltage clamp conditions. The time course of  inactivation can be described by exponential functions  with one or more components, with each component described by an amplitude term (A)  and a time constant (f). With the exception of  the fast  C-type inactivation in HERG, inactivation is generally voltage-independent, but it may be coupled to other voltage-dependent processes or states (Hoshi et al., 1991; reviewed in Yellen, 1998). In Kv channels, the three main types of  inactivation, namely N-type, U-type, and P/C-type, are classified  based on their interactions with intra- or extracellular TEA+ and K+, and they may also differ  in their molecular basis. Table 1.1 summarizes the differences  among the three types of  inactivation. In this section, a brief  description of  the biophysics of  N- and U-type inactivation is followed  by a somewhat more detailed description of the properties of  P/C-type inactivation. Table 1.1 Comparison of  properties of  N-type, U-type, and P/C-type inactivation. Inactivation N-type (fast) U-type P/C-type (slow) Structural basis N-terminal peptide Similar to C-type inactivation (?) S4, Selectivity filter, outer pore Time course Fast (tens of milliseconds) Intermediate (hundreds of milliseconds) Usually slow (seconds) Gating Directly , coupled to open state Coupled to closed states Mainly proceed from open states Extracellular K+ Accelerates recovery from  inactivation Accelerates inactivation Slows rate of inactivation Extracellular TEA+ No effect Accelerates inactivation Slows rate of inactivation Intracellular K+ No effect N/A Slows inactivation kinetics Intracellular TEA' Slows inactivation competitively N/A No effect* * Other quaternary amines with longer mean dwell times can accelerate inactivation (Baukrowitz and Yellen, 1996). 1.3.1 N-Type inactivation Of  the three types of  inactivation, N-type inactivation, also known as fast  inactivation, is by far  the best characterized. In the early 1970's, a transient A-type K current was identified  in neurons of  gastropods (Connor and Stevens, 1971). The A-type current, a term now used to describe all transient K current, was envisioned to arise from  an inactivation process similar to that giving rise to the transient Na current in Nav channels. Just a few  years later, Bezanilla and Armstrong (1977) used internally applied pronase, a proteolytic enzyme, to remove (fast) inactivation in the Nav channel of  squid giant axon, and they proposed a "ball-and-chain" model as the mechanism for  Nav channel (fast)  inactivation. Based on this model, pronase was seen simply to remove the cytoplasmic "inactivation ball." The Shaker  channel from  the fruit  fly  Drosophila melanogaster  also gives rise to a fast inactivating A-type current (Iverson et al., 1988; Timpe et al., 1988). Using an approach similar to that for  studying fast  inactivation in the Nav channel (Bezanilla and Armstrong, 1977), Hoshi et al. (1990) were able to remove (fast)  inactivation from  Shaker  by applying trypsin, another proteolytic enzyme, to the cytoplasmic side of  the channel.. Moreover, deleting residues 2-46 from  the N-terminus of  Shaker  using mutagenesis also resulted in a current that did not inactivate, at least with short depolarizing pulses (Hoshi et al., 1990), and this mutant became known as the inactivation-removed Shaker  channel, or ShakerlR.  Reapplying the N-terminal peptide back into the intracellular medium reconstituted fast  inactivation in ShakerlR  and conferred  fast  inactivation to other Kv channels (Stephens and Robertson, 1995; Zagotta et al., 1990). These results showed conclusively that fast  inactivation is mediated by the N-terminus and thus is now commonly referred  to as N-type inactivation. N-type inactivation has been demonstrated in Shaker  (Hoshi et al., 1990; Zhou et al., 2001a), Kvl .4 (Stuhmer et al., 1989; Tseng-Crank et al., 1993; Zhou et al., 2001a), Kv3.3 (Fernandez et al., 2003), Kv3.4 (Schroter et al., 1991; Stephens and Robertson, 1995), and Kv channels associated with Kvfil.l  subunits (Heinemann et al., 1996) and Kv(3l.3 subunits (Decher et al., 2005), and its mechanism is analogous to a ball-and-chain model. The N-terminal peptides from  ShakerB,  /?Kv3.4, and Kvl.4 have been shown to block the non-(fast)  inactivating mouse Kvl.l channel (Antz and Fakler, 1998; Stephens and Robertson, 1995), and Kvl.4 can be blocked by the N-terminal ball from  Shaker  in addition to its own N-terminal ball (Ruppersberg et al., 1991). However, despite this apparent non-specific  blocking of  Kv channels, the inactivation balls (around 30-40 amino acids in length) show very little homology in their primary sequence or tertiary structure, as revealed by their NMR and crystal structures (Antz et al, 1997; Antz and Fakler, 1998; Schott et al., 1998). The "non-specificity"  arises from  the fact that the N-terminal ball receptor site in the internal cavity is structurally very similar between Kv channels (del Camino et al., 2000; Gulbis et al., 2000; Zhou et al., 2001a). The length of  the linker (i.e.,  the "chain") between the N-terminal ball and the rest of  the channel can affect  the kinetics of  N-type inactivation, such that the rate is slowed if  this linker is lengthened, or accelerated if  part of  the linker is deleted (Zagotta et al., 1990). The ball-and-chain model describes N-type inactivation very well; however, the manner in which the N-terminal ball binds to its receptor site in the central cavity is not as straight forward  as the inactivation cartoons sometime imply. In Kv channels, the Tl domains are folded into a four-fold  symmetric structure that is suspended from  the transmembrane segments like a hanging gondola, as shown in Figure 1.2 B (Kobertz et al., 2000; Kreusch et al., 1998; Long et al., 2005a; Sokolova et al., 2001). Between the Tl-Sl linkers are lateral openings that the N-terminal ball must pass through to gain access to the central pore (Gulbis et al., 2000; Long et al., 2005a; Zhou et al., 2001a). The N-terminal inactivation particle may be in a "pre-inactivated" state as it binds to the Tl domain and/or the S4-S5 cytoplasmic loop prior to activation (Isacoff  et al., 1991; Zhou et al., 2001a). The transition between this pre-inactivated state and the inactivated state may be too fast  to be recorded and thus giving rise to an apparent closed-state inactivation. However, N-type inactivation is strictly coupled to activation (Demo and Yellen, 1991) in that the N-terminal ball can access its binding site only if  the activation gate is open. The binding of  the N-terminal peptide to the internal cavity is mechanistically very similar to that for  an open channel blocker such as intracellular TEA+. Internal TEA+ can slow N-type inactivation by competing with the N-terminal ball for  an overlapping binding site (Choi et al., 1991; del Camino el al., 2000). However, unlike TEA+ and other quaternary ammonium (QA) compounds that can be trapped in the internal cavity upon deactivation, the N-terminal peptide with the tethered chain prevents the closing of  the activation gate (Demo and Yellen, 1991; Ruppersberg et al., 1991). In addition, increasing the extracellular K+ concentration can speed up the rate of  recovery from  N-type inactivation as if  the inward K+ can knock off  the inactivation particle (Demo and Yellen, 1991). Interestingly, mutating the cationic (basic) lysine at position 7 in the N-terminal peptide in Shaker  dramatically reduces fast  inactivation (Hoshi et al., 1990). Furthermore, both the NMR structures of  the N-terminal peptides from  Kv3.4 and Kvl .4 show localized positively-charged surface  residues. On the basis of  the two latter observations, it is tempting to speculate TEA+ and the N-terminal peptide mimic K+ in binding to the receptor site in the central cavity. 1.3.2 U-type inactivation U-type inactivation has been described in Kv3.1 (Klemic et al., 2001), Kv2.1 (Klemic et al., 1998), the Shaker  channel (Klemic et al., 2001), the N-terminal truncated form  of  Kv 1.5 (Kurata et al., 2001; Kurata et al., 2002; Kurata et al., 2005), as well as in N-type and R-type Cav channels (Patil et al., 1998). The term U-type inactivation was coined on the basis of  the U-shaped inactivation-voltage curve, in which the degree of  inactivation at intermediate voltages is greater than at higher voltages (Klemic et al, 2001). It has been proposed that this U-shaped inactivation-voltage relationship arises from  inactivation that proceeds preferentially  from partially-activated states (Klemic et al, 1998). Since the mean dwell time in partially activated y states is longer with intermediate depolarizations, the proportion of  inactivated channels is larger than observed with stronger depolarizations. This property of  U-type inactivation may underlie the strong frequency  dependence of  U-type inactivation, in which the extent of  inactivation with short repetitive depolarization steps is larger than that with a single-pulse depolarization of  the same duration (Klemic et al, 1998; Klemic et al, 2001; Kurata et al, 2001). This "excessive cumulative inactivation" has also been observed in other Kv channels (Aldrich, 1981; DeCoursey, 1990), but whether excessive cumulative inactivation is an exclusive property of  U-type inactivation is still unknown. A number of  properties distinguish U-type inactivation from  N- and P/C-type inactivation. U-type inactivation is accelerated by increasing the extracellular concentration of K+ or TEA+ (Klemic et al, 1998; Klemic et al, 2001; Kurata et al, 2005), which is directly opposite to its effects  on slow inactivation (Choi et al, 1991). In addition, the rate of  recovery from  U-type inactivation at negative potentials is faster  than the rate of  U-type inactivation at depolarized potentials (Klemic etal,  2001). This is different  from  P/C-type inactivation in which the rate of  recovery is usually slower  than the rate of  inactivation. Even though the molecular mechanism for  U-type inactivation is not known, at least in the Shaker  channel, U-type and P/C-type inactivation can be observed simultaneously at different  voltages (Klemic et al, 2001). The coexistence of  these two types of  inactivation argues for  separate underlying mechanisms for  the two processes. It has been proposed that slight changes in the pore structure may be sufficient  for  the channel to switch from  U-type inactivation to P/C-type inactivation and vice versa (Klemic et al, 2001). However, it is still unclear why some channels preferentially inactivate from  the closed-state. Kv4.1 has been proposed to inactivate preferentially  from  a closed-state (Gebauer et al., 2004; Jerng et al., 1999), but its inactivation properties are not completely identical to U-type inactivation, suggesting the existence of  several forms  of  closed-state inactivation (Klemic et al, 2001). Yet, U-type inactivation may be more prevalent than previously thought (Klemic et al, 2001; Patil et al, 1998). 1.3.3 Slow (P/C-type) inactivation When N-type inactivation is removed by N-terminal deletion, the resulting ShakerlR current shows a slow decay process termed slow inactivation (Hoshi et al, 1990). In early studies, the properties of  slow inactivation were found  to depend on the C-terminal splice variant; therefore,  this type of  inactivation was also called C-type inactivation (Hoshi et al, 1991), and the terms have been used synonymously for  some time. By comparing the different C-terminal splice variants of  Shaker,  the residue at position 463 in S6 was found  to determine the kinetics of  slow inactivation: a valine residue (ShakerA)  slowed the decay rate, whereas an alanine at residue 463 (ShakerB)  resulted in a faster  decay. This residue was the first  of  many residues in the pore domain that have been shown to affect  slow inactivation. Slow inactivation manifested  as a time-dependent current decay with a multi-exponential time course, which suggested more than one process was involved. Indeed, a number of  experiments have shown that slow inactivation is at least a two-step process involving conformational  changes in the pore region and the S4 segment (see below); therefore,  slow inactivation is now referred  to as P/C-type inactivation (Chen et al, 2000; Loots and Isacoff,  1998). The terms P-type and C-type inactivation are applied somewhat differently  than originally proposed. Historically, P-type inactivation was proposed to describe an inactivation process in the V369I mutant of  a chimera made by substituting the pore of  Kv2.1 with that from  Kv3.1 (Kirsch et al., 1992). It was found  that the inactivation properties in this mutant were different from  both C-type and N-type inactivation, and the term P-type inactivation was coined because residue 369 is located deep in the pore (De Biasi et al, 1993). Nowadays, the term P-type inactivation is used to describe the conformational  changes in the selectivity filter  and/or the outer pore region (Loots and Isacoff,  1998). Similarly, C-type inactivation was originally used to describe the slow inactivation process observed in ShakerlR-,  however, it now is taken to mean the process resulting in gating charge immobilization (Cha and Bezanilla, 1997; Olcese et al., 1997). We are only beginning to understand the role of  the outer pore mouth, the selectivity filter,  and S4 in mediating this complicated process. 1.3.3.1 Role of  the outer pore in slow inactivation The outer pore has been implicated as one of  the most important structural determinants of  slow inactivation. The classical features  of  slow inactivation (Kiss and Korn, 1998), namely its modulation by the mutations at position 449 in Shaker  (Lopez-Barneo et al., 1993) and its inhibition by external TEA+ (Choi et al., 1991) and external K+ (Lopez-Barneo et al., 1993), are all related to the outer pore. The important role of  Shaker  T449 in slow inactivation is highlighted by the dramatic changes of  current decay observed in the different  T449 mutants. A glutamate (E), lysine (K), alanine (A), glutamine (Q) or serine (S) at the position 449. can dramatically accelerate slow inactivation, whereas a valine (V) or tyrosine (Y) residue at this position results in a very slow or non-inactivating mutant (Lopez-Barneo et al., 1993; Schlief  et al., 1996). Moreover, the T449H mutant shows K+-dependent inhibition of  slow inactivation only at low pH, and in Kvl .3, the histidine residue H399 at the site homologous to Shaker  T449 also slows the rate of  slow inactivation only at low pH (Somodi et al., 2004). In addition, Shaker T449 is also involved in the binding of  external TEA+ (Heginbotham and MacKinnon, 1992; MacKinnon and Yellen, 1990). Mutating this residue to tyrosine (T449Y) or phenylalanine (T449F) increases the TEA+ affinity  by 25- to 50-fold  (Heginbotham and MacKinnon, 1992; Kavanaugh et al., 1992; MacKinnon and Yellen, 1990). The inhibition of  slow inactivation by external TEA+ and K+ was explained by the "foot-in-door" process (Aimers and Armstrong, 1980; Choi et al., 1991; Grissmer and Cahalan, 1989; Lopez-Barneo et al., 1993; Matteson and Swenson, Jr., 1986; Pardo et al., 1992; Swenson, Jr. and Armstrong, 1981). Support for  this hypothesis was provided by Yellen et al (1994) who showed that the outer pore constricted during slow inactivation. Mutating the outer pore residues M448, T449, or P450 to a cysteine (M448C, T449C, or P450C) resulted in a state-dependent block by external Cd2+ and Zn2+ (Yellen et al, 1994), and the T449C mutant was 45,000-fold more sensitive to Cd2+ in the inactivated state than the open state. The M448C and P450C mutants also show a similar state dependent reaction to MTS reagents, and the M448C mutant can also be cross-linked between two subunits by a thiol-reactive compound (Liu et al, 1996). During slow inactivation, the outer pore regions of  the four  subunits are thought to constrict in a highly cooperative manner (Ogielska et al, 1995; Panyi et al, 1995). This constriction may be inhibited physically by external K+ and TEA+ as they bind to the outer pore. It is possible that the external coordination sites (S0 or Sext) are the regulatory sites at which external K+ and TEA+ bind and consequently slow P/C-type inactivation. This hypothesis predicts that ions (K+, TEA+, or other permeant ions such as Rb+, NH4+, Cs+, and Na+) .binding to the external coordination site could prevent the collapse of  the outer pore. Indeed, the extent of attenuation of  C-type inactivation by permeant ions is similar to the permeability ratio with respect to K+ (Lopez-Barneo et al., 1993), which is consistent with the above hypothesis. Similarly, TEA+ was shown to be bound at S0 in a molecular dynamic simulation (Crouzy et al., 2001). In addition, in a KcsA crystal structure, a TEA+ analog was shown to bind to the coordination site S0 and to be coordinated by Y82 (homologous to Shaker  T449) at the outer pore (Lenaeus et al., 2005). These results suggest that the external TEA+ binding site is similar in KcsA and in Shaker,  and that TEA+ may act as a "partially hydrated K+" at S0. However, TEA+ can block certain mutants (e.g.,  Shaker  D447E T449Y) without affecting  the rate of  slow inactivation (Molina et al., 1997). Therefore,  the mechanism by which external TEA+ inhibits slow inactivation may be more complicated than once thought. 1.3.3.2 Role of  the selectivity  filter  in slow inactivation Several lines of  evidence have suggested that the selectivity filter  undergoes conformational  changes during slow inactivation, and that the slow inactivation gate may in fact be the selectivity filter  in a collapsed, non-conducting state (Cordero-Morales et al., 2006a; Cordero-Morales et al., 2006b; Kiss et al., 1999; Kiss and Korn, 1998). The possible involvement of  the selectivity filter  in slow inactivation was first  implied by the inhibition of slow inactivation by permeant ions (Lopez-Barneo et al., 1993). It was later suggested that when the selectivity filter  was depleted of  permeant ions, either following  N-type inactivation (Baukrowitz and Yellen, 1995) or block by internal quaternary amines with long mean dwell times (Baukrowitz and Yellen, 1996), slow inactivation could be dramatically accelerated. It was also proposed that slow inactivation could not proceed before  the last ion had evacuated the selectivity filter  (Baukrowitz and Yellen, 1996; Yellen, 1998). According to this hypothesis, external TEA+ and external permeant ions may attenuate slow inactivation by preventing the evacuation of  K+ from  the selectivity filter  (Kiss and Korn, 1998). The involvement of  the selectivity filter  in slow inactivation can also be appreciated from the various mutations in the P-loop that result in a change in the rate of  slow inactivation. Mutating W434 in the pore helix of  Shaker  to phenylalanine (W434F) results in a non-conducting mutant (Perozo et al., 1993) that is thought to be permanently P-type inactivated (Olcese et al., 1997; Yang et al., 1997), and the homologous mutation in Kvl .5 (W472F) also results in a similar non-conducting mutant (Chen et al., 1997)! When these residues are mapped onto the KcsA structure, the equivalent residue (W67) is in a network of  hydrophobic residues that includes W68 and Y78 (in the G-Y-G-D signature sequence; Figure 1.4) and which is proposed to hold the selectivity filter  in the open conformation  (Doyle et al., 1998). Interestingly, the W434F mutant becomes conducting with the T449V mutation (Kitaguchi et al., 2004), which suggests the two mutations affect  the same process but in an opposite manner. The tryptophan-to-phenylalanine mutation is thought to disrupt the hydrophobic network resulting in the collapse of  the selectivity filter,  and the channel becomes permanently P-type inactivated. This idea is supported by the fact  that normal gating charge movement is observed in this mutant, which also suggests that the gating charges are not immobilized and that the channel is not C-type inactivated. An apparent "collapse" of  the selectivity filter  is also observed in the absence of  external permeant ions, during which some K channels may enter a non-conducting state that is related to r slow inactivation (Yellen, 1997) and distinct from  the defunct  state (Loboda et al., 2001; Melishchuk et al., 1998). Thus, in Kvl.4, removal of  external permeant ions results in a conductance collapse due to a large decrease of  channel availability. This external K+-dependent conductance collapse is absent in the K533Y-I535M mutant (Pardo et al., 1992). The involvement of  K533, which is homologous to Shaker  T449, suggests that the collapse of  Kvl.4 conductance is related to a slow inactivation process. In the absence of  external K" or other permeant ions, the BK channel and the plant inward rectifier  K channel KAT1 and ZmK2.1 also fail  to conduct current (Hertel et al., 2005; Su et al., 2005; Vergara et al., 1999). This "conductance collapse" is also observed in Kvl.3 at pH 6.0, the Shaker  T449A, T449E, T449K mutants, and the Kvl.5 H463G mutant (Jager et al., 1998; Jager and Grissmer, 2001; Lopez-Barneo et al., 1993; Su et al., 2005). These channels are all proposed to enter a stable non-conducting state resulting in channels being unavailable for  activation. In the inward rectifier Kir 1.1, reducing external [K+] promotes an inactivated mode of  gating which is similar to the conductance collapse described above (Sackin et al., 2001). Together, these results highlight the importance of  permeant ions in channel function,  in which K+ in the pore prevents a "conductance collapse" related to an inactivation process. Another clue to the involvement of  the selectivity filter  in slow inactivation is the change of  selectivity observed during slow inactivation (Yellen, .1998). In the absence of  K+ on both sides of  the membrane, slow inactivation was estimated to be complete in milliseconds (Baukrowitz and Yellen, 1996). It was expected that, under this K+-free  condition, the outer pore of  the channels would become constricted due to slow inactivation, and channels would be non-conducting. However, removing K+ from  both sides of  the channels resulted in measurable currents through Shaker,  Kvl .5, and Kv2.1 that were carried by Na+ (Korn and Ikeda, 1995; Starkus et al., 1997; Wang et al., 2000). That is, inactivated channels became Na+ permeable (Kiss et al., 1999). This Na+ current was also observed in the permanently P-type inactivated "non-conducting" Shaker  W434F mutant in the absence of  K+ on both sides of  the membrane (Starkus et al, 1998). It appeared that the selectivity filter  became constricted during slow inactivation such that the coordination sites could accommodate Na+ better than in the "open" conformation. Besides the more "defined"  functional  studies, a number of  recent structural studies have proposed the molecular basis for  the conformational  changes during slow inactivation. From the KcsA crystal structure, a structural rearrangement at T75 and V76 during K+ conduction was predicted by a molecular dynamics study to occur spontaneously, and this rearrangement was suggested to be an initial step in slow inactivation (Berneche and Roux, 2005). In a recent study, the residues E71 and D80 in KcsA are suggested to undergo a conformational  change during some gating processes (Cordero-Morales et al, 2006a). It is uncertain whether this conformational  change is related to a slow inactivated state in Kv channels, but these studies have opened up new possibilities for  further  investigations. 1.3.3.3 Role ofS4  in slow inactivation In addition to participating in voltage sensing during activation, S4 is also involved in slow inactivation. Studies using fluorescent  probes have shown that changes in fluorescence project from  the pore out to S4 during slow inactivation (Gandhi et al, 2000; Loots and Isacoff, 1998; Loots and Isacoff,  2000). However, the precise interaction of  the voltage-sensing domain and the pore domain during slow inactivation is still largely unknown. The involvement of  S4 in slow inactivation is implicated most directly in the process called gating charge immobilization (Fedida et al, 1996; Olcese et al, 1997). Gating charge immobilization was first  described in Nav channels (Armstrong and Bezanilla, 1977). With short depolarizing pulses, the ratio of  off-gating  charge to on-gating charge (Q of f:Q on ratio) was close to one, suggesting that most of  the gating charges activated (mobilized) during activation returned to the resting level upon repolarization. However, with longer depolarizing pulses, the Qof f:Q on ratio became progressively smaller, and the kinetics of some of  the o/f-gating  current was much slower. Together, these observations indicated that the gating charges moved back to the resting state but only very slowly, as if  some of  the gating charges were being "immobilized." This apparent loss of  gating charges is described as gating charge immobilization. In ShakerlR,  a similar leftward  shift  of  the Qof f-  V  curve (i.e.,  charge immobilization) was observed after  the channels were inactivated, but all the gating charge could be seen to return to the resting state if  a strong hyperpolarizing pulse was given (Olcese et al., 1997). It was shown that the time course of  gating charge immobilization followed  that of  slow inactivation, and the time course of  recovery from  gating charge immobilization also correlated well with that of recovery from  slow inactivation (Olcese et al., 1997). In addition, gating charge immobilization in Kvl .5 was inhibited by 4-aminopyridine (4-AP), which inhibited slow inactivation by inhibiting channel opening (Fedida et al., 1996). It was proposed that gating charge immobilization is part of  the C-type inactivation process (Loots and Isacoff,  1998). The conformational  changes occurring during charge immobilization and C-type inactivation are still unknown, but an interaction between the pore domain and S4 was postulated (Loots and Isacoff,  2000). A possible interacting pair is the activated S4 and a conserved glutamate residue (E418 in Shaker)  at the base of  the turret (Larsson and Elinder, 2000; Loots and Isacoff,  2000). In the Kvl .2 crystal structure the pore domain (in particular the S5) of  one subunit is adjacent to the S4 of  another subunit (Long et al., 2005a). In Shaker,  E418 is suggested to be positioned adjacent to the top of  S4 during activation (Loots and Isacoff,  2000), and it is thought to interact with G452 in the P-S6 linker to stabilize the open state. This interaction is thought to be broken during slow inactivation, as shown by E418 coming close to V451 in an inactivated state (Loots and Isacoff,  2000). Yet, it is uncertain which residues actually move during slow inactivation and how S4 fits  into this picture. Nevertheless, as suggested by the Shaker  ILT mutant, the movement of  S4 itself  may not trigger slow inactivation. In this mutant, the movement of  S4 is "energetically separated" from  the concerted opening of the activation gate (Pathak et al., 2005); that is, the voltage range over which S4 movement occurs is not sufficient  to open the activation gate, which requires a much stronger depolarization. Even at potentials in which S4 is fully  activated, this mutant does not seem to undergo slow inactivation until the activation gate is open. Conversely, slow inactivation has been suggested to occur from  one or more closed states (Olcese et al., 1997; Yang et al., 1997). It is uncertain whether slow inactivation occurring from  open state is structurally equivalent to that occurring from  closed states. To summarize, slow (P/C-type) inactivation has been shown to result from  a constriction of  the outer pore and/or a constriction at the selectivity filter,  both of  which may underlie P-type inactivation. A slower process that involves an interaction between the pore domain and S4 results in gating charge immobilization that underlies C-type inactivation. Surprisingly, the turret region, which forms  part of  the outer vestibule, has, except for  a few  reports (Perchenet and Clement-Chomienne, 2001; Steidl and Yool, 1999; Zilberberg et al, 2001, been largely ignored in the study of  slow inactivation,. That the turret may play an important role in slow inactivation is suggested by results described in this thesis. 1.4 Kvl.5 and scope of  thesis investigation The human voltage-gated potassium channel Kvl .5, encoded by the gene KCNA5, is a delayed rectifier  K channel expressed in atrial and, to a smaller extent, ventricular myocytes (Fedida et al, 1993; Mays et al, 1995), vascular and intestinal smooth muscles (Overturf  et al, 1994), pulmonary arteiy smooth muscles (Archer et al, 1998), pancreatic islets (Philipson et al, 1991), and microglia in the central nervous system (Jou et al, 1998). In human atrial myocytes, Kvl.5 mediates the ultra-rapid delayed rectifier  current (IKur), which is partly responsible for repolarizing the atrial action potential and for  the determination of  the plateau duration (Fedida et al, 1998; Feng et al, 1997). Simulations have shown that a reduction in IKu r can lead to a prolongation of  the atrial action potential (Gomez et al, 2005), and have suggested that a reduction in Kvl .5 current may have therapeutic benefit  for  treating atrial fibrillation  (Brendel and Peukert, 2003; Matsuda et al, 2001). The role of  Kvl.5 in the central nervous system is not well understood. The homotetrameric Kvl.5 channel is a Shaker-related  channel with some biophysical properties that differ  from  those of  Shaker.  In contrast to the Shaker  channel, wild-type Kvl .5 does not have an N-terminal inactivation ball and thus does not undergo N-type inactivation unless associated with Kv (31.1 or (51.3 subunits (Heinemann et al,, 1996). Furthermore, wild-type Kvl .5 is not sensitive to external TEA+ block, presumably due to the positively charged arginine residue at position 487, homologous to Shaker  T449, given that the R487V mutation confers  (external) TEA+ sensitivity to Kvl .5 (Fedida et al, 1999). Similar to the Shaker  channel, Kvl.5 is sensitive to block both by 4-AP and quinidine (Bouchard and Fedida, 1995; Fedida, 1997; Wang et al, 1995). The blocking mechanism for  4-AP and quinidine is thought to be similar in Kvl .5 and Shaker,  in which 4-AP stabilizes an activated closed-state (Armstrong and Loboda, 2001) and quinidine acts as an open channel blocker (Fedida, 1997). Given that the single channel conductance is similar between Kvl .5 and Shaker,  it is thought that the conducting pore of  these two channels is similar. As mentioned above, in the absence of  intracellular and extracellular K+, both the Kvl.5 (Wang et al, 2000) and the Shaker  channel (Starkus et al., 1997) are permeable to Na+, and both the Shaker  W434F mutant and the equivalent Kvl .5 W472F mutant do not conduct K+ current (Chen et al, 1997; Hesketh and Fedida, 1999; Perozo et al, 1993). Interestingly, the Kvl .5 R487V mutation strongly attenuates slow inactivation in Kvl.5 when Na+ is the charge carrier, but no significant  difference  in the inactivation kinetics is observed with K+ as the charge carrier (Wang et al, 2000), a phenomenon that is not yet explained. This is in contrast to the finding  in the Shaker  channel where a valine at the equivalent position (T449V) strongly inhibits slow inactivation with K+ current (Lopez-Barneo et al, 1993; but see Holmgren et al, 1996). In addition, external K+ has been shown to inhibit slow inactivation in the Shaker  channel (Lopez-Barneo et al, 1993), but external K+ seems to have a much more muted effect  on the rate of  slow inactivation in Kvl .5 (Fedida et al., 1999). The molecular basis for  these differences  has not yet been identified,  but the molecular mechanisms underlying slow inactivation are assumed to be similar in Kvl .5 and in Shaker. This dissertation examines the mechanistic basis for  the modulation of  /zKvl.5 (Kvl.5) slow inactivation by external H+, Ni2+, K+, and some other divalent cations. The goal for  this research project is to increase our understanding of  slow inactivation gating of  ion channels, which may in turn lead to a better understanding of  the structure-function  relationship of  slow inactivation and the conformational  changes associated with slow inactivation. The impetus for this work was the observation in our laboratory that external Zn2+ could inhibit Kvl .5 current in a K+0-dependent manner (Zhang et al., 2001). However, the binding site for  Zn2' and the mechanistic basis for  its inhibitory effect  were not identified  in that study. Therefore,  the first goal of  this research project was to ascertain the molecular basis for  the Zn2 r-induced current inhibition (Chapter 2). To do this, Kvl .5 macroscopic currents were studied in various conditions using conventional voltage clamp techniques in the whole-cell and outside-out configurations.  The turret histidine residue at position 463 (H463) was proposed to be the binding site for  Zn2+, and this binding of  cations resulted in a current inhibition mediated by a process involving the outer pore arginine residue (R487). In addition, the properties of  the current inhibition were similar to the classical features  of  slow inactivation. It was also found that external H+ also caused an inhibition of  Kvl.5 with similar properties as that induced by Zn2+. Therefore,  the current inhibition induced by external Zn2+ and H+ in Kvl .5 was / hypothesized to result from  an enhancement of  a slow inactivation process that led to the reduction in channel availability. This hypothesis was tested further  with external Ni2+ and other divalent cations, and a similar current inhibition was observed, albeit with somewhat different  effects  on other biophysical properties (Chapter 3). From the unitary currents recorded with Ni2+, the current inhibition was found  to result from  an increased probability that a channel failed  to report current upon depolarization, which was again consistent with our hypothesis. The single channel analysis was continued in Chapter 4 but focussed  on the effects  of  changes of  external pH on gating behaviour. When the unitary current behaviour of  Kvl .5 was systematically analysed at different  pHs, a shift  in the mode of  gating was revealed, and the shift  was proposed to result from  an inactivation process proceeding from  closed states. Further support for  channels being in an inactivated state at low pH was obtained by analysing the first  latency behaviour and the gap lengths between bursts (Chapter 5). Using long (> 6 s) depolarizing pulses, channels were sometimes shown to open with a long delay (first  latency), which was proposed to result from  the recovery from  an unavailable mode of  gating (channel failed  to conduct; mode U)  to a normal mode of  gating (channel able to conduct; mode A). The mean (long) first  latency was found  to correlate with the mean (long) gap length. This result suggested that mode U  gating involved an inactivated state. Chapter 5 also presents evidence that K+ antagonizes the H+-induced current inhibition by promoting mode A gating. Together, the results support the hypothesis that the external H+-induced current inhibition resulted from  promotion of  an inactivation process from one or more closed states. 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Molecular determinants of  the inhibition of  human Kvl.5 potassium currents by external protons and Zn2+ 2.1 Introduction We have previously shown in human Kvl .5 (hKv\.5)  channels that external Zn2+ ions caused a depolarizing shift  of  the activation curve, an effect  referred  to as the gating shift,  as well as a reduction of  the current amplitude, which we termed block, that was relieved by external permeant ions such as K+ and Cs+ (Zhang el al., 200lb). Although a gating shift  is very often associated with a change of  the external concentration of  divalent cations, as first  detailed by Frankenhaeuser and Hodgkin (1957), the block of  voltage-gated K+ channels by divalent cations, in particular Zn2+, is not a general finding  and its mechanistic basis if  therefore  of  some interest. In a follow-up  study of  the effects  of  Zn2+ on gating currents (Zhang et al., 2001a), we found  that Zn2+ ions caused a rightward shift  of  the voltage dependence of  on-gating charge movement or the QON(V)  relationship. This Zn2+-induced shift  of  the OJV)  was approximately 2-fold  greater than that observed for  the conductance-voltage (g(V))  relationship and implied that Zn2+ binding at a site in the outer channel pore could inhibit the ionic current as well as the movement of  the voltage sensor. The latter effect  is consistent with recent evidence for  a close proximity of  the outer pore mouth and the S4 segment which comprises a major part of  the voltage sensor (Blaustein et al., 2000; Cha and Bezanilla, 1998; Li-Smerin et al., 2000; Loots and Isacoff,  1998) and with the view that the voltage sensing domain wraps around the outer rim, i.e., the turret of the pore (Loots and Isacoff,  2000). A version of  this chapter has been published. Kehl, S. J., Eduljee, C., Kwan, D. C. H., Zhang, S., and Fedida, D. (2002) Molecular determinants of  the inhibition of  human Kvl.5 potassium currents by external protons and Zn2+. J. Physiol. 541(l):9-24. Identifying  the potential site(s) of  action of  Zn2+ is facilitated  by the fact  that Zn2+ shows high affinity  binding to the imidazole ring of  histidine (H) or to the sulphur atom of  cysteine (C) residues. Lower affinity  binding of  Zn2+ can also occur at the side chain carboxyl group of  the acidic amino acid residues glutamate (E) and aspartate (D) (Vallee and Auld, 1990). In the six transmembrane segment (6TM) a-subunit of  MCvl .5 there is in the pore-forming  (P-) region and the extracellular segments linking it to S5 (S5-P) and S6 (P-S6) only one high affinity  Zn2+ ligand, H463, which, based on the crystal structure of  KcsA channels (Doyle et al., 1998), is found  in the turret. Other potential Zn2+ binding sites include, at the NH2-terminal end of  S5-P, a glutamate residue, E456, that is strictly conserved in voltage-gated channels (Ortega-Saenz et al., 2000) and which appears to interact with S4 (Loots and Isacoff,  2000). E456 is, however, an unlikely Zn2+ binding site at least in part because its side chain carboxyl group is believed to form hydrogen bonds with residues in the P-S6 region (Larsson and Elinder, 2000). Two aspartate residues are also found  in the outer pore mouth of  hKvl  .5, D469 in the outer pore helix and D485 which forms  part of  the "GYGD" motif  in the pore signature sequence. Interestingly, the inhibition of  rat Kvl.5 (rKvl .5) currents by extracellular protons (H+0) (Steidl and Yool, 1999) has features  similar that of  the Zn2+ block of  hKvl .5. Although not evident with Zn2+ during 300 ms pulses (Zhang et al., 2001b), an acceleration by H+0 of  current inactivation during long-lasting (> 1 s) depolarizations raised the possibility that the current inhibition was due to an accumulation of  channels in the C-type inactivated state (Steidl and Yool, 1999). The term "C-type inactivation" originated with the observation that Shaker  splice variants with different  carboxy-terminal regions (including S6) showed different  rates of  slow inactivation (Hoshi et al., 1991). Subsequently it was reported that mutations in the pore (P) region of  Kv2.1 channels increased the inactivation rate by a process having different characteristics than C-type inactivation and the term "P-type inactivation" was coined (De Biasi et al., 1993). A growing body of  evidence now suggests that slow inactivation in Shaker-related channels such as Kvl .5 is in fact  a complex process involving either multiple and independent inactivation mechanisms or a single inactivation process that involves multiple steps (Kiss et al., 1999; Loots and Isacoff,  2000; Olcese et al., 1997; Wang and Fedida, 2001; Yang et al., 1997). P-type inactivation appears to involve a partial constriction of  the outer pore mouth that eliminates K+ currents but has no effect  on gating currents including their ability to undergo charge immobilization following  the "closed to open" transition (Yang et al., 1997). C-type inactivation might be coupled to P-type inactivation and has been proposed to involve a farther conformational  change of  the outer pore that stabilizes the S4 segments in the activated or outward position. This "stabilization" contributes to a voltage-dependent slowing of  return or off-gating  charge movement, an effect  that is also known as charge immobilization (Olcese et al., 1997; Wang and Fedida, 2001). The possibility that a common mechanism of  action, possibly involving an inactivation process, might account for  the inhibition of  Kvl.5 current by Zn2+ and H+ also pointed to the potential for  a common site of  action. In this connection it is known that Zn2+ and H+ bind to histidine residues and a rKvl.5 mutant in which histidine residues in the pore turret are replaced by glutamine (Q) (rKvl .5 H452Q) has a substantially reduced acid sensitivity (Steidl and Yool, 1999). Against this background the experiments described here had two major goals. First, to determine the concentration dependence of  the inhibition of  hKv\  .5 channels by protons and to discover if,  as with Zn2+, the inhibition by H+0 was affected  by changes of  K+0. After  confirming a K+0-sensitive inhibition of  MCvl.5 currents by H+0, the second goal was to gain at least a preliminary insight into the molecular basis for  that inhibitory action by assessing the effects  of point mutations. We provide evidence that the binding of  H+ or Zn2f  to histidine residues (H463) in the channel turret is a necessary but not a sufficient  condition for  the inhibitory effect.  Instead, H463s appear to function  as sensors, such that H+ or Zn2+ binding permits a conformational change that involves an arginine (R) residue near the pore mouth (R487). An examination of gating currents revealed that H+0 (pH 5.4) has no effect  on the total gating charge movement (Qmax)  a n d that charge immobilization persists following  strong depolarizations. Based on these observations, we propose that protons and Zn2+ ions inhibit /zKvl .5 currents by affecting  channel availability. The possibility that the H+ 0 inhibition of  /zKvl .5 currents arises by the facilitation  of a transition to a non-conducting state, possibly the P-type inactivated state, is discussed. 2.2 Materials and Methods 2.2.1 Cell preparation Wild type (wt) /?Kvl.5 channels were studied in a human embryonic kidney cell line (HEK-293) as reported previously (Wang et al., 2000). Cells were dissociated for  passage by using trypsin-EDTA and were maintained in minimum essential medium (MEM), 10% fetal bovine serum, penicillin-streptomycin and 0.5 mg ml'1 gentamicin in an atmosphere of  5% C02 in air. All tissue culture supplies were obtained from  Invitrogen (Burlington, ON, Canada). Point mutations of  the wt hKv\  .5 a-subunit in the plasmid expression vector pcDNA3 were made using the Quikchange Kit (Stratagene, La Jolla, CA, USA) to convert the histidine (H) residue at position 463 to glutamine (Q) (H463Q) or glycine (G) (H463G). The double mutant H463Q,R487V was created by subcloning a cassette of  /zKvl.5 H463Q into MCvl.5 R487V (Wang et al., 2000) using BstEII and Clal restriction enzymes (New England BioLabs, Beverly, MA, USA). Stable transfections  of  HEK-293 cells were made using 0.8 |ig of  /zKvl .5 H463Q, /zKvl.5 H463Q,R487V or /?Kvl.5 H463G cDNA and 2 |iL of  Lipofectamine  2000 (Invitrogen): Geneticin (0.5mg/mL) was added 48 hrs after  transfection. 2.2.2 Recording solutions The standard bath solution contained, in mM, 140 NaCl, 3.5 KC1, 10 HEPES, 2 CaCl2, 1 MgCl2, 5 glucose and its pH was adjusted to 7.4 with NaOH. Where the effect  of  the external concentration of  potassium (K+0) on the proton block was examined, zero K+0 solution was made by substituting NaCl for  KC1 and, for  K+0greater than 3.5 mM, NaCl was replaced by KC1. Cs+-containing solutions were prepared by substitution of  CsCl for  KC1 (3.5 mM Cs+0) or for  both KC1 and NaCl (20 mM and 140 mM Cs+0). In experiments addressing the effect  of  Na+0 on the current inhibition, N-methyl-D-glucamine (NMG+) replaced Na+ and the pH was adjusted with HC1. The external concentration of  H+ (H+0) was adjusted with 10 mM of  buffer  where the buffer was HEPES for  the pH range 6.8 to7.4, MES for  pH 5.4 to 6.7 or TAPS for  pH 8.4. Zn2+-containing test solutions were made by the addition of  ZnCl2 from  0.1 or 1 M stock solutions. The low solubility of  Zn(OH)2 limits the maximum concentration of  Zn2+ that can be used at pH 7.4 to less than 5 mM. Our standard patch pipette solution for  recording K+ currents contained 130 KC1, 4.75 CaCl2 (pCa2+ - 7.3), 1.38 MgCl2, 10 EGTA , 10 HEPES and was adjusted to pH 7.4 with KOH. For gating current recordings the bath solution contained, in mM, 140 NMGC1, 1 MgCl2, 10 HEPES (pH 7.4) or MES (pH 5.4), 2 CaCl2, 10 glucose and the pH was adjusted with HC1; the patch pipette solution consisted of  140 NMGC1, 1 MgCl2, 10 HEPES, 10 EGTA and was adjusted to pH 7.2 with HC1. Chemicals were from  the Sigma Aldrich Chemical Co. (Mississauga, ON, Canada). In an experiment, a section of  glass coverslip with cells attached to it was placed in the recording chamber (0.5 ml volume) and perfused  with 5-10 ml of  control solution. After recording control currents the chamber was flushed  with 5-6 ml of  test solution to ensure complete replacement of  the bath solution before  treated responses were recorded. If  after perfusing  5-6 ml of  control solution the post-treatment currents did not recover to within ± 10% of  the pre-treatment amplitudes the entire data set was discarded. In most cells, however, virtually complete recovery was observed. We found  no difference  between experiments done with discontinuous perfusion,  as described above, and experiments where the cells were continuously perfused  (not shown). 2.2.3 Signal recording and data analysis Macroscopic currents were recorded at room temperature (20-25°C) using the patch clamp technique primarily in the whole cell configuration.  In some of  the cell lines expressing mutant MCvl.5 channels at a high level, i.e., the H463Q and some of  the R487V mutants, the large amplitude of  the whole cell currents necessitated recording macroscopic currents from outside-out patches. Voltage clamp experiments were done with an EPC-7 patch clamp amplifier and Pulse+PulseFit software  (HEKA Electronik, Germany). Patch electrodes were made from thin-walled borosilicate glass (World Precision Instruments, FL, USA) and had a resistance of 1.0 to 2.5 MQ measured in the bath with standard internal and external saline. Capacitance and series resistance compensation, typically 80%, were used. An on-line P/N  method, for  which the holding potential was -100 mV and the scaling factor  was 0.25, was used to subtract leak and any uncompensated capacitive currents. Current signals filtered  at 3 kHz (-3dB, 8-pole Bessel filter) were digitized (16 bit) at a sampling interval of  100 |is (10 kHz). Voltages have been corrected for  the liquid junction potentials. To quantify  the inhibition of  currents, tail currents were recorded at -50 mV following  a depolarizing pre-pulse. Peak tail current amplitudes were then obtained by extrapolation of  a single exponential function  fitted  to the tail current decay to the start of  the step to -50 mV. After normalization of  tail currents either to the maximum current of  the control or the treated response, data points were fitted  to a single Boltzmann function: where, when y is the current normalized with respect to the control response, A is the proportion of  the control gmax. When y is the current normalized with respect to the maximal treated current, A is the best fit  value for  the normalized maximal response and ideally has a value of  unity. V, : is the half-activation  potential or mid-point of  the activation curve, Fis the voltage during the pre-pulse and s is the slope factor,  in mV, reflecting  the steepness of  the voltage dependence of gating. To quantify  gating charge movement during activation or deactivation, charge-voltage (QfV))  curves were generated by time integration of  on- or o//-gating currents as described previously (Chen et al., 1997). Activation gating in ZjKvI .5 is best fit  by the sum of  two Boltzmann functions  where the larger component, known as Q2, represents -80% of  the total charge movement (Hesketh and Fedida, 1999). However, for  simplicity, QfV)  data obtained at pH 7.4 and 5.4 were fitted  to Equation (2.1) where y is the charge moved, A is the maximal charge (Qm a x) and Vis  the voltage at which the o«-gating charge (OJ  or off-gating  charge (Q of f)  is evoked. V V i and .s' remain as described above. Concentration-response data for  Zn2+ were fitted  to the Hill equation: A (2.1) y 1 + [Zn 2+] V KZn  J (2.2) where y is the proportion of  the control gmla, K Zn is the equilibrium dissociation constant for  Zn2+ binding and nH  is the Hill coefficient  reflecting  the number of  Zn2+ ions binding per channel. For protons, Equation (2.2) was modified  to account for  the fact  that the data points were normalized to the response at pH 7.4: 7 K H""  +(1Q-7-4)"" K H"»  +[H +)0"" (2.3) where K H  is the equilibrium dissociation constant for  proton binding. The equation used to model the binding interaction between K+0, H+ 0 and the /?Kvl.5 channel has been described previously (Zhang et al, 2001b) and is also known as the ternary-complex model of  interaction that is used to describe the binding of  two ligands to the same receptor. y = (2.4) where K H  is the equilibrium dissociation constant for  proton binding in zero K+0, K K  is the equilibrium dissociation constant for  K+ binding at pH 7.4 and the parameter a is known as the cooperativity factor  or the coupling constant/factor.  A value for  # greater than unity indicates negative cooperativity. To reduce the number of  parameters in the model the Hill coefficient  for K+ binding was assumed to be unity. Data are expressed as the mean ± the standard error of  the mean (SEM) except for  the values obtained by non-linear least-squares fitting  routines (Igor, Wavemetrics, OR, USA) which are expressed as the mean ± the standard deviation (SD). The paired-sample t test (control versus treated) was used to assess the actions of  protons and Zn2+ on the inactivation rate. A p-\alue of 0.05 or less was considered significant. 2.3 Results 2.3.1 Increasing [H+]0 causes a gating shift  and reduces the maximum conductance (g^) Representative traces in Figure 2.1 show the effect  of  changing pH0 from  7.4 to pH 6.4 in nominally K+-free  medium (zero K+0) in which Na+ was the major extracellular cation. For the control currents the voltage protocol consisted of  a 300 ms pulse command to between -40 and 40 mV in 5 mV increments followed  immediately by a 300 ms command to -50 mV to record the tail current. The robust pulse and ensuing tail currents, shown at a higher gain in the inset, obtained during or following  strong depolarizations in pH 7.4 medium (Figure 2.1 A) are consistent with a failure  of  MCvl .5 currents to disappear or "collapse" in zero K+0 (Jager and Grissmer, 2001). After  switching to pH 6.4 medium the range of  the pulse voltages was changed from  -30 to 60 mV to compensate for  a small rightward shift  of  the voltage dependence of  gating, the so-called gating shift.  As noted by Steidl & Yool (1999), there appeared also to be a slight slowing of  the activation kinetics with extracellular acidification  but this was not systematically studied and was certainly not as pronounced as the slowing caused by Zn2+ (Zhang et al., 2001b). A more profound  effect  of  the increased extracellular acidity, and the main focus  of  this report, was a large reduction of  pulse and tail current amplitudes. Figure 2.1 C, which was derived in part from  the traces shown in Figure 2.1 A and B, plots the tail current amplitudes at -50 mV measured as described in Materials and Methods and fitted  to a Boltzmann function.  In this cell, increasing H+ 0 caused V, : to shift  from  -6.2 mV to 4.3 mV and the maximal tail current amplitude at pH 6.4 was 14% of  that measured at pH 7.4. Both the gating shift  and the current reduction reversed completely and rapidly (e.g.,  Figure 2.9) after  returning to pH 7.4. External protons have been reported to reduce the amplitude of  rKvl.5 currents (Steidl and Yool, 1999); however, the reduction (-40%) was substantially less than shown in Figure 2.1 (-85%) and in the left  column of  Figure 2.2 which summarizes the results obtained in 12 such experiments. Since our previous work showed that the reduction of  hKvl  .5 currents by Zn2+ was affected  by K+0, we next addressed the possibility that this apparently greater potency of  the inhibition by protons of  MCvl .5 currents shown in Figure 2.1 was due to the use of  a zero K+ bathing solution. 2.3.2 Increasing [K+]0 inhibits the reduction of  g ^ by extracellular acidification Current traces in the left,  centre and right columns of  Figure 2.2 were recorded from  cells in which K+0 was zero, 3.5 and 140 mM , respectively, and the pH0 was changed from  7.4 (upper row of  traces) to 6.4 (lower row of  traces). The voltage protocol was similar to that described for Figure 2.1. Graphs at the foot  of  each column show the tail currents from  a number of  similar experiments for  control (empty  circles)  and treated responses normalized either with respect to the maximum control tail current (filled  circles')  or to the maximum treated tail current (filled A pH 7.4, 0 mM K*0 40 mV -50 mV -80 mvJ -40 mV 1 4 nA 500 pA 100 ms pH 6.4, 0 mM K*. 60 mV 50 mV 100 pA Figure 2.1 Extracellular acidification decreases the maximum conductance (g^) and causes a rightward shift  of  the conductance-voltage (g(V))  relationship for  Kvl.5 currents. Panels A and B show, respectively, representative control (pH 7.4) and treated (pH 6.4) currents evoked by the voltage protocol shown above each family of  traces. Successive pulse command voltages were incremented by 5 mV but for clarity only alternate traces are shown. The change of  the range of  pulse voltages at pH 6.4 was necessary to compensate for  the gating shift.  The holding potential in this and other figures  was -80 mV, except where noted. Inset traces show the tail currents at a higher gain. Tail current amplitudes, obtained by extrapolating the fit  of  a single exponential function  to the start of  the step to -50 mV, are plotted in C and fitted  to a Boltzmann function  to obtain the equivalent of  the g(V)  relationship. Acidification shifted  the V, / : from  -6.2 mV to 4.3 mV and the maximum current decreased from 1.7 nA to 0.24 nA which corresponds to a gmax relative to that at pH 7.4 (relative gmax) of  0.14. Voltage (mV) squares). That K+ 0 inhibits the proton-induced current reduction is shown by the increase of  the relative gmax from  0.19 ± 0.02 (n = 12) in zero K+0, to 0.56 ± 0.01 (n  = 6) in 3.5 mM K+0 and finally  to 0.81 ± 0.12 (n = 6) in 140 mM K+0. As with the Zn2+ block, the gating shift  at pH 6.4 was not significantly  affected  by changes of  K+0 (see Figure 2.2 legend for  V,,  and 5 values) suggesting that the proton-induced gating shift  and current inhibition are independent effects. [K*]0 = 0 mM [K*]0 = 3.5 mM [K*)0 = 140 mM pH 7.4 < , c pH 6.4 1.0 -, <N I 100 ms -40 -20 0 20 Voltage (mV) f f Voltage (mV) Voltage | Figure 2.2 Increasing K+0 reduces the inhibition of  AKvl.5 current by protons. Traces obtained from  three different  cells showing the current at pH 7.4 (control, top row) and pH 6.4 (treated, lower row) in, from  left  to right, zero, 3.5 and 140 mM K+0. In zero K+0, control and treated pulse currents were evoked by 300-ms pulses from  -50 to 45 mV in 5 mV steps; in 3.5 mM K+0, the pulse command range was -50 to 45 mV at pH 7.4 and -30 to 65 mV at pH 6.4; in 140 mM K+0, the range for  pulse voltages was -40 to 55 mV. For clarity, only alternate current traces are shown. The corresponding control (o) and treated g(V)  relationships, obtained from  a number of  similar experiments, are shown in the graph at the bottom of  each column. Treated data were normalized with respect both to the gmax at pH 6.4 (•) and to the control gmca (•). The relative gmax at pH 6.4 in zero, 3.5 and 140 mM K+0 was 0.19 ± 0.02 (n  = 12), 0.56 ± 0.01 (n  = 6), and 0.81 ± 0.12 (n  = 6), respectively. In zero K+0, V m and 5 changed from  -21.4 ± 4.3 and 4.7 ± 0.5 mV at pH 7.4 to -8.2 ± 4.0 and 7.1 ± 0.3 mV at pH 6.4, respectively. In 3.5 mM K+0, the corresponding values were -18.3 ± 1.9 mV and 3.9 ± 0.4 mV at pH 7.4 and -10.5 ± 1.2 mV and 3.9 ± 0.4 mV at pH 6.4; and, in 140 mM K+0 -26.2 ± 1.1 mV and 3.8 ± 0.3 mV at pH 7.4 and -12.3 ± 1.1 mV and 3.8 ± 0.4 mV at pH 6.4.' Data obtained by repeating experiments of  the type shown in Figure 2.2 over a range of pHs were fitted  to the Hill equation to generate the concentration-response curves shown in Figure 2.3. In zero K+0 medium in which Na+ was the predominant metal cation {filled  circles) the best fit  to the data gave a K H  of  153 ± 13 nM (pK H  ~ 6.8) and a Hill coefficient,  nH,  of  1.5 ± 0.2 which suggests that inhibition requires protonation of  at least two sites. To determine if  Na+ ions affect  the current inhibition by protons, the zero K+0 experiments were also done with NMG+ as the major extracellular cation. With NMG+0, the K„  was 128 ± 53 nM (pK H  ~ 6.9) and nH  was 1.2 ± 0.5 (open circles  and dashed  line of  Figure 2.3). This suggests that external Na+ ions do not affect  the current inhibition by protons. With 5 mM K+0, the K H  increased to 590 ± 85 nM (pK H ~ 6.2), but the value for  nH  of  1.6 ± 0.4 was not significantly  different  from  that with zero K+0. In comparison to the substantial rightward shift  caused by increasing K+0 from  zero to 5 mM, a much smaller increase of  the K H  to 1.1 ± 0.11 |iM (pK H  ~ 6) was obtained when K+0 was increased from  5 to 140 mM. The nH  inl40 mM K+0 was 1.8 ± 0.3. 2.3.3 K+0 relief  of  the effect  of  protons is fitted  by a model of  non-competitive inhibition As noted with Zn2+ block of  /zKvl .5 channels (Zhang et al., 2001b), the greater relief  of the proton-induced current inhibition when K+0 was changed from  zero to 5 mM K+0 compared to when it was changed from  5 to 140 mM K+0, suggested that K+ ions and protons were not competing for  a common site. For that reason we modelled the interaction between K+ 0 and H+ 0 as an allosteric inhibition (Equation (2.4)), by which we mean that the interaction is mediated via separate binding sites and is therefore  non-competitive. For this analysis (Figure 2.4), we focussed  in particular on the current inhibition at pH 6.4 with K+0 concentrations of  zero, 1, 3.5, 5, 10, 20, 80 and 140 mM. The fit  of  these data to [H+]0 (M) Figure 2.3 The concentration dependence of  the inhibition of  Kvl.5 currents by protons in zero (•,©), 5 ( • ) and 140 mM (A) K+0. Data for  zero K+0 were obtained with either 143.5 mM Na+(«) or 143.5 NMG+ (o) as the major extracellular cation. The lines represent the best fit  to equation (2.3). The fitted  values for  the equilibrium dissociation constant for protons CK H),  thepK H  and nH  were, in zero K+0 and 143.5 NMG+: 128 ± 53 nM (mean ± SD), 6.9 and 1.2 ± 0.5; in zero K+0 and 143.5 mMNa+: 153 ± 13 nM, 6.8 and 1.5 ± 0.2; in 5 mM K+0: 590 ± 85 nM, 6.2 and 1.6 ± 0.4; and in 140 mM K+0: 1.1 ± . 11 [iM,  6.0, and 1.8 ± 0.3. Although the pK H  estimates with either Na+ or NMG+ as the extracellular cation are similar, the increase of  the relative gmar with NMG+ at pH 8.4 was significantly  greater. Consistent with a non-competitive versus competitive interaction between H+ 0 and K+0 (see Figure 2.4) the increase of  K H  going from  zero to 5 mM K+0 was greater than that going from  5 mM to 140 mM. Equation (2.4) gave mean values (± SD) of  150 ± 1900 nM for  K H,  1.33 ± 17 for  nH,  0.68 ± 9 for K K  and 6.2 ± 14.9 for  a, the factor  by which bound H7K+ inhibits the binding of  K+/H+. To reduce the SD of  the estimates for  K K  and A, we fixed  the values for  K„  and nH  in the fitting routine at 153 nM and 1.5, respectively, based on the data of  Figure 2.3 (zero K+0, 143.5 Na+0). This was justified  on the basis of  the similarity to the values for  K H  and the nH  from  Figure 2.3 and the preliminary fit  (i.e.,  with the four  parameters free)  of  the data at pH 6.4. With K t, and nH fixed,  the fit  of  the data at pH 6.4 gave estimates for  K K  and A of  0.65 ± 0.27 mM and 5.5 ± 0.7, respectively. 1.2 - | 1 . 0 - pH 6.9 pH 6.4 $ 0.4 - pH 5.9 0.2 0.0-r ~i r "1 :—l 1 1 80 100 120 140 0 20 40 60 [K ] 0 (mM) Figure 2.4 The concentration dependence of  the antagonism by K+0 and Cs+0 of  the inhibition of/iKvl.5  currents by H+0. The relative gm;LX  at different  K+0 concentrations is plotted for  pH 6.9 (•) , pH 6.4 (A) and pH 5.9 ( • ) . The data for  pH 6.9 and 5.4 were obtained with zero, 5, 20 and 140 mM K+0. At pH 6.4, K+0 was zero, 1, 3.5, 5, 10, 20, 80 and 140 mM. Assessment of  the block-relieving effect  of  Cs+0 (v) was done with concentrations of  3.5, 20 and 140 mM. The lines represent the best fit  of  the data to equation 2.4 (see Materials and Methods). With the values for  K H  and nH  fixed  to those obtained directly from  the data in Figure 2.3 (153 nM and 1.5, respectively) the best fit  values for  K K  and arwere 0.65 ± 0.27 mM and 5.5 ± 0.7. Cs+0 appears to be equivalent to K+ 0 in its antagonism of  the proton block. The best fit  of the data at pH 6.9 was obtained with 0.93 ± 2.8 mM for  K K  and 6.2 ± 9.1,for  a, at pH 5.9 the corresponding values were 0.66 ± 0.48 mM and 6.2 ± 1. These estimates for  K K  are very near those estimated for  the K+0 relief  of  the Zn2+ block (-0.5 mM) (Zhang et al., 2001b). At pH 6.9 and pH 5.9 the relative gmax was measured with zero, 5, 20 and 140 mM K+0. For the data at pH 5.9 the best fit  values for  K K  and a, with K„  and nH  constrained as above, were 0.66 ± 0.48 mM and 6.2 ± 1, respectively; at pH 6.9 the corresponding values were 0.93 ± 2.8 mM and 6.2 ±9.1. 2.3.4 External Cs+ ions mimic the block-relieving effect  of  K+ In /zKvl .5 channels the permeability of  Cs+ ions relative to K+ ions is approximately 0.2 (Fedida et al., 1999) and the K D for  the relief  by Cs+0 (K a) of  the Zn2+ block is some 5-6 fold higher than the K k (Zhang et al., 2001b). Surprisingly, with the same experimental protocol but using Cs+ at concentrations of  3.5, 20 and 140 mM (open inverted  triangles  of  Figure 2.4) the ability of  Cs+0 to antagonise the current inhibition by protons was indistinguishable from  that of K+0. 2.3.5 Sensitivity to H+0 and Zn2+0 inhibition is reduced in /rKvl.5 H463Q The range of  pK Hs  for  the inhibition of  hKvl.5  is consistent with the titration of  one or more histidine residues and, as noted above, in rKvl.5 channels in which glutamine (Q) is substituted for  H452, the homologue of  H463 in hKv  1.5, there is a substantially reduced proton sensitivity (Steidl and Yool, 1999). Based on the crystal structure of  KcsA (Figure 2.5; Doyle et al., 1998) H463 is presumed to be located in the outer rim or "turret" of  the pore mouth. Since Zn2+ ions also bind avidly to histidine residues this raised the possibility that the current inhibition caused either by Zn2+ or H+ 0 involves binding to one or more of  the H463s in the turret of  the homotetrameric /zKvl .5 assembly. To test that hypothesis we examined the concentration dependence of  the conductance decrease by H+0 and Zn2+ in the mutant hKvl.  5 H463Q. These experiments were done in zero K+0 so that the interpretation of  the results would not be complicated by a change, if  any, of  the affinity  of  the site at which K+ ions bind to produce an allosteric inhibition of  the actions of  Zn2+ and H+. The g(V)  relationships (Figure 2.6 A) and concentration-response curves (Figure 2.6 B) for the proton sensitivity of  AKvl .5 H463Q, confirmed  the results reported for  rKvl .5. Thus, the gating shift  was apparently intact but the decline of  gmax was seen only with much higher proton concentrations. Fitting of  the concentration-response data (Figure 2.6 B) to the Hill equation Figure 2.5 The structure of  the S5, S6 and the pore (P) loop of  Kvl.5 inferred  from  the crystal structure ofKcsA.  A. The sequence alignment for  Kvl .5, Shaker  and KcsA between the turret and the outer pore mouth. B. A side view of  the KcsA channel in which the foreground  and background a-subunits have been removed for clarity. The a-subunit of  voltage-gated K+ channels has an additional 4 transmembrane segments (S1-S4) that are not illustrated. Sites at which mutations were made, namely H463 and R487, are shown at their homologous positions in the KcsA crystal structure. The orientation of the side chains of  these two residues is tentative. gave an estimate for  K„  of  4.7 ± 1.9 [iM  (pK H  = 5.3) and an nH  of  1 ± 0.4 versus the corresponding values of  0.15 |iM (pK H  = 6.8) and 1.5 in wt /zKvl.5. The acid sensitivity of MCvl .5 H463Q is therefore  quite comparable to that of  rKvl.5 H452Q where the pK H  is -5.2 (Steidland Yool, 1999). Tests of  the effects  of  Zn2+ on the H463Q mutant showed that the outcome (Figure 2.6 C and D) mirrored that seen with protons. Because of  the limited solubility of  Zn(OH)2, the highest concentration of  Zn2+ we tested was 2.5 mM, and consequently a full  concentration-response curve could not be obtained. From the limited concentration range over which data were collected the extrapolated K Zn was 1.7 ± 1 mM or roughly 25-fold  higher than for  wt / j K v I .5 (Zhang et al:, 2001b). The nH  for  the inhibition by Zn2+ of  wt hKvl .5 and / jKvI .5 H463Q currents was 0.9 (Zhang et al., 2001b) and 0.5 (Figure 2.6), respectively. In the course of  this series of  experiments we became aware of  a report that currents A HFSSIPDAFWWAVVTMTTVGYGDMR K v l . 5 FFKSIPDAFWWAVVTMTTVGYGDMT Shaker QLITYPRALWWSVETATTVGYGDLY KcsA B A 12-1.0-N.' 0.8-i x" <0 0.6-e 0.4-0.2-0.0-B 1.2 —, 1.0 1 0.8-1 o> g> 0.6 -1 — 04-1 <0 ^ an 0.2 0.0 H463QH Block c H463Q Zn2+ Block •+*—I 1 r -20 0 20 40 Voltage (mV) 2.5 mM Zn r -20 0 20 40 Voltage (mV) ® e T T T 1.2 1.0 <0 p 0.8 o> 0) > • 0.6 m 0.4 (V Od 0.2 0.0 10"8 10"7 10"* [H10 (M) 1CT i 11 mii|—i 111inij— 0.01 0.1 11 iinij—i 11 mii| 1 10 [Zn2*] mM Figure 2.6 A point mutation in the turret (S5-P loop), H463Q, reduces the inhibition but not the gating shift  caused by H+0 and Zn2+0. A. The g(V)  relationship in zero K+0 at pH 8.4 (•), pH 7.4 (o), pH 6.4 (•) , pH 5.9 ( • ) , and pH 5.5 (T) after  normalization with respect to the gmax at pH 7.4. Values for  the relative gmax, V l/ ;, and 5 were: at pH 8.4, 1.1 ± 0.02, -23.9 ± 1.3 mV and 5.4 ±0.5 mV (n = 5); at pH 7.4, 1,-20.1 ± 1.0 mV and 4.6 ± 0.2 mV(n = 28); at pH 6.4, 1.06 ± 0.03, -13.0 ± 0.5 mV, and 4.0 ± 0.5 mV (n = 3); at pH 5.9, 0.86 ± 0.08, 7.6 ± 0.7 mV, and 5.6 ± 0.4 mV (n = 8); and, at pH 5.5, 0.63 ± 0.04, 19.2 db 1.7 mV and 5.7 ± 0.4 mV (n - 7). B. The concentration dependence of  the reduction of  gmax by protons. Fitting of  the data to the Hill equation gave a K D of  4.7 ± 1.9 nM (pK„  of  5.3) and nH  of  1.0 ± 0.4. The gmax-H+0 concentration relationship for  wt hKvl .5 is represented by the dashed line. C. The g(V)  relationship as described for  (A) but with zero (o), 50 \xM  (•) , 200 \iM(T), 1 mM ( • ) and 2.5 mM (A) of  Zn: The relative gmax, F„„ and 5 were 1, -14.9 ± 0.9 mV, and 4.8 ± 0.3 mV for  the.control (n = 27); 0.87 ± 0.08, -1.9 ± 0.9 mV, and 6.2 ± 0.9 mV for  50 |iM Zn2+ (n  = 5); 0.77 ± 0.09, 7.1 ± 1.1 mV, and 5.9 ± 0.2 mV for  200 \iM Zn2+ (n = 6); 0.47 ± 0.04, 18.5 ± 2.0 mV, and 6.0 ± 0.7mV for 1 mM Zn2+ (n = 10); and 0.52 ± 0.009, 27.2 ± 2.3 mV, and 6.4 ± 0.8mV for  2.5 mM Zn2+ (n  = 5). D. As described for  (B) but with Zn2+. The best fit  values for  K Zn and nH  were 1.7 ± 1 mM and 0.5 ± 0.2. The dashed line indicates the concentration-response relationship for  wt Kvl.5 in zero K+ 0 (K Zn = 69 |iM, nH  = 0.9) (Zhang et al., 2001b). 2+ through / jKvI .5 H463G channels were completely suppressed upon changing from  4.5 mM to zero K+0 medium at pH 7.4 (Jager and Grissmer, 2001). This result was surprising since no such effect  is apparent with the hKvl.5  H463Q mutant under the same recording conditions (Figure 2.6 and 2.7 4^). Our experiments with hKvl .5 H463G confirmed  this conductance collapse in zero K+0 at pH 7.4 (Figure 2.7 C), and we also noted that there was a striking increase in the inactivation rate in 3.5 K+0 (Figure 2.7 B) that was not previously reported. Thus, in contrast to wt AKvl.5 (Figure 2.1) and /zKvl.5 H463Q (Figure 2.7 A) where there is little or no current decay evident during 300 ms pulse commands, in /iKvl .5 H463G the current decay at 40 mV is well-fitted  by a single exponential function  with a time constant of  73 ± 8 ms (n  = 4) (Figure 2.7 B). H463Q, 0 mM K* 2 nA B 2 nA 100 ms H463G, 3.5 mM K* 100 ms H463G, 0 mM K* 2 nA 100 ms Figure 2.7 In AKvl.5 H463G slow inactivation is greatly accelerated and the conductance collapses in zero K+0 at pH 7.4. A. Shown for  comparison are the currents from  hK\\  .5 H463Q evoked in zero K+0 by 300 ms pulses to between -40 and 40 mV in 10 mV increments. B. /zKvl .5 H463G currents recorded using the same stimulus protocol but with 3.5 mM K+0. The solid line superimposed on the current at 40 mV represents the best fit  of  the current decay to a single exponential function.  The mean time constant for  inactivation at 40 mV was 73 ± 8 ms {n = 4). C. From the same cell as in B and using the same voltage command protocol after  switching to zero K+ 0 at pH 7.4. Unlike either wt Kvl.5 H463 (Figure 2.1) or Kvl,5 H463Q, K+0 is required for  /zKvl .5 H463G channels to function  normally at pH 7.4. Complete recovery was obtained after  returning to K+-containing bath solution (not shown). 2.3.6 H+0 and Zn2+0 accelerate inactivation In rKvl .5, H+ 0 has been shown to accelerate inactivation, an effect  that was evident only with long depolarizing commands (Steidl and Yool, 1999). Similarly, in /zKvl.5 there was no. obvious change of  inactivation kinetics during 300 ms depolarizations but an increased inactivation rate was evident with H+0 as well as Zn2+ during depolarizations lasting for  several seconds (not shown). Fitting a single exponential function  to the current decay during a 7 to 10 s depolarization at 60 mV in external medium with 5 mM K+ at pH 7.4 gave a time constant for inactivation (r inac l) of  2.63 ± 0.11 s (n = 4).' In the same cells, extracellular acidification  to pH 6.4 caused a roughly 50% reduction of  rmact to 1.19 ± 0.04 s (p  < 0.05). Using the identical stimulation protocol, we found  that the changeover from  Zn2+-free  medium at pH 7.4 to medium at the same pH and containing 1 mM Zn2+ reduced r m n c t by approximately 30% from  3.0 ± 0.18 s to 2.14 ± 0.16 s {n  = 4,/? < 0.05). Although these results confirm  that current inhibition by H+0 and Zn2+ is associated with a moderately increased rate of  inactivation we suggest below that this cannot account for  the reduction of  gmax. 2.3.7 Current inhibition by protons and Zn2+0 is reduced in AKvl.5 R487V To more directly address the possibility that the reduction of  gmux reflected  an effect  on one or more inactivation processes we next examined the actions of  H+0and Zn2+ in a /?Kvl .5 mutant in which an arginine (R) residue in the P-S6 region was mutated to valine (V) (R487V, Figure 2.5). This was motivated by the fact  that mutations at the homologous site (T449) in N-type (fast)  z'nactivation- removed Shaker  channels (ShakerlR)  either accelerates (T449E, T449K, T449A) or slows (T449Y, T449V) inactivation (Lopez-Barneo et al., 1993). A previous study.of /zKvl.5 R487V showed that inactivation was indeed dramatically slowed when channel currents were carried by Na+ but, curiously, the time course of  K+ currents were relatively unchanged (Wang et al., 2000). It has also been proposed that a charged residue at position 487 is critical for  the current inhibition by HH0 (Jager and Grissmer, 2001). Figure 2.8 summarizes the results of  experiments assessing the inhibition of  hKvl  .5 R487V by protons and Zn2+ ions in zero K+0(143.5 mM Na+) medium. The g(V)  relationships derived from  tail current measurements (Figure 2.8 A) show that the gating shift  was apparently intact in the R487V mutant. However, there was a dramatic change of  the concentration dependence of  the H+0-induced conductance decline. For example, whereas in wt hKvl.5  the relative gmcu at pH 5.9 was 0.07 ± 0.01 (n = 9; Figure 2.4), in MCvl .5 R487V the relative gmax at the same pH was 0.92 ± 0.03 (n = 5; Figure 2.8 A and B). An extrapolatedpK H  of  4.6 obtained from  the best fit  of  the concentration-response data of  Figure 2.8 B suggests a shift  of  ~2 pH units from  the pK H  of  wt hKvl  .5 channels. Tests of  the sensitivity of  MCvl.5 R487V channels to Zn2+ (Figure 2.8 C and D) showed that the gating shift  was, again, substantially unaffected  and, as with H+0, there was a clear increase of  the Zn2+ concentration required to cause 50% inhibition. Thus, whereas wt hKvl  .5 currents were half-inhibited  by 0.07 mM Zn2+ (Zhang et al., 2001b), in the R487V mutant 41.2 ± 1.7% {n  = 5) of  gmax persisted in 2.5 mM Zn2+. Closer inspection of  the concentration-response data of  Figure 2.8 D suggested that two Zn2+ binding sites might be involved in the inhibition of hKvl  .5 R487V currents. Subsequent experiments with the double mutant hKvl  .5 H463Q, R487V (open triangles  of  Figure 2.8 D) implied that the higher affinity  site (K Zn = 29 ^M) which accounted for  approximately 20% of  conductance decline in the R487V mutant, was apparently eliminated. The latter observation could be accounted for  in many ways, perhaps the simplest being that the higher affinity  site in the R487V mutant reflects  the binding of  Zn2+ to one or more R487V H* Block R487V Zn2* Block 2 5 mM Zn 24 (Hlo (M) [Zn 2*] (mM) Figure 2.8 A mutation near the pore mouth, R487V, substantially reduces the sensitivity to inhibition by H+0 and Zn 2\. A. The g(V)  relationship in zero K+0 at pH 8.4 ( • ) , pH 7.4 (O), pH 6-4 ( • ) , pH 5.9 (A), and pH 5.5 ( • ) after  normalization with respect to gmwc at pH 7.4. The values for  the relative gmax, V 1/ 2, and s were, respectively, 1.04 ± 0.02, -28.5 ±1.1 mV, 4.6 ± 1.0 mV at pH 8.4 (n  = 3); 1,-18.1 ± 0.9 mV, 4.5 ± 0.2 mVatpH7.4 (n=  17); 1.04 ± 0.06,-1.8 ± 1.3 mV, 5.6 ± 0.4 mV at pH 6.4 (n  = 5); 0.92 ± 0.03, 6.4 ± 1.3, 4.9 ± 0.4 at pH 5.9; and, 0.87 ± 0.03, 15.5 ± 1.6 mV, 5.5 ± 0.2 mV at pH 5.5 (n = 5). B. The concentration-response relationship for  the reduction of  gmax by protons. The continuous line, representing the best fit  of  the data to the Hill equation, was obtained with K H  = 23 |iM (pK H  of  4.6) and nH  = 0.8. C. The g(V) relationship in zero K+0 and with Zn2+ concentrations of  10 |j,M (•), 25 (iM (•), 100 |iM (0), 200 \iM  (T), 1 mM (•) , and 2.5 mM after  normalization with respect to the control (O) gmax. The relative gmax, V i2, and .r were, respectively, 1, -13.4 ± 1.5 mV, 4.5 ± 0.3 mV for  the control^? = 15), 0.99 ± 0.01, -5.9 ± 1.5 mV, 5.4 ± 0.4 mV in 10 |iM Zn2+ (n = 4); 0.92 ± 0.05, -5.7 ± 0.1 mV, 4.7 ± 0.5 mV in 25 [iM Zn2+ (n = 3); 0.80 ± 0.02, 2.8 ± 1.8 mV, 4.8 ± 0.5 mV in 100 [iM Zn2+ (n = 4); 0.78 ± 0.02, 5.2 ± 1.6 mV, 5.3 ± 0.3 mV in 200 |iM Zn2+ (n = 5); 0.70 ± 0.05, 21.0 ± 1.2 mV, 5.9 ± 0.3 mV in 1 mM Zn2+ (n = 3); and, 0.59 ± 0.02, 28.9 ± 1.1 mV, 5.9 ± 0.3 mV in 2.5 mM Zn2+ (n  = 5). D. As described for  (B) but with Zn2+. The continuous line represents the best fit  of  the h Kvl .5 R487V data to the sum of  two Hill equations. Binding at the higher affinity site (K Zn = 29 ± 0.2 p,M) accounted for  -20% of  the inhibition. The apparent elimination of  the higher affinity  site in the double mutant Kvl.5 R487V, H463Q (A and dashed  line) suggests that it may reflect  Zn2+ binding to H463. The extrapolated K Zn for  the lower affinity  site in the R487V mutant was 6.4 ± 0.07 mM. Again, the dotted lines in B and D represent the corresponding concentration-response curves for  wt hKvl.5 (Zhang et al., 2001b). H463 residues. The concentration dependence of  the reduction of  gmax in the double mutant was best fitted  by a single Hill function  with a K Zn of  2.2 mM, representing an approximately 30-fold increase over that measured in wt /?Kvl .5 under the same recording conditions. 2.3.8 Current inhibition by H+0 and Zn2+0 is apparently not use-dependent If,  as has been proposed to account for  the block of  rKvl .5 currents by H+ 0 (Steidl and Yool, 1999), the inhibition of  hKvl  .5 currents by Zn2+ or extracellular acidification  were due to an accumulation of  inactivation, then the degree of  inhibition would be expected to show use-dependence. Figure 2.9 shows the results of  a representative experiment addressing this issue. Peak tail current amplitudes following  300 ms depolarizations from  -80 mV to 60 mV at pH 5.9 are bracketed by control and recovery responses at pH 7.4. K+0 was 3.5 mM. Two features  of  the current behaviour at pH 5.9 are significant.  First, inhibition of  the current is apparent with the first  pulse and is more-or-less constant for  each of  the subsequent pulses during a train of  10 pulses delivered at 5 s intervals. Second, a 2 min stimulus-free  interval in which the membrane was held at either -80 mV or -100 mV had no block-relieving effect.  Consequently, despite the fact  that both Zn2+ and H+0 slightly enhance the rate of  inactivation of  residual hKvl.5 currents, there is no support for  the hypothesis that accumulation of  inactivation accounts for  the reduction of  gmax. Finally, Figure 2.9 also demonstrates the rapid reversal, i.e., within the time course of fluid  exchange in the bath, of  the current inhibition after  beginning the perfusion  with pH 7.4 solution. The latter observation argues against a mechanism involving a change of  the internal pH concomitant to extracellular acidification. 1000 < (1) 800 — -o Q. 600-E TO C 400-<D t D 200-U (0 pH 5.9 Vholding = -100 mV V 200 I 400 I 800 I— 1000 1200 600 Time (s) Figure 2.9 The effect  of  the stimulus frequency  and holding potential on the inhibition of wt  hKvl.5 currents by H+0. This graph, which is representative of  the results obtained from  six such experiments, three at pH 5.9 and three with 1 mM Zn2+, plots the amplitude of  tail currents measured at -50 mV following  a 300 ms step to 60 mV to maximally activate channels. After  10 consecutive control responses in standard external saline (pH 7.4, 3.5 mM K+0) and evoked at 5 s intervals from  a holding potential of  -80 mV, pulsing was stopped and 5 ml of  test solution was perfused  to change the extracellular pH to 5.9 for  the duration indicated by arrows. Resumption of  the step commands approximately 2 minutes after  extracellular acidification  showed an immediate -75% reduction of  the tail current amplitude. The identical effect  was obtained for each of  two subsequent pulse trains confirming  that the inhibition was not affected  by a period without stimulation. Changing the holding potential to -100 mV also had no effect  on the current amplitude. Returning to pH 7.4 medium while pulsing shows the effect  rapidly (within 15 s) and completely reverses implying that a change of  the internal pH is not involved. 2.3.9 Protons cause a depolarizing shift  of  the Q(V) relationships but do not affect  Q ^ Because gating currents can provide useful  evidence on the conformational  states available to a channel, we recorded gating currents in a stable HEK-293 cell line expressing AKvl.5 W472F mutant channels (Chen et al., 1997). This mutant is analogous to the Shaker W434F non-conducting mutant in that it has no measurable K+ current, perhaps because of permanent or greatly accelerated P-type inactivation (Yang et al., 1997). Representative examples of  gating current traces from  /zKvl.5 W472F recorded at pH 7.4 and pH 5.4 in the same cell are shown in Figure 2.10 A-D. To prevent contamination of  gating currents by endogenous HEK-293 ionic currents, these recordings were made in symmetrical 140 mM NMG+. At pH 7.4, on-gating currents were evoked between -60 and 100 mV from  a holding potential of  -100 mV and at pH 5.4 the voltage range was from  -60 to 150 mV to compensate for  the proton-induced gating shift.  As reported previously (Chen et al., 1997), on-gating currents at pH 7.4 were first  apparent at -60 mV and as the strength of  the depolarization increased both the peak amplitude and decay rate of  the on-gating current increased. Following depolarizations up to 0 mV the return- or off-gating  currents decayed rapidly as channels deactivated at -100 mV. In contrast, following  depolarizations to 0 mV or more the off-gating currents are superimposable and have a clear rising phase that is followed  by a slow decay. This slowing of  charge return is such that integration of  the off-gating  current over a 15 ms period produces a Qof f  that is reduced relative to Qon. This decrease of  Qaf/Q on or charge immobilization has been attributed to the conformational  change underlying C-type inactivation (Chen et al., 1997; Yellen, 1997) since it is affected  by the presence-of  permeant metal cations, much as C-type inactivation of  ionic currents is affected  by extracellular cations (Baukrowitz and Yellen, 1995; Lopez-Barneo et al., 1993). C-type inactivation is greatly accelerated in the recording conditions used here because there are no permeant metal cations on either side of  the membrane. The transition of  the voltage sensor from  its outward "immobilized" position to the inward position remains voltage dependent but stronger hyperpolarizations are required to overcome the stabilizing interaction between the sensor and the C-type inactivated state. This accounts for  the leftward  shift,  relative to the Qon(V)  relationship, of  the voltage dependence of  charge return (Olcese et al., 1997; Wang and Fedida, 2001; see Figure 2.10 E). Figure 2.10 B shows that changing the external pH from  7.4 to 5.4 caused a rightward shift  of  the voltage dependence of  the on-gating currents such that the on-gating current evoked at 150 mV at pH 5.4 was comparable to that at 100 mV at pH 7.4. At pH 5.4 there was also a A __ B 100 mV < c D 5 ms -160 mV -200 mV O Q. s 1 O 1 -0-o -1 a. 5 - 2 H O Q  pH 7.4 • CLPH5.4 QOQOujf -200 -50 Voltage (mV) Figure 2.10 Extracellular acidification  to pH 5.4 causes a depolarizing shift  of  the Q0„(V) and Q0jj(V)  relationships but does not reduce Q^.  Panels^ and B show at pH 7.4 and 5.4, respectively, the on- and off-gating  currents recorded when the membrane was depolarized for  12 ms from  a holding potential of  -100 mV to between -60 and 100 mV (A)  or -60 and 150 mV (B)  in 10 mV increments before  stepping back to -100 mV. Outward charge movement (Q on) induced by the depolarization was determined by integrating the on-gating currents at pH 7.4 (O) and 5.4 (•), and is plotted in panel E. For the Qon(V)  relationship in E, the fitted  values for  V l2 and s were, respectively, -2.2 mV and 6.5 mV at pH 7.4 and 50.2 mV and 11.8 mV at pH 5.4. Q ^ was not significantly affected  by extracellular acidification.  C, D. From the same cell as in A and B, these panels show the off-gating  currents following  a 12 ms step from  -80 mV to 50 mV in pH 7.4 (Q or to 100 mV at pH 5.4 (D) to move QmiLX.  Off-gating  current was recorded in 10 mV increments between -200 and -10 mV at pH 7.4 and between -200 and 40 mV at pH 5.4. Charge return at pH 7.4 (A) and pH 5.4 (A) is plotted against the repolarization voltage in E to obtain the Qo//V)  relationship. Extracellular acidification  changed the V r2 of  Qof j(V)  from  -100.5 mV to -72.9 and 5 increased from  9.4 mV to 13.1 mV. Both at pH 7.4 and pH 5.4 there is a leftward  shift  of  the voitage dependence of  Qoff relative to Qnn(V). substantial increase of  the peak amplitude and an increase of  the decay rate of  offgating  currents following  strong depolarizations. To quantify  the effects  of  changes of  pH0 on activation gating, the on-gating currents in Figure 2.10 A and B were integrated to obtain the voltage dependence of on-gating charge movement shown in Figure 2.10 E. A fit  of  the Qon(V)  relationship at pH 7.4 (open  circles) to a single Boltzmann function  gave a maximum charge movement Qmax of  / +2.5 pC , V,,  = -2.2 mV and 5 = 6.5 mV. At pH 5.4 {filled  circles  of  Figure 2.10 E), Qmax, V,,  and r 5 were +2.5 pC, 50.2 mV and 11.8 mV. In the six cells examined, V V i was 4.3 ± 2.2 mV at pH 7.4 and 48.9 ± 1.2 raV at pH 5.4• s increased from  7.1 ± 0.5 mV at pH 7.4 to 10.5 ± 0.4 mV at pH 5.4; and, the relative Qmax (Q max,pHJQ mas,pH?.,)  was 1.0 ± 0.003. Thus, changing pH0 from  7.4 to 5.4 caused a -45 mV rightward shift  of  the V,,  of  the Qon(V)  relationship and a decrease of  the voltage-sensitivity of  activation. Both of  these effects  are replicated by Zn2+ (Zhang et al., 2001a) and interestingly, as with Zn2+, the shift  of  V,- : of  the Qon(V)  relationship is roughly twice that measured for  the g(V)  curve. For example, at pH 5.9 the K,  of  the g(V)  relationship was shifted  by -21 mV (not shown). Panels C and D of  Figure 2.10 illustrate the outcome of  experiments to determine if  the change of  off-  gating current in Figure 2.10 B was due to a shift  of  the voltage-dependence of  off-gating charge movement (Olcese et al., 1997). The voltage clamp protocol consisted of  a 12 ms step from  the holding potential of-80  mV to 50 mV at pH 7.4 (C) or 100 mV at pH 5.4 (D)  to evoke maximal charge movement. This was followed  immediately by a pulse to between -10 and -200 mV at pH 7.4 or to between 40 and -200 at pH 5.4. Integration of  the o^-gating currents yielded the Qof/V)  curves shown in Figure 2.10 E at pH 7.4 (open triangles)  and pH 5.4 (closed triangles).  Considering first  the data at pH 7.4, it can be seen that, as in the Shaker  non-conducting mutant (Olcese et al., 1997), the voltage dependence of  return charge movement was shifted  leftward  (V T I  = -100.5 mV) by -100 mV relative to the QON(V)  curve. Of  particular importance is that a similar effect  is seen at pH 5.4 where the V,, : of  the QOJ/V)  curve was -72.9 mV, representing a leftward  shift  of  - 124 mV from  the V,,  of  the QON(V)  relationship. The values, respectively, for  V,,,  and 5 of  the QOF/V)  relationship in 3 such experiments were, at pH 7.4,-102.8 ± 1.4 mV and 11.4 ± 1.0 mV and, at pH 5.4, -75 .3 ± 1.4 mV and 14.8 ± 1.1 mV. Thus, at pH 7.4, there was, following  a depolarization that moved Qm(XX, a -107 mV leftward  shift  of  the voltage dependence of  return gating charge movement. A comparable leftward  shift  of  -124 mV of  the voltage dependence of  gating charge movement was seen at pH 5.4. 2.4 Discussion The first  series of  experiments (Figures 2.1 - 2.4) described in this paper show that, as with Zn2+ ions, external protons cause a concentration-dependent and reversible inhibition of hKvl  .5 currents. Although this effect  is associated with a depolarizing shift  of  the activation (g(V))  curve, the two actions appear to be mechanistically unrelated. Both effects  have been reported for  rKvl .5 channels (Steidl and Yool, 1999) but we have extended the previous work by showing that external ions such as K+ and Cs+, but not Na+, are able to relieve the inhibition but not the gating shift  caused by protons. In zero K+0 the apparent pK H  of  the protonation site is 6.8 and this decreases to 6.2 with 5 mM K+0. The latter pK H  accords well with the pK H  of  6.2 for rKvl.5 responses recorded in 2 mM K+0 (Steidl and Yool, 1999) and thepK H  of  6.1 in N-terminal deleted ferret  Kvl.4 with 3 mM K+0 (Claydon et al., 2000). The influence  of  K+0 on this inhibition of  /zKvl.5 currents was modelled as a non-competitive interaction between K+ and. protons (Figure 2.4) and the estimated K D for  this antagonism by K+0 is very near that estimated for  the Zn2+ block (Zhang et al., 2001b), i.e., K K  = 0.5 -1.0 mM. This implies that the same K+ binding site is involved in both cases and is perhaps homologous to the site (K D -0.75 mM) at which K+ binds to lock Ba2+ ions within the pore of  ShakerB  channels (Harris et al., 1998). Binding sites with a similar affinity  for  K+ have also been shown to influence  the availability of Shaker  T449A channels (K D = 0.8 mM) (Lopez-Barneo et al., 1993) and to competitively inhibit C-type inactivation in ShakerlR  channels (K D= 1 - 2 mM) (Baukrowitz and Yellen, 1996). Interestingly, the external lock-in site of  ShakerB  channels and the site at which K+ binds to antagonise the inhibitory actions of  Zn2+ or H+0 in MCvl.5 also share the property of  having a low affinity  for  Na+ ions. One of  two clear differences  between the actions of  H+0 and Zn2+ is that while the K D for the relief  by Cs+0 of  the Zn2+ block is roughly 5-fold  higher than that for  K+0 (Zhang et al., 2001b), Cs+0 is as effective  as K+0 in antagonizing the current inhibition by protons (Figure 2.4). In the case of  Zn2+, the higher K 0 for  Cs+ was assumed to reflect  the lower permeability of  the Cs+ in the pore. With H+ 0 it is conceivable that protonation of  a negatively-charged, cation-binding site decreases that site's negativity and alters the selectivity sequence to one favouring  Cs+ binding (Hille, 2001). If  so, the selectivity sequence of  a binding site in the outer pore mouth must be involved since we have no evidence of  a change of  the reversal potential with extracellular acidification.  The nH  of  ~ 1.5 for  the proton block suggests that at least two sites, most likely H463 residues in the tetrameric channel assembly, must be protonated. Although the nH  for  Zn2+ block is near unity (Zhang et al., 2001b), this might still involve coordinated binding of  histidine residues of  two or more subunits. 2.4.1 Evidence against a pore blocking mechanism The block of  cardiac voltage-gated Na+ (Nav) channels by Zn2+ occurs by occlusion and is eliminated by the mutation of  a cysteine residue in the pore (Backx et al., 1992). Similarly, the block by external protons of  Nav channels in nerve and skeletal muscle has a voltage-dependence suggesting a site of  action within the pore (Woodhull, 1973). It seems unlikely, however, that the inhibition of  /?Kvl.5 current by H+0 and Zn2+ is due to pore block. First, in the voltage range where the open probability is maximal there is no indication of  a voltage-dependent decline of the inhibition by Zn2+ (Zhang et al., 2001b) or H+0, e.g. Figure 2.1 C, as would be expected were these ions binding at a pore site within the electric field.  The latter observation is consistent, however, with an interaction with one or more H463 residues which, being in the channel's turret, are outside the electric field.  It is well-established that Zn2+ and H+ bind to histidine residues, and we have shown directly that the H463Q substitution shifts  the pK H  measured in zero K+0 from  6.8 to approximately 5.4 (Figure 2.6). The acid sensitivity that persists in this mutant and in hKvl.5  R487V (Figure 2.8) is similar to that reported for  Shaker  (pK H  -5.4) (Perez-Cornejo et al., 1998) and Kvl.2 channels (pK H  -4.9) (Ishii et al., 2001). Exactly where protons and Zn2+ act in these mutants is not known, but given their typical pK H  values of  4-5, likely candidates are the aspartate residues in the outer pore mouth (see Introduction). Since each of  the H463s is approximately 14-16 A from  the central axis of  the pore (Aiyar et al., 1995; Doyle et al., 1998), it is very unlikely that binding of  either ion to H463 residues would directly occlude the permeation pathway since Zn2+ has an ionic radius of  0.74 A and H+ is orders of  magnitude smaller. Assuming that the site at which external K+ binds to antagonise the current inhibition by Zn2+ and H+0 is in the outer pore mouth, our observation that this interaction is best described by a non-competitive versus a competitive model of  inhibition also argues against direct pore block as a mechanism of  action of  either cation. It appears therefore  that protonation or "zincification"  of  H463 residues indirectly leads to current inhibition. From this view of  H463 as sensor arises the next question: what is the nature of  the effector? 2.4.2 A connection between current inhibition and an inactivation process Though it is clear from  this study and that of  rKvl.5 currents (Steidl and Yool, 1999) that inactivation is faster  at acidic pHs, our simulation studies (not shown) indicate that this increased rate of  inactivation cannot itself  account for  the reduction of  gmax. Furthermore, although increasing K+0 can speed recovery from  C-type inactivation of  Kvl.3 currents (Levy and Deutsch, 1996), an explanation for  the inhibition that involves a slowing of  recovery from  inactivation and an accumulation of  inactivation can be rejected since a two minute period without voltage pulsing has no effect  on the degree of  inhibition (Figure 2.9). Nonetheless, a simple interpretation of  the effect  of  K+0 (or Cs+0) on the reduction of  gmax caused by H+ 0 or Zn2+ is that, by a "foot-in-the  door" mechanism, K+0 acts as a competitive antagonist of  a conformational change at the pore mouth that is believed to underlie inactivation. In this connection we think it is significant  that a point mutation at a site (position 487; T449 in Shaker)  which has been implicated in the regulation of  inactivation (Lopez-Barneo et al, 1993) dramatically affects  the proton block (Figure 2.8). In the studies of  mutant Shaker  channels the terms "potentiation'V'conductance collapse" were used to describe the increase/decrease of  gmax when K+ 0 was increased/decreased. It is likely that potentiation/collapse in ShakerlR  is analogous to block relief/block  in hKv\  .5 but there are some differences.  Foremost among these is that, in contrast to the Shaker  mutants, in Kvl .5 the block (conductance collapse) is K+0 and  pH sensitive. That is to say at pH 7.4 removing K+0 has little or no effect  on wt /iKvl .5 currents whereas at pH 6.4 decreasing K+0 causes a substantial conductance decline. Additionally, although the tendency for  the conductance of  Shaker  mutants to collapse in zero K\  is strongly correlated with an accelerated inactivation rate, this does not extend to hiCvl.5 where wt hKvl.5 is much more prone to block at pH 6.4 than is MCvl.5 R487V even though both inactivate at approximately the same rate at pH 7.4 (Fedida et al., 1999). Nonetheless, the fact  that current inhibition by H+0is substantially reduced by increasing K+0 or by the R487V mutation implies that an inactivation process is involved. Additional insight into the possible basis for  the proton block and in particular about the role of  C-type inactivation was provided by gating current results (Figure 2.10). 2.4.3 External acidification  and on-gating charge movement Based on the data of  Figure 2.10, we can immediately exclude a mechanism of  action in which protonation of  H463 residues impedes on-gating charge movement and consequently prevents the opening of  the activation gate since at pH 5.4 there is no significant  reduction of Qmax- This is a second major distinction between the actions of  H+0 and Zn2+. At a concentration that reduces gmax by more than 90%, Zn2+ decreases Qmax by 10-15% as though it were preventing the late, weakly voltage-dependent transitions in the activation pathway (Zhang et al., 2001a). However, even though Qmax is unchanged by extracellular acidification,  we cannot rule out the possibility that opening of  the activation gate becomes uncoupled from  the outward movement of the voltage sensor. Interestingly, as with Zn2+ (Zhang et al., 2001a), the proton-induced depolarizing shift  of  for  the Qm(V)  relationship is roughly 2-fold  greater than that measured from  the ^ ( ^ relationship. We have attributed this differential  effect  on the g(V)  and QfV) curves to the presence of  two distinct binding sites. In our view the protonation of  an as yet unidentified  site on the channel surface  affects  the movement of  the voltage sensor and culminates in a rightward shift  both of  the g(V)  and the Q(V)  curves. The protonation of  a second site, which is probably H463, has two direct or indirect effects:  it decreases gmax and it causes a rightward shift  of  the voltage dependence of  activation gating. The latter effect  possibly reflects the close proximity of  S4 and H463 in the S5-P loop (Loots and Isacoff,  2000). Since H463-protonated and therefore  non-conducting channels can report the gating shift  in gating current measurements, but not in ionic current measurements, the gating shift  attributed to protonation of this second site is evident only in the Q(V)  curve. 2.4.4 External acidification  and off-g&tmg  charge movement At pH 7.4 and pH 5.4 the mid-point of  the Qof/V)  relationship was shifted  leftward, relative to the corresponding Qon(V)  curve, by 107 mV and 124 mV, respectively (Figure 2.10). That this shift  occurs at both pHs is significant  because it has been attributed to a stabilization of S4s in the outward position by a conformational  change linked to C-type inactivation (Olcese et al., 1997; Wang and Fedida, 2001). We take this to mean that at pH 5.4 channels are not C-type inactivated prior to a depolarizing pulse but are able to become so when sufficiently  depolarized. In other words, at a low pH wt /jKvI .5 apparently behaves like the non-conducting mutant /zKvl.5 W472F. That is to say, the gating shift  notwithstanding, on- and off-gating  charge movement is relatively normal but the channels are never or, at best, only very briefly  in a conducting state. We speculate, as proposed for  homotetrameric ShakerlR  W434F (Yang et al., 1997), that in wt /zKvl .5 protonation of,  or Zn2+ binding to, H463s allows an inactivation process to occur either from  one or more of  the closed states or at a greatly accelerated rate following  the outward movement of  the voltage sensor and channel opening. Since our data indicate that the transition to the C-type inactivated state is intact even at low pHs, this leaves P-type inactivation as a possible basis for  the H+0-induced current inhibition. Some support for  this suggestion comes from  a study of  the ShakerlR  FWFW mutant (Yang et al, 1997) where, as described for P-type inactivation (De Biasi et al., 1993), peak FWFW current was increased by external TEA+ and where there was also an enhancement of  FWFW current when K+0 increased. In a similar manner, in MCvl .5 the block-relief  by K+0 would be due to the occupancy of  a site, presumably near the outer pore, that inhibits P-type inactivation. The cooperativity factor,  a, of  equation (2.4) would then be interpreted to mean that protonation of  H463s, by virtue of  a conformational change, inhibits the binding of  K+ at its site, and vice versa. This proposed scheme, which remains to be tested by single channel analysis, is at least functionally  equivalent to closed-state inactivation proposed to account for  the loss of  current in ShakerlR  T449 mutants (Lopez-Barneo et al., 1993), to the decrease of  channel availability (N) proposed for  the current loss in zero K+0 in Kvl.4 (Pardo et al, 1992), and to the non-conducting "open" state proposed for  Kvl.3 (Jager et al, 1998) and MCvl.5 (Wang et al., 2000) channels. 2.4.5 What is the connection between H463 and R487? To reiterate, our view is that H463 acts as a sensor and R487 is a required component in the effector  mechanism, e.g., inactivation. Concerning the nature of  the coupling between H463 and R487, it has been proposed that the charge of  H463 reduces the pK H  of  R487 by an electrostatic effect  (Jager and Grissmer, 2001). However, a number of  our observations argue against such an electrostatic interaction. First,.a strong, mutual  electrostatic interaction between R487 and H463 would be expected to affect  the pK H  of  H463. In this connection, a histidine residue substituted at the same position in the turret of  Shaker  channels (F425H) has a pK H  of  6.4 in 2 mM K+0 (Perez-Cornejo et al., 1998) that is similar to that for  wt AKvl.5 (pK H  ~6.2 in 5 mM K+0, Figure 2.3). This suggests that the pK H  of  a histidine in the turret is weakly influenced,  if  at all, by the nature of  the residue apposed to it in the tertiary structure (Doyle et al., 1998), be it either charged as with R487 in hKvl .5, or polar and uncharged as with T449 in Shaker.  This also implies that an effect  of  the R487Y mutation on the binding equilibrium for  H+ or Zn2+ at H463 does not account for  the decreased sensitivity of  /zKvl.5 R487V currents to inhibition by either cation. Another argument against an electrostatic interaction between a protonated H463 and R487 is that the proposed shift  of  the pK H  of  R487 by ~6 units would require that these two residues be in much closer apposition (Elinder et al., 2001) than the 8 A (Ca to Ca) suggested by the crystal structure of  KcsA (Doyle et al., 1998).. We have also found,  contrary to the expectation of  an electrostatic mechanism, that increasing the Debye length by decreasing the ionic strength of  the external solution does not substantially affect  the block of  wt ZKvl.5 by Zn2+ (Minshall & Kehl, unpublished data). On these grounds, an electrostatic interaction between protonated H463 and R487 seems unlikely but the pH sensitivity of  /zKvl .5 R487H (Jager and Grissmer, 2001) does imply that a positive charge near the pore mouth is necessary for  the virtually complete suppression of  outward current seen in zero K+0. An alternative view of  the coupling between H463 and R487 is that, perhaps because of the change of  its charge and a consequent increase of  its hydrophilicity, the protonation of  H463 permits a conformational  change requiring R487. Although we have no direct evidence for  such a conformational  change, it is intriguing that studies of  Kv2.1 have shown that the distribution of channels between two outer vestibule conformations  is regulated by K+0 (Immke et al., 1999). Additionally, a lysine residue (K356) which is positively charged at neutral pH, and which is homologous in position and charge to a protonated H463 of  MCvl .5, is crucial in this K+-dependent conformational  change (Immke et al., 1999). The K356 residue is also involved in the enhancement of  Kv2.1 currents by K+0 (Wood and Korn, 2000). 2.4.6 Inactivation and the influence  of  the charge on and size of  the residue at position 463 Jager & Grissmer (2001) recently reported, and we have confirmed  here, that in the mutant hKv\  .5 H463G the conductance collapses at pH 7.4 after  switching to zero K+0 (Figure 2.7). We also found  that this mutant inactivates much faster  than wt /zKvl .5 which underscores the association in Shaker,  noted above, between an increased inactivation rate and a tendency for the current to collapse in zero K+0. The differences  in the properties of  wt /zKvl .5 and the H463Q and H463G mutants also imply that both the charge on and the size of  the residue at position 463 influence  the structural rearrangement leading to a conductance collapse in zero K+0. The importance of  charge is evident in wt hKvl  .5 at pHs where, when H463 residues are protonated, the conductance collapses in zero K+0. An influence  of  the size of  the residue at position 463 is suggested by different  behaviours of  the H463Q and H463G mutants. Thus, there is no conductance collapse in zero K+0at pH 7.4 in /JKVI.5 H463Q where the substituted glutamine is uncharged, but polar, and occupies only a slightly smaller volume than histidine (~ 150 A3). In contrast, in the H463G mutant the uncharged but much smaller glycine residue (-60 A3) does allow the conductance to collapse in zero K+0 at pH 7.4. Additional indirect support for  the idea that the size of  the residue at this position in the turret affects  inactivation comes from  the report that substitution of  glutamine for  glycine at the homologous position in Kvl.3 (G380Q) slows inactivation roughly 7-fold  (Nguyen et al., 1996). Finally, the results of  voltage clamp fluorimetry  in Shaker  suggest that, rather than being restricted to a structural collapse at the selectivity filter,  slow inactivation may involve a coordinated movement extending to the outer rim (turret) of  the pore (Loots and Isacoff,  2000). This is consistent with our results that in hKvl.5  both the charge on and the volume of  the residue at position 463 in the turret influences  a coordinated movement to an inactivated state. Acknowledgements Supported by a grant to S.J.K. from  the Natural Sciences and Engineering Research Council and by grants to D.F. from  the Canadian Institutes for  Health Research and the Heart and Stroke Foundation (HSF) of  British Columbia and Yukon. S.Z. was in receipt of  a Research Fellowship from  the HSF of  Canada and D.C.H.K. was supported by a Trainee Award from  the Michael Smith Foundation for  Health Research. We thank Qin Wang who prepared the cells and Dr. Simon Baudrexel who assisted with some of  the experiments. 2.5 References Aiyar, J., J. M. Withka, J. P. Rizzi, D. H. Singleton, G. C. Andrews, W. Lin, J. Boyd, D. C. Hanson, M. Simon, and B. Dethlefs.  1995. Topology of  the pore-region of  a K+ channel revealed by the NMR-derived structures of  scorpion toxins. Neuron 15:1169-1181. Backx, P. H., D. T. Yue, J. H. Lawrence, E. Marban, and G. F. Tomaselli. 1992. Molecular localization of  an ion-binding site within the pore of  mammalian sodium channels. Science 257:248-251. Baukrowitz, T. and G. Yellen. 1995. Modulation of  K+ current by frequency  and external [K+]: a tale of  two inactivation mechanisms. Neuron 15:951-960. Baukrowitz, T. and G. Yellen. 1996. Two functionally  distinct subsites for  the binding of internal blockers to the pore of  voltage-activated K+ channels. Proc. Natl. Acad. Sci. U. S. A. 93:13357-13361. Blaustein, R. O., P. A. Cole, C. Williams, and C. Miller. 2000. Tethered blockers as molecular 'tape measures' for  a voltage-gated K+ channel. Nat. Struct. Biol. 7:309-311. Cha, A. and F. Bezanilla. 1998. Structural implications of  fluorescence  quenching in the Shaker K+ channel. J. Gen. Physiol. 112:391-408. Chen, F. S., D. Steele, and D. Fedida. 1997. Allosteric effects  of  permeating cations on gating currents during K+ channel deactivation. J. Gen. Physiol. 110:87-100. Claydon, T. W., M. R. Boyett, A. Sivaprasadarao, K. Ishii, J. M. Owen, H. A. O'Beirne, R. Leach, K. Komukai, and C. H. Orchard. 2000. Inhibition of  the K+ channel Kvl.4 by acidosis: protonation of  an extracellular histidine slows the recovery from  N-type inactivation. J. Physiol. * 526:253-264. De Biasi, M., H. A. Hartmann, J. A. Drewe, M. Taglialatela, A. M. Brown, and G. E. Kirsch. 1993. Inactivation determined by a single site in K+ pores. Pflugers  Arch. 422:354-363. Doyle, D. A., C. J. Morais, R. A. Pfuetzner,  A. Kuo, J. M. Gulbis, S. L. Cohen, B. T. Chait, and R. MacKinnon. 1998. The structure of  the potassium channel: molecular basis of  K+ conduction and selectivity. Science 280:69-77. Elinder, F., R. Mairnikko, and H. P. Larsson. 2001. S4 charges move close to residues in the pore domain during activation in a K channel. J. Gen. Physiol. 118:1-10. Fedida, D., N. D. Maruoka, and S. Lin. 1999. Modulation of  slow inactivation in human cardiac Kvl.5 channels by extra- and intracellular permeant cations. J. Physiol. 515:315-329. Frankenhaeuser, B. and A. L. Hodgkin. 1957. The action of  calcium on the electrical properties of  squid axons. J. Physiol. 137:218-244. Harris, R. E., H. P. Larsson, and E. Y. Isacoff.  1998. A permanent ion binding site located between two gates of  the Shaker  K+ channel. Biophys. J. 74:1808-1820. Hesketh, J. C. and D. Fedida. 1999. Sequential gating in the human heart K+ channel Kvl.5 incorporates Q, and Q, charge components. Am. J. Physiol. 277:H1956-H1966. Hille, B. 2001. Ion Channels of  Excitable Membranes. Sinauer Associates, Inc. Sunderland, Massachusetts. Hoshi, T., W. N. Zagotta, and R. W. Aldrich. 1991. Two types of  inactivation in Shaker  K+ channels: effects  of  alterations in the carboxy-terminal region. Neuron 7:547-556. Immke, D., M. Wood, L. Kiss, and S. J. Korn. 1999. Potassium-dependent changes in the conformation  of  the Kv2.1 potassium channel pore. J. Gen. Physiol. 113:819-836. Ishii, K., K. Nunoki, T. Yamagishi, H. Okada, and N. Taira. 2001. Differential  sensitivity of Kvl.4, Kvl.2, and their tandem channel to acidic pH: involvement of  a histidine residue in high sensitivity to acidic pH. J. Pharmacol. Exp. Ther. 296:405-411. Jager, H. and S. Grissmer. 2001. Regulation of  a mammalian Shaker-related  potassium channel, /zKvl.5, by extracellular potassium and pH. FEBS Lett. 488:45-50. Jager, H., H. Rauer, A. N. Nguyen, J. Aiyar, K. G. Chandy, and S. Grissmer. 1998. Regulation of mammalian ST^er-related K+ channels: evidence for  non-conducting closed and non-conducting inactivated states. J. Physiol. 506:291-301. Kiss, L., J. LoTurco, and S. J. Korn. 1999. Contribution of  the selectivity filter  to inactivation in potassium channels. Biophys. J. 76:253-263. Larsson, H. P. and F. Elinder. 2000. A conserved glutamate is important for  slow inactivation in K+ channels. Neuron 27:573-583. Levy, D. I. and C. Deutsch. 1996. Recovery from  C-type inactivation is modulated by extracellular potassium. Biophys. J. 70:798-805. Li-Smerin, Y., D. H. Hackos, and K. J. Swartz. 2000. A localized interaction surface  for  voltage-sensing domains on the pore domain of  a K+ channel. Neuron 25:411-423. Loots, E. and E. Y. Isacoff.  1998. Protein rearrangements underlying slow inactivation of  the Shaker  K+ channel. J. Gen. Physiol. 112:377-389. Loots, E. and E. Y. Isacoff.  2000. Molecular coupling of  S4 to a K+ channel's slow inactivation gate. J. Gen. Physiol. 116:623-635. Lopez-Barneo, J., T. Hoshi, S. H. Heinemann, and R. W. Aldrich. 1993. Effects  of  external cations and mutations in the pore region on C-type inactivation of  Shaker  potassium channels. Receptors. Channels 1:61-71. Nguyen, A., J. C. Kath, D. C. Hanson, M. S. Biggers, P. C. Canniff,  C. B. Donovan, R. J. Mather, M. J. Bruns, H. Rauer, J. Aiyar, A. Lepple-Wienhues, G. A. Gutman, S. Grissmer, M. D. Cahalan, and K. G. Chandy. 1996. Novel nonpeptide agents potently block the C-type inactivated conformation  of  Kvl.3 and suppress T cell activation. Mol. Pharmacol. 50:1672-1679. Olcese, R., R. Latorre, L. Toro, F. Bezanilla, and E. Stefani.  1997. Correlation between charge movement and ionic current during slow inactivation in Shaker  K+ channels. J. Gen. Physiol. 110:579-589. Ortega-Saenz, P., R. Pardal, A. Castellano, and J. Lopez-Barneo. 2000. Collapse of  conductance is prevented by a glutamate residue conserved in voltage-dependent K+ channels. J. Gen. Physiol. 116:181-190. Pardo, L. A., S. H. Heinemann, H. Terlau, U. Ludewig, C. Lorra, O. Pongs, and W. Stuhmer. 1992. Extracellular K+ specifically  modulates a rat brain K+ channel. Proc. Natl. Acad. Sci. U. S. A. 89:2466-2470. Perez-Cornejo, P., P. Stampe, and T. Begenisich. 1998.. Proton probing of  the charybdotoxin binding site of  Shaker  K+ channels. J. Gen. Physiol. 111:441-450. Steidl, J. V. and A. J. Yool. 1999. Differential  sensitivity of  voltage-gated potassium channels Kvl.5 and Kvl.2 to acidic pH and molecular identification  of  pH sensor. Mol. Pharmacol. 55:812-820. Vallee, B. L. and D. S. Auld. 1990. Zinc coordination, function,  and structure of  zinc enzymes and other proteins. Biochemistry 29:5647-5659. Wang, Z. and D. Fedida. 2001. Gating charge immobilization caused by the transition between inactivated states in the Kvl.5 channel. Biophys. J. 81:2614-2627. Wang, Z., X. Zhang, and D. Fedida. 2000. Regulation of  transient Na+ conductance by intra- and extracellular K+ in the human delayed rectifier  K+ channel Kvl.5. J. Physiol. 523:575-591. Wood, M. J. and S. J. Korn. 2000. Two mechanisms of  K+-dependent potentiation in Kv2.1 potassium channels. Biophys. J. 79:2535-2546. Woodhull, A. M. 1973. Ionic blockage of  sodium channels in nerve. J. Gen. Physiol. 61:687-708. Yang, Y., Y. Yan, and F. J. Sigworth. 1997. How does the W434F mutation block current in Shaker  potassium channels? J. Gen. Physiol. 109:779-789. Yellen, G. 1997. Single channel seeks permeant ion for  brief  but intimate relationship. J. Gen. Physiol. 110:83-85. Zhang, S., S. J. Kehl, and D. Fedida. 2001a. Modulation of  Kvl.5 potassium channel gating by extracellular zinc. Biophys. J. 81:125-136. Zhang, S., D. C. Kwan, D. Fedida, and S. J. Kehl. 2001b. External K+ relieves the block but not the gating shift  caused by Zn2+ in human Kvl.5 potassium channels. J. Physiol. 532:349-358. 3. The external K+ concentration and mutations in the outer pore mouth affect  the inhibition of  Kvl.5 current by Ni2+ 3.1 Introduction Kvl .5 (KCNA5) channels, which are expressed in cardiac myocytes (Fedida et al., 1993; Tamkun et al., 1991) and in smooth muscle cells of  airways, the intestine and the vasculature (Adda et al., 1996; Clement-Chomienne et al., 1999), are members of  a major structural class of K+ channels in which the a-subunit consists of  6 transmembrane (TM)  segments with a pore-forming  or P-region positioned between transmembrane segment five  (S5) and S6. A characteristic feature  of  the 6TM-1P  subunit is the charge-bearing S4 domain whose movement upon membrane depolarization (Baker et al., 1998; Larsson et al., 1996) is linked to the opening of  the activation gate which is believed to comprise the cytoplasmic ends of  the four  S6 regions in the tetrameric channel assembly. Macroscopic currents through Kvl.5 channels resemble delayed rectifier  currents. Thus, with a strong sustained depolarization, channel activation is rapid and voltage-dependent whereas inactivation is voltage-independent and occurs on a timescale of  seconds. Kvl.5 channels exhibit only outer pore (P/C-type) inactivation (Fedida et al., 1999) and in this regard are different  from  Shaker  channels which also show inner pore (N-type) inactivation (Hoshi et al., 1991). The term C-type inactivation was coined to describe the slow inactivation process in Shaker  that was uncovered when ball-and-chain or N-type /nactivation was removed (ShakerlR)  by deletion of  the cytoplasmic N-terminal residues 6-46. C-type inactivation is A version of  this chapter has been published. Kwan, D. C. H., Eduljee, C., Lee, L., Zhang, S., Fedida, D., and Kehl, S. J. (2004) The external K+ concentration and mutations in the outer pore mouth affect  the inhibition of  Kvl.5 current by Ni2+. Biophys. J. 86(4):2238-2250. coupled to channel activation and is believed to involve a conformational  change in the outer pore mouth that extends to the selectivity filter  delimited by the highly conserved GYG sequence. Because C-type inactivated ShakerlR  (Starkus et al, 1997) and Kvl .5 (Wang et al, 2000a) channels are able to conduct Na+ ions, the current view is that the conformational  change at the outer pore mouth involves an incomplete constriction rather than a complete collapse. An important consequence of  C-type inactivation is a leftward  shift  of  the gating charge versus voltage relationship, or Q-Vcurve,  and charge immobilization (Fedida et al, 1996; Olcese et al, 1997). In ShakerlR  channels the residue at position 463 in the S6 segment was the first  shown to influence  the rate of  C-type inactivation (Hoshi et al, 1991). Subsequently, point mutations of the threonine residue (T449) in the outer pore mouth were shown to dramatically accelerate (T449E, T449A, T449K) or slow (T449Y,T449V) C-type inactivation (Lopez-Barneo et al, 1993). In Kvl.5 channels the residue homologous to T449 is R487 and it has been shown that inactivation is substantially slowed in Kvl.5 R487V when Na+ is the charge carrier but not when K+ is the permeant ion (Fedida et al, 1999; Wang et al, 2000a). The finding  that ShakerlR/Kvl.5  channels with the pore mutation W434F/W472F were Na+- but not K+-conductive and showed wild-type gating charge behaviour, including gating charge immobilization following  channel inactivation (Chen et al, 1997; Olcese et al, 1997), was one of  the first  indications of  the complexity of  outer pore inactivation. To account for  the properties of  the ShakerlR  W434F non-conducting mutant it was proposed that there was also a so-called P-type inactivation process that prevented K+ conduction but which was different  from C-type inactivation in that it did not affect  gating charge movement (Olcese et al, 1997; Yang et al, 1997). Restoration of  ionic current in the double mutant Shaker  W434F, T449Y supports the -hypothesis that enhanced inactivation accounts for  the ShakerlR  W434F conductance loss (Yang et al., 2002). An intriguing divergence in the structure-function  relationships of  Kvl .5 and ShakerlR  is seen in the response to extracellular acidification.  In Kvl .5 external protons cause, in addition to a rightward shift  of  the g-V  curve that is often  referred  to as the gating shift,  a concentration-dependent decrease of  the maximum macroscopic conductance (gmax) as well as an acceleration of  the inactivation rate of  residual currents (Kehl et al., 2002; Steidl and Yool, 1999). In contrast, in ShakerlR  channels increasing [H+]0 does not reduce gmax but the gating shift  and the speeding of  inactivation are observed (Perez-Cornejo, 1999; Starkus et al., 2003). A number of lines of  evidence now support the view that protonation of  a histidine residue (H463), the equivalent of  Shaker  F425, in the pore turret (S5-P linker) plays an important role in the proton-induced conductance loss/block in Kvl .5. Thus, in the Kvl .5 H463Q mutant there is a large rightward shift  of  the concentration dependence of  the H+0 block (Kehl et al., 2002). The finding that the H+ 0 block is antagonized by K+c and is also reduced in the R487V mutant (Jager and Grissmer, 2001; Kehl et al., 2002) has suggested that the protonation of  H463 facilitates  an inactivation process requiring R487. An alternative explanation involving direct pore block by protons has been ruled out on the basis of  single channel recordings (Kwan et al., 2003) and the finding  that the Na+ current through inactivated Kvl .5 channels is maintained following extracellular acidification  (Zhang et al., 2003). Additional support for  a crucial role of  H463 in the H+0-induced decrease ofg max is provided by reports showing that divalent cations known to bind to histidine residues also affect Kvl.5 currents. Harrison et al. (1993) first  reported that extracellular Zn2+ blocks Kvl.5 currents and, as with the H+0 block, this effect  of  Zn2+ is inhibited either by increasing K+0 or by mutating H463 and/or R487 (Kehl et al., 2002). Ni2+ is also a histidine ligand and although it too has been reported to block Kvl.5 currents expressed in Chinese hamster ovary (CHO) cells (Perchenet and Clement-Chomienne, 2001), the mechanism of,  and the molecular determinants for  the block have not been resolved. To test the hypothesis that the mechanistic basis for  the Ni2+ block is essentially the same as that outlined above for  Zn2+ and H+0, we set out in this study to address the following  questions. Is the block of  Kvl.5 by Ni2+antagonized by increasing [K+]0? Does Ni2+ speed the inactivation rate of  residual Kvl .5 currents? Is the effect  of  Ni2+ affected  either by mutating H463, a putative Ni2+ coordination site, or by mutating R487, a site implicated in the regulation of  outer pore inactivation? Are gating currents affected  by Ni2+? And finally,  is the blocking effect  of  Ni2+ replicated by other divalent cations such as Co2+, Cd2+ and Mn2+? 3.2 Materials and Methods 3.2.1 Cell preparation As described previously (Wang et al., 2000a), wild type {wt)  and mutant human Kvl .5 channels, henceforth  referred  to simply as Kvl.5 channels, were studied in a human embryonic kidney cell line (HEK-293) (Wang et al., 2000b). Cells were dissociated for  passage by using trypsin-EDTA and were maintained in minimum essential medium (MEM), 10% fetal  bovine serum, penicillin-streptomycin and 0.5 mg ml"1 gentamicin in an atmosphere of  5% C02 in air. All tissue culture supplies were obtained from  Invitrogen (Burlington, ON, Canada). Point mutations of  the wt Kvl.5 a-subunit in the plasmid expression vector pcDNA3 were made using the Quikchange Kit (Stratagene, La Jolla, CA, USA) to convert the histidine (H) residue at position 463 to glutamine (Q) (H463Q) or the arginine (R)) at position 487 to valine (R487V). Stable transfections  of  HEK-293 cells were made using 0.8 |ig. of  Kvl.5 H463Q or Kvl.5 R487V cDNA and 2 [iL of  Lipofectamine  2000 (Invitrogen). Geneticin (0.5 mg/mL) was added 48 hrs after  transfection.  Because Shaker-related  channels such as Kvl .5 are homotetramers (MacKinnon, 1991), a given point mutation will exist in each of  the four  subunits of  the channel assembly. 3.2.2 Recording solutions The standard bathing solution contained, in mM, 140 NaCl, 3.5 KC1, 10 HEPES, 2 CaCl2, 1 MgCl2, 5 glucose and its pH was adjusted to 7.4.with NaOH. HEPES was replaced by MES when the pH of  the extracellular solution was less than 6.8 in the experiments directly comparing the proton block and the Ni2+ block. Where the effect  of  the external concentration of  potassium ([K+]0) on the divalent metal cation block was examined, a nominally K+-free  solution was made by substituting NaCl for  KC1 and, for  [K+]0greater than 3.5 mM, NaCl was replaced by KC1. The standard patch pipette solution for  recording K+ currents contained 130 KC1, 4.75 CaCl2 (pCa2+ = 7.3), 1.38 MgCl2, 10 EGTA, 10 HEPES and was adjusted to pH 7.4 with KOH. Solutions of divalent metal ions were made by dilution of  0.1 to 1 M stock solutions of  the chloride salt in distilled water. At pH 7.4 the concentration of  Ni2+ that can be used was limited to 10 mM or less by virtue of  the solubility product for  Ni(OH)2 (~ 2 x 10"16). Mouse fibroblasts  expressing Kvl.5 channels at a low density were used to record unitary currents from  outside-out patches. The inside face  of  the patch was exposed to standard patch pipette solution and the outside face  was exposed to standard bath solution either with or without added Ni2+. For gating current recordings the bath solution contained, in mM, 140 NMGC1, 1 MgCl2, 10 HEPES, 2 CaCl2, 10 glucose and the pH was adjusted to 7.4 with HC1. The patch pipette solution contained 140 NMGC1, 1 MgCl2, 10 HEPES, 10 EGTA and was adjusted to pH 7.2 with HC1. Chemicals were purchased from  the Sigma Aldrich Chemical Co. (Mississauga, ON, Canada). 3.2.3 Signal recording and data analysis Macroscopic currents were recorded at room temperature (20-22°C) using the patch clamp technique primarily in the whole cell configuration.  In some of  the cell lines expressing mutant Kvl.5 channels at a high level, i.e., the H463Q and some of  the R487V mutants, the large amplitude of  the whole cell currents necessitated recording macroscopic currents from  outside-out patches. Voltage clamp experiments were done with an EPC-7 patch clamp amplifier  and Pulse+PulseFit software  (HEKA Electronik, Germany). Patch electrodes were made from  thin-walled borosilicate glass (World Precision Instruments, FL, USA) and had a resistance of  1.0 to 2.5 MQ measured in the bath with standard internal and external solutions. Typically, 80% series resistance compensation was used and an on-line P/N  method, for  which the holding potential was -100 mV and the scaling factor  was 0.25, was used to subtract the leak current as well as any uncompensated capacitive.currents. Current signals filtered  at 3 kHz (-3dB, 8-pole Bessel) were digitized (16 bit) at a sampling interval of  100 (is (10 kHz). Voltages have been corrected for  the liquid junction potentials. In an experiment, a section of  glass coverslip with cells attached to it was placed in the recording chamber (0.5 ml volume) and was continuously perfused  with bathing solution. After recording currents in the control solution the inflow  was switched to the test solution and once 5-6 ml had been flushed  through the bath the treated responses were recorded. Recovery currents were taken after  flushing  the bath with 5-6 ml of  control solution. If  the recovery currents were not within ±15% of  the pre-treatment amplitudes the data for  that cell were discarded. By this criterion most cells showed recovery. To quantify  the effect  of  Ni2+ and other metal cations on Kvl .5, tail currents were recorded at -40 or -50 mV following  depolarizing pre-pulses of  differing  magnitude. Peak tail current amplitudes were obtained by fitting  a polynomial function  and taking the fitted  value for the maximum current. After  normalization of  tail currents either to the maximum current of  the control or the treated response, data points were fitted  to a single Boltzmann function: A y = l + e x p ( ^ ) ^ where, when^ is the current normalized with respect to the control response, A is the proportion of  the control gmax. When y is the current normalized with respect to the maximal treated current, A is the best fit  value for  the normalized maximal response and ideally has a value of  unity. I7,, is the half-activation  potential or mid-point of  the activation curve, V  is the voltage during the pre-pulse and .v is the slope factor,  in mV, reflecting  the steepness of  the voltage dependence of gating. To quantify  gating charge movement during activation, charge-voltage (Qon-V) curves were generated by time integration of  on-gating currents as described previously (Chen et al., 1997). Activation gating in Kvl .5 is best fit  by the sum of  two Boltzmann functions  where the larger component, known as Q2, represents -80% of  the total charge movement (Hesketh and Fedida, 1999). However, for  simplicity, Q-Vdata  obtained at pH 7.4 and 5.4 were fitted  to Equation (3.1) whereY  is the charge moved, A is the maximal charge (Q MA X)  and Kis the voltage at which the on-gating charge (Q T  J is evoked. V, 2 represents the mid-point of  the Q-V curve and 5 reflects  the steepness of  the voltage dependence of  charge movement. Concentration-response data were fitted  to the Hill equation: 1 ) ^ K d ) where y is the proportion of  the control gmcLX,  K D is the equilibrium dissociation constant for  the test cation (X2') andtnw is the Hill coefficient  reflecting  the number of  test cations binding per channel. Microscopic currents were low-pass filtered  at 3 kHz (8-pole Bessel), sampled at 10 kHz and digitally-filtered  at 1 kHz for  the data analysis using TAC and TACFit (Bruxton, Seattle). Leak and uncompensated capacitive currents were subtracted using a template generated from blank sweeps. Half-amplitude  threshold analysis was used to idealize single channel recordings for  the generation of  dwell time histograms. Data are expressed as the mean ± SEM except for  the values obtained by non-linear least-squares fitting  routines (Igor, Wavemetrics, OR, USA) which are expressed as the mean ± SD. The paired-sample t test was used to compare the inactivation rates of  residual currents in Ni2+ and H+0. A /rvalue of  0.05 or less was considered significant. 3.3 Results Shown in Figure 3.1 A are traces confirming  the block of  Kvl .5 currents by external Ni2+. From a holding potential of  -80 mV and with 0 mM K+0, currents were evoked by a family  of 300 ms depolarizations from  -45 to +35 mV with a cycle length of  5 s. Tail currents were recorded at -40 mV. After  obtaining the control responses, the perfusate  was switched to a test ([X 2+]) bathing solution containing 0.1 mM Ni2+ and then to one containing 1 mM Ni2+. Complete recovery was obtained after  returning to Ni2+-free  solution. As noted previously (Perchenet and Clement-Chomienne, 2001), and in contrast to the effects  with Zn2+ (Zhang et al., 2001b), with Ni2+ there was neither a significant  change of  the activation kinetics nor an obvious effect  on the decay of  residual pulse currents. The effect  of  Ni2+ on the current behaviour during longer depolarizing pulses is examined below (Figure 3.4). 0 mM K 35 mV -40 mV | 1 nA 1 mM Ni .2+ B. f  1.2 • c a? | 0 . 8 • o 5 Z 0.4 • ro 6 0 . 0 • control . JPn0o0ouo0u recovery 0.25 mM Ni . . . 4 0 . 5 m M Ni + T~ "I -40 -20 20 40 60 pulse \oltage (mV) 0.1 mM Ni 2+ recovery C. 1.2 1.0 H 0.8 0.6 0.4 0.2 0.0 tgM9geBguH 0.25 mM Ni ~i 1 r -40 -20 0 20 40 pulse voltage (mV) Figure 3.1 Ni2+ block of  Kvl.5 currents in 0 mM K+0. A. Control currents evoked by a family of  300 ms depolarizations from  -45 to +35 mV, here shown in 10 mV increments, from  a holding potential of  -80 mV. Tail currents were recorded at -40 mV. Perfusion  of  solution containing 0.1 mM Ni2+ and then 1 mM Ni2+ caused a concentration-dependent inhibition of  the current. Recovery traces illustrate the complete reversal of  the Ni2+ block. B and C. Ni2+ decreases gmca and shifts  the g-V.curve  slightly rightward. B. Peak tail current at -40 mV following  a 300 ms depolarization to the voltage indicated on the x-axis. Note the absence of  any voltage dependence of  the inhibition between 0 and 50 mV. C. The g-V  relationship derived by normalizing tail currents with respect to the maximum tail current shows that Ni2+ caused a 10 mV shift  of  the half-activation  voltage. Current tails in 0.5 mM Ni2+ were too small to be unequivocally analysed. 3.3.1 Increasing [K+]0 causes a rightward shift  of  the concentration dependence of  the Ni2+ block In order to quantify  the block by Ni2+, g- V  curves were constructed from  peak tail currents as described in the Methods section. Panel B of  Figure 3.1 plots the peak tail current amplitude versus the pulse voltage for  the same cell in 0 mM K+0 without Ni2+ and with 0.25 or 0.5 mM Ni2+. In this cell 0.25 mM and 0.5 mM Ni2+ decreased the maximum tail current, and by extension the maximum conductance (g max), by approximately 70% and 90%, respectively. To more clearly illustrate the effect  of  Ni2+ on the midpoint (V, !) of  the g- V  curve, the currents in panel B were normalized with respect to the maximum current for  the same treatment group and are presented in panel C. It is evident that Ni2+ caused a rightward shift  of  the g-V  curve and this is assumed to reflect  a change of  surface  charge due to screening and/or binding to the channel. With 0.25 mM Ni2+ the shift  of  V, A_ determined from  the best fit  of  the g-V  data to the Boltzmann function  was 10.6 ± 0.9 mV (n = 4). The gating shift  with 0.5 mM Ni2+ was not determined because the standard deviation in the fitted  values for  V,,  was quite large. Figure 3.2 shows the concentration-response relationship for  the block of  Kvl.5 by Ni2+ and the influence  of  [K+]0 thereon. Panel A illustrates representative current traces from  3 different  cells in 0 mM (left),  3.5 mM (middle)  and 140 mM (right)  K+0. In the absence of  Ni2+ (-Ni2+) the current in each of  the K+0 concentrations had a similarly slow rate of  decay. The inward tail current recorded at -40 mV in 140 mM K+0 is due to the shift  of  EK  to ~0 mV. To produce a similar degree of  block in the three different  experiments it was necessary to increase the Ni2+ concentration to offset  the effect  of  increasing [K+]0. Note that in each example the Ni2+ block was not associated with an acceleration of  pulse current decay. The latter observation, together with the fact  that the tail current decay was not slowed, as best seen with the traces in 140 mM K+0, supports the conclusion that a block of  the open channel occurring with intermediate-to-slow kinetics (vis  a vis the activation rate) is not involved. For the graph in Figure 3.2 B, the gmax relative to the control value has been plotted against the concentration of Ni2+ for  experiments in which [K+]0 was 0 mM (open  circles),  3.5 mM (open  triangles)  or 140 mM (open  squares). The solid lines overlaying the three data sets represent the best fit  to Equation 3.2. With 0 mM K+0 the K D for  the Ni2+ block was 0.15 ± 0.01 mM and nH  was 1.3 ± 0.1. Increasing [K+]0 to 3.5 mM increased the K D to 0.44 ± 0.02 mM and nH  was 1.6 ± 0.2. With 140 mM K+0 the K D was 3.1 ± 0.3 mM and nH  was 0.9 ± 0.1. These results clearly demonstrate that, as with the block by H+0 and Zn2+, the block of  Kvl.5 by Ni2+ is antagonized by increasing [K+]0. 3.3.2 The time courses of  the onset and the offset  of  the Ni2+ block are similar Using a fast  solution application system, Perchenet and Clement-Chomienne (2001) noted that the offset  of  the Ni2+ block was rapid but that the onset was comparatively much slower. They found,  with test pulses delivered at 15 s intervals and using 1 mM Ni2+ and 5 mM K+0, that steady-state block was reached only after  5 to 7 minutes. Since a similar phenomenon is not seen with H+0 or Zn2+, we felt  it was important to characterize the time dependence of  the Ni2+ block and did so by comparing the time course of  the current inhibition by Ni2+ with that by H+0. Graphs summarizing the outcome of  this comparison are shown in Figure 3.3. For each graph, the peak tail current, measured at -40 mV following  a 300 ms pulse to 50 mV applied at 10 s intervals, was plotted against the elapsed time. In Figure 3.3 A, Ni2+ and H+0 were applied for  the duration indicated by the horizontal bar at concentrations of  150 |J,M and 0.16 |iM (pH 6.8) (Kehl et al., 2002), respectively, and in 0 mM K+0 to cause roughly 50% block of  the current at the A. 0 mM K+„ 3.5 mM K + 0 140 mM K + 0 ± 0.25 mM Ni2* ± 0.5 mM Ni2+ ± 5 mM Ni2+ Ni concentration (mM) Figure 3.2 Increasing [K+]0 changes the concentration dependence of  the block of  Kvl.5 by Ni2+. A. Representative traces obtained from  three different  cells showing, superimposed, the currents evoked in the K+ 0 concentrations indicated either without (-) or with (+) the Ni2+ concentration indicated. The voltage protocol consisted of  a 300 ms step from  -80 mV to 50 mV followed  by a step to -40 mV. Increasing [K+]0 necessitates a higher concentration of  Ni2+ to produce roughly the same degree of  block. The time calibration is the same for  the three sets of traces. B. The concentration-response relationship for  Ni2+ in 0, 3.5 and 140 mM K+0 shows that increasing [K+]0 from  nominally K+-free  to 3.5 mM shifted  the K D from  0.15 ± 0.01 mM to 0.44 ± 0.02 mM. Increasing K+0 to 140 mM shifted  the K D for  the Ni2+ block to 3.1 ± 0.3 mM. Each point represents the mean ± SEM of  measurements from  3-7 cells. steady state. In four  such experiments we consistently found  that the time courses for  the onset and offset  of  the block by Ni2+ and H+0 were similar. Since the failure  to uncover any asymmetry in the on- and off-  time courses might be attributed to the absence of  K+0, experiments were also done with 5 mM K+0 which necessitated using higher concentrations of  Ni2+ and H+ 0 to compensate for  the effect  of  K+0 on the block. Figure 3.3 B shows that the outcome was still the same: after  switching from  the control to the test perfusate  the relaxation to the steady-state was complete in less than a minute, a time frame  that appears to reflect  primarily the dynamics of solution exchange in the bath and is much shorter than the onset noted by Perchenet and Clement-Chomienne (2001). 0 . 0 -150 pM Ni . 0 mM K 0 pH 6.8, 0 mM K , • ^ M H m * ~n 1 1— 200 300 400 elapsed time (s) ~~I 600 1.2 n 1 . 0 -•2 0.4-0 . 0 -600 pM Ni 5 m M K , pH 6.2. 5 mM K c 200 300 400 elapsed time (s) Figure 3.3 A comparison of  the time course of  the onset and offset  of  the inhibition of  Kvl.5 by Ni2+and H+0. Concentrations of  Ni2+ or H+ 0 producing approximately 50% steady-state inhibition with 0 mM K+0 (A)  and, in a different  cell, • with 5 mM K+0 (B)  were used. Peak tail currents measured at -40 mV following  a 300 ms depolarization to 50 mV applied at a 10 s interval are plotted against the elapsed time. The horizontal bar indicates the duration of  each application. The results do not reveal any obvious asymmetry in the onset versus the offset  of the block either with or without K+0. As with H+0, the time course for  the development or reversal of  the Ni2+ block was similar and in both cases is presumed to reflect  the time course of  solution exchange in the bath. 3.3.3 Ni2+ block is associated with a slight acceleration of  inactivation of  residual currents In addition to blocking Kvl .5 currents, extracellular acidification  accelerates the rate of inactivation of  residual currents (Kehl et al., 2002; Steidl and Yool, 1999) and this was the motivation for  determining if  there was a similar association between block and inactivation with Ni2+. Our approach to addressing this question was to use cells expressing Kvl .5 channels at a very high density so that despite the reduction of  gmax by 80-95% the residual, currents were virtually unfettered  by endogenous HEK currents and could therefore  be unambiguously analysed. The voltage protocol consisted of  a 5 s step from  -80 mV to 50 mV followed  by brief depolarizations to 50 mV to track recovery from  inactivation (Fedida et al., 1999). An interval of 120 s between the 5 s pulses was used to permit complete recovery from  inactivation in the experiments with Ni2+. Initially, these experiments were done with 0 mM K+0, but the interpretation of  the data was confounded  by a very slowly rising phase of  current with a time constant of  1-1.5 s in 1 mM Ni2+ and 200-300 ms at pH 5.9 which followed  a normally activating component of  current (not shown). This slow component was not observed with 3.5 mM K+0, consequently this was the [K+]0 used when comparing the effects  on inactivation of concentrations of  H+0 and Ni2+ that reduce gmax by 80 to 95% (Figure 3.2 and Kehl et al., 2002). Results representative of  those obtained in five  experiments with 2 mM Ni2+ and 5 experiments with pH 5.4 are shown in Figure 3.4 A and B where the effects  of  H+0 and Ni2+, respectively, were tested on the same cell. At pH 5.4 the inactivation of  the residual current during the 5 s pulse was well-fitted  by a single exponential with a time constant of  91 ms and the steady-state current was -25% of  the peak amplitude. At pH 5.4 the mean inactivation time constant (r /nac,) at 50 mV was 101 ± 3 ms (n  = 5 cells). In Figure 3.4 A recovery from  inactivation, tested by 50 ms depolarizations delivered from  0.5 s up to 96 s after  the 5 s depolarization, was fitted  to a single exponential with a time constant of  4.3 s. The mean T recovery at pH 5.4 was 4.2 ± 0.1 s. In contrast, currents recorded after  switching from  pH 5.4 solution to perfusate  containing 2 mM Ni2+ at pH 7.4 (Figure 3.4 B) showed much slower inactivation as well as slower recovery from inactivation: vmacl = 1.69 s and T recover y = 24.8 s. In the five  cells tested with 2 mM Ni2+ the mean -value for  vmaa and T recover y was 1.71 ± 0.07 s and 23.5 ± 2.1 s, respectively. Because of  their very large amplitude, currents in Ni2+-free  medium at pH 7.4 could not be recorded from  these cells, however the best fit  to a single exponential of  the current decay during 7-10 s depolarizations to 60 mV at pH 7.4 in Kvl.5 is typically of  the order of  2-3 s (Kehl et  al.,  2002) and the T ncmerv measured at -80 mV in 5 mM K+0 and using a similar voltage protocol is 1.1 s (Fedida et  al., 1999). A. pH 5.4 2 mM Ni2 + Figure 3.4 Inactivation and recovery kinetics of  Kvl.5 at pH 5.4 or with 2 mM Ni2+. A comparison of  the residual current behaviour, done in 3.5 mM K+0 with a concentration of  H+0 (A) or Ni2+ (B)  estimated to block 80-95% of  the channels, reveals divergent effects  on inactivation. From a holding potential of-80  mV, the voltage protocol consisted of  a 5 s step to 50m V followed  by 50 ms steps at variable intervals to 50 mV to monitor recovery from  inactivation. Current during the 5 s pulse is shown expanded on the right side of  the figure.  A. At pH 5.4 the inactivation of  current during the 5 s pulse is well-fitted  by a single exponential with a time constant of  91 ms. Peak currents, marked by the filled  circles, that were evoked by the 50 ms test pulses were fitted  to a single exponential with a time constant of  4.3 s. B. In the same cell after switching to solution containing 2 mM Ni2+ at pH 7.4, inactivation was =20 times slower ( t m a a = 1.69 s) and recovery from  inactivation was =5 times slower (T recover y = 24.8 s) than at pH 5.4. 3.3.4 The K D for  the Ni2+ block is increased in the H463Q and R487V mutants We next examined the effect  of  Ni2+ in Kvl .5 channels in which either a putative Ni2+ binding site in the S5-P linker (turret) was mutated to a glutamine residue (H463Q) or the residue analogous to Shaker  T449 was changed from  arginine to valine (R487V). To circumvent the potential problem of  changes of  the K+0-dependence of  the block relief,  the analysis of  the effect of  Ni2+ on currents from  these mutated channels was done with 0 mM K+0. Concentration-response curves for  the Ni2+ block of  currents from  Kvl.5 H463Q (filled  squares) and Kvl.5 R487V (filled  circles)  are shown superimposed in Figure 3.5. As with the block by H+0 and Zn2+ (Kehl et al., 2002), the concentration dependence for  the block by Ni2+ was shifted  substantially to the right by either mutation. In Kvl .5 R487V the K D was estimated to be 2.8 ± 0.004 mM or roughly 20-fold  higher than in wt Kvl .5. With Kvl .5 H463Q the concentration dependence of the block was much more shallow (n H  -0.4) than in wt Kvl .5 and the K D was estimated by extrapolation to be 24 ± 8 mM which is 100- to 200-fold  higher than in wt Kvl.5. 3.3.5 Ni2+ decreases channel availability Macroscopic current amplitude (I) is, in general, the product of  the number of  channels available (N),  the single channel current (/) and the channel open probability (Pj.  To gain a clearer insight into which of  these variables was affected  by Ni2+ ions, recordings were made from  outside-out patches containing a single channel. Figure 3.6 A shows representative, consecutive control sweeps evoked by a 300 ms depolarization from  -80 mV to 100 mV applied at a frequency  of  0.1 Hz. As reported previously (Chen and Fedida, 1998), channel openings occurred in bursts of  varying duration and within bursts channel closings were frequent  but brief. With seconds-long pulses (not shown), we observed closed states with longer mean dwell times o 4-u/Kv1 5 in 140 mM K o 0.0 0.01 TTTTTJ 1 1 I I I I I l| 1 1 1 I I 0.1 1 2+ Ni concentration (mM) • 1 0 TTq 100 Figure 3.5 Ni2+ sensitivity is reduced in Kvl.5 H463Q and Kvl.5 R487V. Experiments with the mutant channels were done with 0 mM K+0 to preclude a change of  the K+0 binding as the basis for  the change of  the sensitivity to Ni2+. Because Ni2+ is known to bind to histidine (H) residues, a mutant was constructed in which glutamine (Q) was substituted for  H463, a residue in the S5-P linker that forms  part of  the outer pore vestibule. Kvl .5 H463Q {filled  squares) was 100- to 200-fold  less sensitive to Ni2+ (K D = 24 ± 8 mM; nH  = 0.4 ± 0.04) compared to the wt Kvl .5 responses (dashed line taken from  Figure 3.2) measured in the same recording condition, i.e., 0 mM K+0. In another mutant construct, the arginine (R) residue near the entrance to the pore mouth that has been implicated, by alignment with Shaker  T449, in the outer pore inactivation mechanism, was mutated to valine. The sensitivity of  Kvl .5 R487V currents (filled  circles) to Ni2+ was approximately 20-fold  less (K D = 2.8 ± 0.004 mM; nH  = 0.7 ± 0.001) than that measured under the same recording conditions in wt Kvl .5. The dotted line, which was taken from  Figure 3.2, represents the line fitted  to the block of  wt Kvl .5 in 140 mM K+0. which are assumed to reflect  a multistep inactivation pathway. Double-Gaussian fits  to the control all-points amplitude histogram (e.g.,  Figure 3.6 C) indicates an open channel current (/) of  1.7 ± 0.1 pA (n  = 8 patches). After  switching to medium with 0.5 mM Ni2+, which is the-K D for  the block in 3.5 mM K+0 (Figure 3.2), there was no significant  change of  the single channel current (e.g.,  Figure 3.6 Z); 1.6 ± 0.1 pA, n = 6,patches), and the P0 in sweeps containing channel activity was also not significantly  affected  (P 0.M,=  0.64 ± 0.06 versus P0 _N j = 0.61 ± 0.06). There were, however, many more blank sweeps in the presence of  the Ni2+ (Figure 3.6 B). Channel availability (N),  defined  as the number of  sweeps with channel activity divided by the total number of  sweeps, decreased significantly  from  the control value of  0.90 ± 0.06 (n = 6 patches) to 0.43 ± 0.14 (n = 6 patches) in Ni2+. A. ' 2+ 0 mM Ni 0 B-0.5 mM Ni 0 c. littmiiittfftiiii J p i W r w f 4 0 0 -c J W I M d l W f  3 0 0 c a 8 2 0 0 -A r t W W L L U L lu l 1 0 0 -J w W i w ° V -2 D. JlWWWf'l'iW jifwnnppwrnijiP 1 6 0 0 -c •9 f  8 0 0 -I I W W W l l 3 O L> 4 0 0 -?A i j w t i w W M ^ 100 ms JIKWmVm -2 k T 0 t 2 3 amplitude (pA) J LmflTTTTW T=— I r - 1 0 1 2 3 amplitude (pA) Figure 3.6 Ni2+ effects  at the single channel level. A. Shown here are 10 representative and consecutive control sweeps in a one-channel, outside-out patch that were evoked by a 300 ms pulse from  -80 mV to 100 mV applied at 0.1 Hz and with [K+], = 140 mM and [K+]0 = 3.5 mM. Data were digitally filtered  at 1 kHz. B. From the same patch as in ,4,'10 consecutive sweeps evoked with the same voltage protocol but with 0.5 mM Ni2+ in the external perfusate.  The main effect  of  Ni2+ is to reduce channel availability. Representative all-point amplitude histograms from  a different  one-channel patch in control and 0.5 mM Ni2+-containing perfusate  are shown in panels C and D, respectively. A double Gaussian fit  to data gave a mean current in each case of 1.6 pA. 3.3.6 Ni2+ causes a rightward shift  of  the Qon-Vcurve but does not affect  Q ^ A possible explanation for  the current block by Ni2+ is that one or more transitions in the gating pathway is prevented. To address that possibility; gating currents were recorded in an HEK-293 cell line expressing Kvl.5 W472F channels. The W472F mutation produces channels that are not K+ conductive, but which have normal gating currents. Figure 3.7 A shows gating current traces in control solution and in 1 mM Ni2+. On-gating currents were evoked by 12 ms pulses between -60 and 130 mV from  a holding potential of  -100 mV. In the control traces, charge movement was first  evident at approximately -50 mV and the peak amplitude and decay rate increased as the intensity of  the depolarization increased. Following depolarizations up to -10 mV the off-gating  current at -100 mV was rapid (e.g.,  Figure 3.7 B, upper traces) but following  stronger depolarizations there was a clear rising phase to the off-gating  current and the peak current was substantially smaller and occurred much later (e.g.,  Figure 3.7 B, lower traces) than was the case following  steps to -10 mV or less. This pronounced change of  o^-gating current following  stronger depolarizations has been attributed at least in part to a weakly voltage-dependent transition in the return pathway between the open and closed states (Perozo et al., 1993). To construct the charge-voltage (Q-V)  curves shown in Figure 3.7 C, on-gating currents were integrated and Qon was normalized with respect to the control maximal charge movement (<Qmax). Although charge movement is better fitted  by a double Boltzmann function  to account for a smaller component of  charge movement with depolarizations up to -20 mV (Hesk'eth and Fedida, 1999), the data of  Figure 3.7 C were fitted  to a single Boltzmann function. In 6 experiments of  the type illustrated in Figure 3.7, the control F, and s were -6.8 ± 1.2 mV and 7.0 ± 1.4 mV. After  switching to 1 mM Ni2+ F, was 2.2 ± 0.8 mV and s was 9.2 ± 1.2 mV. The difference  in V,.  between 1 mM Ni2+ and control medium was 9.0 ± 3.0 mV. Aside from  this gating shift,  the gating current was essentially unaffected  by 1 mM Ni2+. In contrast to the situation with 1 mM Zn2+ where Qmax decreased by approximately 15% (Zhang et al., 2001a), Qmax was unchanged by 1 mM Ni2+. \oltage (mV) Figure 3.7 Gating charge movement with 1 mM Ni2+ in Kvl.5. To determine if  the conductance loss caused by Ni2+ was due to an inhibition of  transitions in the activation pathway, the effect  of  1 mM Ni2+ on gating charge movement in Kvl.5 W472F, a non-conducting mutant, was examined. Internal and external permeant ions were replaced by NMG+ to prevent ionic currents through endogenous HEK-293 channels. The family  of  traces in the top of  panel A shows control on-gating currents evoked between -60 and 90 mV from  a holding potential of -100 mV and off-gating  currents at -100 mV; the lower traces of  panel A show the gating currents in 1 mM Ni2+. Control and treated traces, taken at the voltages indicated to account for  the gating shift,  have been superimposed in B to show that the kinetics of  the on- and off-  gating currents are not substantially affected  by Ni2+. C. The Qon- V  curve constructed from  6 cells by integrating the on-gating currents and normalizing with respect to the control Qmax confirms  that, although Ni2+ caused a ~ 10 mV rightward shift  of  the F,„ the Qmax did not decrease. Fitting to a Boltzmann function  gave control and treated F, values of  -6.8 ± 1.2 mV and 2.2 ± 0.8 mV, respectively, and values of  7.0 ± 1.4 mV and 9.2 ± 1.2 mV for  s. ' 3.3.7 Co2+ and Cd2+ , but not Mn2+, block Kvl.5 Other divalent transition metals that can bind to histidine include Cu2+, Fe2+, Co2+, Cd2+ and Mn2+. Because a precipitate formed  with Cu2+ and Fe2+, only the effects  of  Co2+, Cd2+ and Mn2+ could be compared to those of  Ni2+. The experimental protocol was the same as that described for  Figure 3.1 and was confined  to tests with a 0 mM K+0 solution. Figure 3.8 A shows a representative example of  the effect  of  Co2+ on currents evoked by the voltage protocol illustrated above the control responses. Switching from  the control solution to one containing 0.1 mM Co24 had no significant  effect  on the current but 10 mM Co2+ decreased the peak tail current following  a +60 mV pulse by more than 90%. Virtually complete recovery occurred after returning to the control solution. Fitting of  g-V  curves (not shown) to Equation (3.1) revealed that V V l shifted  by 11.4 ± 0.9 mV with 1 mM Co2+ and by 25.3 ± 1.3 mV with 10 mM Co2+. Neither concentration of  Co2+ significantly  affected  the slope factor  of  the g-V  curve (not shown). A fit  of  the Hill equation to the concentration-response data for  Co2+ (Figure 3.8 B) gave an estimate for  nH  of  1.3 ± 0.1 and a K D (1.4 + 0.1 mM) that was roughly 10-times larger than that " for  Ni2+ under the same recording conditions. 60 mV B. -50 mV -40 mV control 0.1 mM Co 10 mM Co recovery | 1 nA 50 ms 1.2 -i 1 . 0 -. 0.8 « I 0 6 -TO <U 0.4 -0.2 0.0 r~ 0.01 1—I I I Mill 1—I I I I Mil "TTTT 0.1 ^ 1 Co * concentration (mM) TTT]—i—m 10 Figure 3.8 Co2+0 also causes a concentration-dependent block of  Kvl.5 currents but is an order of  magnitude less potent than Ni2+. Shown in panel A are control and treated current traces evoked in 0 mM K+0 with the voltage protocol indicated above the control responses. In contrast to the -35% block of  the current with 0.1 mM Ni2+ (see Figure 3.2), 0.1 mM Co2+ had no effect.  However, with 10 mM Co2+ the maximum peak tail current amplitude decreased by -90%. As with Ni2+, the block by Co2+ was completely reversible. B. Concentration-response data obtained in 0 mM K+0, with each point representing the mean ± SEM of  measurements in 3-8 cells, were fitted  to Equation (3.2) which gave aK D of  1.4 ± 0.1 mM and an nH  of  1.3 ± 0.1. The effects  of  Cd2+ are not illustrated but closely resembled those of  Co2+. The K 0 was 1.5 ± 0.4 mM and the nH  was 1.3 ± 0.3. In 1 mM Cd2+ the V V i for  the g-V  relationship was shifted  rightward by 19.5 ± 1.2 mV. Of  the divalent cations we tested for  an ability to block Kvl.5, Mn2+ proved to be the least effective.  At 10 mM, the highest concentration used, gmCLX  was 73 ± 3% of  the control value. The mid-point of  the g-V  relationship was shifted  rightward by 21.5 ± 0.7 mV (n  = 3). 3.3.8 Co2+ and Zn2+ mimic the effect  of  Ni2+ on Kvl.5 inactivation Figure 3.9 A illustrates representative results of  the effect  of  10 mM Co2+ on inactivation and recovery from  inactivation using a voltage protocol identical to that described for  Figure 3.4. Again, a slowly rising phase of  current seen in 10 mM Co2+, K+0-free  medium (not shown) necessitated recording with 3.5 mM K+0. In 10 mM Co2+ both the onset of  and recovery from inactivation was comparable to that seen with 2 mM Ni2+ (Figure 3.4). In the four  cells studied with 10 mM Co2+, Tinacl and t r e c m e r y were 1.3 ± 0.1 s and 24.6 ± 1.7 s, respectively. As noted above, Zn2+ also causes a concentration and K+0-dependent inhibition of  Kvl.5 currents and for that reason its effects  on inactivation were also examined (Figure 3.9 B). Using a Zn2+ concentration of  2 mM, which is estimated to reduce gmax by 80-90% in 3.5 K+0, Tmac, was 1.64 ± 0.3 s and trecovery was 27.7 ±2.1 s (n  = 5 cells). Thus, a feature  which is shared by Ni2+, Co2+ and Zn2+ is an ability to substantially slow recovery from  inactivation and to modestly accelerate inactivation. In this regard at least these divalent cations are clearly distinct from  extracellular protons which, by comparison, accelerate inactivation to a far  greater extent (x m a a ~ 100 ms at pH 5.4) and slow recovery from  inactivation much less (Trecoven. ~ 4 s at pH 5.4). A . 10 mM Co2 ' B . 2 mM Zn2* Figure 3.9 Co2+ and Zn2+ mimic the effect  of  Ni2+ on macroscopic inactivation. A. Using a voltage protocol identical to that described for  Figure 3.4, the x maa with 10 mM Co2+ in 3.5 mM K+0 was well-fitted  by a single exponential with a time constant of  1.6 s. The fit  of  an exponential function  to the peak currents evoked by 50 ms test pulses following  the 5 s pulse to 50 mV gave a ^recovery of  21.8 s. B. With 2 mM Zn2,+ in 3.5 mM K^ vmaa was 1.86 s and the vrecovery was 29.3 s. Because Zn2+ slowed the activation rate the duration of  test pulses used to monitor recovery was increased to 200 ms. These data indicate a clear difference  in the effect  on Kvl .5 inactivation of divalent cations versus external protons (Figure 3.4 A). 3.4 Discussion As reported previously (Perchenet and Clement-Chomienne, 2001), external Ni2+ ions were shown to reversibly block human Kvl.5 currents (Figure 3.1). We have also shown here that Ni2+ block is affected  by [K+]0 (Figure 3.2). Thus, with 0 mM K+0 the K D for  the Ni2+ block is approximately 150 [iM whereas with 3.5 mM K+0 the KD increases to 400 [xM. The latter value is consistent with the K D of  570 [iM obtained with 5 mM K+0 in CHO cells (Perchenet and Clement-Chomienne, 2001). Increasing K+0 to 140 mM increased the K D to ~3 mM. The nH  of 1.2 to 1.6 derived from  concentration-response data in 0 mM to 5 mM K+0 (see also Perchenet and Clement-Chomienne, 2001) suggests that the block requires the binding of  at least two Ni2+ ions. In the study with Kvl .5 expressed in CHO cells the Ni2+ block was shown, regardless of the pulse frequency,  to develop slowly over a 2 to 5 minute period (Perchenet and Clement-Chomienne, 2001) despite the use of  a fast  drug application system. These data were interpreted to reflect  a large disparity in the association and dissociation rate constants for  Ni2+ binding to the closed state of  the channel. Although we agree that the Ni2+ block can occur from  the closed state, we found  no evidence for  a slow development of  that block (Figure 3.3). One possible interpretation of  the inhibition of  the Ni2+ block by K+0 is that it reflects  an interaction in the channel pore either by competition for  the same binding site or by an electrostatic effect  between separate Ni2+ and K+ binding sites. However, as noted with the block by Zn2+ and H+0 (Kehl et al., 2002), the block by Ni2+ shows no voltage dependence over a range of  voltages where the open probability is maximal (Perchenet and Clement-Chomienne, 2001). This observation supports the conclusion that the Ni2+ binding site is at least not in a region of  the pore that is within the electric field  and, by extension, that Ni2+ is not blocking by occlusion of the pore. The fact  that Kvl.5 currents are blocked by H+0, Zn2+ and Cd2+, whereas Shaker ) channels are not, also suggests a binding site external to the pore (e.g.,  in the turret) since in Kvl .5 and Shaker  there is complete homology from  the N-terminal end of  the pore helix to the GYG pore signature sequence. As is the case with the block of  Kvl.5 by H+0 and Zn2+, the sensitivity of  Kvl.5 channels to Ni2+ is greatly affected  (Figure 3.5) either by mutating H463 in the pore turret or by mutating R487, a residue in the outer pore mouth that has been shown in Shaker  channels to play a pivotal role in P/C-type inactivation. These results with the 463Q and 487V mutant channels, as well as the sensitivity of  the Ni2+ block to K+0 and the outcome of  other substitutions at position 463 (see below), are consistent with a model in which the binding of  Ni2+ to one or more H463 residues in the pore turret facilitates  an inactivation process that involves the outer pore mouth. Although this model is the same as that proposed for  the H+ 0 and Zn2+ block of  Kvl.5, there is not complete overlap of  the effects  of  these three metal cations. For example, the inactivation rate of  the residual currents is markedly different  with the divalent cations (Ni2+, Co2+, Zn2+) compared to H f 0 (Figure 3.4 and Figure 3.9). Thus, for  example, using concentrations that produce a similar degree of  block in 3.5 mM K+0, the residual currents inactivated roughly twenty times faster  with H+0 (pH 5.4) than with Ni2+ (Figure 3.4). Additionally, the shift  of  the midpoint of  the g-V  curve ^  the Qon-Vcurve  by Ni2+ was also much less than with either H+0 or Zn2+. Finally, the dramatic slowing of  the activation rate observed with Zn2+ (Zhang et al., 2001b) is not seen with either Ni2+ or H 0 . It seems unlikely, though we cannot disprove, that these differences  are due solely to the nature of  ligand co-ordination by the histidine residues in the turret. Particularly in the case of  H+0, the involvement of  additional binding sites seems likely. This is suggested by the fact  that although ShakerlR  channels are largely resistant to the conductance collapse in low pH, acidification  does accelerate current inactivation (Perez-Cornejo, 1999; Starkus et al., 2003). Furthermore, we and others have shown that manipulations that reduce the block of  Kvl.5 by metal cations do not affect  the gating shift  (Kehl et al., 2002; Trapani and Korn, 2003). From the data in Figure 3.5, it is also apparent that neither of  the outer pore mutations completely prevents current inhibition by Ni2+. Currents through the Kvl.5 H463Q construct decreased by -30% in 5 mM Ni2+ and, as with the Zn2+ block of  this mutant channel (Kehl et al, 2002), the nH  fitted  to the concentration dependence of  this block was quite small ( 0.5) suggesting the involvement of  a binding site and mechanism of  action that is different.  In the case of  the R487V mutant, the K D and the nH  for  the Ni2+ block with 0 mM K+0 are similar to that estimated for  wt Kvl.5 in 140 mM K+0. Paradoxically, neither of  these manipulations, increasing [K+] or mutating R487, substantially affects  the inactivation rate of  macroscopic currents carried by K+ during sustained depolarizations (Fedida et al., 1999). Although the latter observations might be construed as evidence against an involvement of  outer pore inactivation in the Ni2+ block, that is to say neither manipulation can be shown directly to affect  the current decay rate, an alternative explanation is that these manipulations inhibit an outer pore inactivation process occurring from  a closed state but are much less effective  against inactivation from  the open state. In this connection, a K+0-sensitive (K D ~ 0.8 to 10 mM) inactivation process occurring from,  a closed state has been suggested to account for  the decline of  the macroscopic conductance seen in fast-inactivating  ShakerlR  T449 mutants when the [K+]0 is decreased (Lopez-Barneo et al., 1993) and there is evidence in ShakerlR  supporting, not exclusively, " multiple, independent pathways of  which C-type is only one" (Yang et al., 1997). As with some of  the T449 mutations in Shaker,  there are mutations of  Kvl .5 H463 that can dramatically affect  outer pore inactivation. For example, mutants in which glycine (G) (Kehl et al., 2002) or arginine (R)) (Eduljee et al., 2003) is substituted for  H463 display rapidly / inactivating currents (r mac, = 35 to 75 ms) and, again as in the Shaker  T449X mutants, these rapidly inactivating mutants show a collapse of  the macroscopic conductance in 0 mM K+0 Furthermore, in the H463G mutant the conductance collapse in 0 mM K+0 is prevented by the R487V mutation (Trapani and Korn, 2003). The outcome of  these H463G and H463R mutations is significant  because it shows directly that the physico-chemical properties of  the residue at this position can dramatically affect  the time course of  open- (and closed?) state inactivation and thus offers  additional support for  the proposition that non-covalent chemical modification  of  H463 by the binding of  Ni2+, in addition to other metal cations, can affect  inactivation. Another significant  property of  the H463G mutant is that K+0 affects  the gmax with a K 0 of approximately 1 mM (Eduljee et al., 2003). This low millimolar K D is comparable not only to that estimated for  the fast-inactivating  ShakerlR  mutants (Lopez-Barneo et al., 1993) but to that obtained for  the relief  by K+0 of  the H+0 and Zn2+ block (Kehl et al., 2002; Zhang et al., 2001b). A detailed study of  the K+0-dependence of  the Ni2+ block was not undertaken here. However, using the K D of  the Ni2+ block in zero and 3.5 mM K+0, and assuming, for  simplicity, a competitive interaction, the K D for  the relief  of  the block by K+0 is calculated to be ~ 1.5 mM. A consistent pattern that emerges from  these studies, whether it is the spontaneously-occurring conductance collapse in ShakerlR  and Kvl.5 mutant channels or the metal ion-induced block/conductance collapse in wt Kvl.5, is that inhibition of  the conductance loss occurs with low millimolar K+0 concentrations and that this inhibition occurs in the absence of  a change of the inactivation rate measured during depolarizing pulses. This implies that there is an outer pore inactivation process, perhaps that occurring from  the closed state, that is much more sensitive to K+0 and, we suggest, given that its inactivation rate is not distinguishable from  wt Kvl .5 channels, that in Kvl .5 the R487V mutation selectively affects  this same inactivation process. With the fast-inactivating  ShakerlR  mutants, Lopez-Barneo et al. (1993) remarked that the tendency for  the conductance to collapse (inactivate from  the closed state?) in 0 mM K+0 is associated with fast  current inactivation. This correlation also applies to Kvl.5 H463G where the inactivation rate is some 20-fold  faster  than in wt Kvl .5 channels but it is much less evident with the Ni 2 + , Co2+ and Zn2+ block where the inactivation rate of  residual currents is only ~ 2-fold faster  than in controls (Figure 3.4 and 3.9). Particularly in view of  the low concentrations of  K+0 needed to relieve the metal cation block, a question that inevitably arises is whether the external K+ binding site can also be populated by outward K+ flux  through the open channel. Though it has not been studied for  Ni2+ block, our recent finding  (Zhang et al., 2003) of  virtually identical K Ds for  the block by H+0 of outward K+ or Na+ currents argues against a contribution of  outward K+ currents in the block relief.  One explanation for  this apparent absence of  an effect  of  K+ efflux  through the open pore is that K+ ions at the outer pore mouth rapidly equilibrate with the external solution. Alternatively, if  Ni2"-bound channels are inactivating from  a closed state, or if  the open time is very brief  (Zhang et al., 2003), there would be no opportunity for  block relief  by outward K+ currents. A comparison of  currents from  one channel outside-out patches (Figure 3.6) prior to and following  the application of  0.5 mM Ni2+ showed: 1) that open channel current (/) at 100 mV did not change; 2) that the open probability (PJ  during 300 ms sweeps containing channel activity was not changed; and, 3) channel availability (AO, decreased from  a value of  ~ 0.9 in the control to ~ 0.4 during treatment. Although a detailed analysis and comparison of  open and closed time behaviours have not yet been done, these preliminary data are consistent with a model in which Ni2+ binding facilitates  a reversible transition from  an available to an unavailable (closed-state inactivated?) state. Gating current analyses (Figure 3.7) showed that, as with H+0 (Kehl et al., 2002), Ni2+ did not affect  Qmax. This finding  rules out the possibility that the prevention of  one or more of  the transitions in the activation pathway accounts for  the Ni2+-induced decrease of  gmax. Ni2+ treatment also caused a ~ 10 mV shift  of  the Qon-V  curve but this was much less than the 50-60 mV shift  seen with H+ 0 or Zn2+ (Kehl et al., 2002; Zhang et al., 2001a). As noted above, it is not clear if  this disparity in the gating shift  reflects  differences  in ligand coordination with H463 residues or if  the larger shift  with H+0 and Zn2+ reflects  interactions with additional binding sites. Transition metal ions that have now been shown to block Kvl .5 currents are Zn2+, Cd2+, Ni2+ and Co2+ (Figure 3.8). For the first-row  transition metals the rank order for  the inhibition of Kvl.5 in 0 mM K+0 is Zn2+(K D -0.07 mM) > Ni 2 + (£ D -0.15 mM) > Co2+ (K 0 -1.4 mM) > Mn2+ (7Cd>10 mM) and, as such, is in accord with the Irving-Williams order (Glusker, 1991). Zn2+, Ni2+ and Co2", which are intermediate Lewis acids, are known to bind to the thiolate side group of cysteine and the imidazole nitrogen of  the histidine. Zn2+ is also able to bind to carboxylate and carbonyl oxygen atoms. Cd2+, a second row transition metal, is a soft  Lewis acid and typically has a higher affinity  for  a soft  base such as the thiolate ion. Preliminary work with the H463C mutant shows a sensitivity to block by Cd2+ that is greater than for  wt Kvl .5. In Cav2.3 (alE) channels, external Ni2+ causes, in addition to a rightward shift  of  the g-V curve, a reduction of  the slope conductance with an estimated K t of  300 [iM (French et al., 1996). The blocking reaction appears to be bimolecular and is also affected  by the type of  permeant ion (e.g.,  Ca2+ versus Ba2+). It was suggested that the Ni2+ block of  Cav2.3 reflected  changes of permeation due to direct occlusion of  the pore in addition to a possible change of  the permeant ion concentration at the pore mouth. In voltage-gated K+ channels, divalent cations have proved to be useful  probes of  gating and permeation. However, whereas Zn2+ and Cd2+ have been studied in some detail (Gilly and Armstrong, 1982; Spires and Begenisich, 1994), Ni2+ has been used somewhat sparingly. In HERG K+ channels external Ni2+, as well as Cd2+, Co2+ and Mn2^, increased the maximum current amplitude, an effect  that was imputed to an alteration of inactivation gating (Paquette et al., 1998). Interestingly, in HERG channels mutations at a number of  sites in the S5-P linker can dramatically alter inactivation (Liu et al.,'2002), a finding that underscores the findings  with Kvl.5 that, either by substitution through point mutation, or by chemical modification  through ligand binding, residues in this region can profoundly  influence the rate and extent of  one or more inactivation processes occurring at the outer pore mouth. / 3.5 References Adda, S., B. K. Fleischmann, B. D. Freedman, M. Yu, D. W. Hay, and M. I. Kotlikoff.  1996. Expression and function  of  voltage-dependent potassium channel genes in human airway smooth muscle. J. Biol. Chem. 271:13239-13243. Baker, O. S., H. P. Larsson, L. M. Mannuzzu, and E. Y. Isacoff.  1998. Three transmembrane conformations  and sequence-dependent displacement of  the S4 domain in Shaker  K+ channel gating. Neuron 20:1283-1294. Chen, F. S. and D. Fedida. 1998. 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Mechanisms of  the inhibition of  Shaker  potassium channels by protons. Pflugers  Arch. 447:44-54. Steidl, J. V. and A. J. Yool. 1999. Differential  sensitivity of  voltage-gated potassium channels Kvl.5 and Kvl.2 to acidic pH and molecular identification  of  pH sensor. Mol. Pharmacol. 55:812-820. . Tamkun, M. M., K. M. Knoth, J. A. Walbridge, H. Kroemer, D. M. Roden, and D. M. Glover. 1991. Molecular cloning and characterization of  two voltage-gated K+ channel cDNAs from human ventricle. FASEB J. 5:331-337. Trapani, J. G. and S. J. Korn. 2003. Effect  of  external pH on activation of  the Kvl.5 potassium channel. Biophys. J. 84:195-204. Wang, Z., J. C. Hesketh, and D. Fedida. 2000a. A high-Na+ conduction state during recovery from  inactivation in the K+ channel Kvl.5. Biophys. J. 79:2416-2433. Wang, Z., X. Zhang, and D. Fedida. 2000b. Regulation of  transient Na+ conductance by intra-and extracellular K+ in the human delayed rectifier  K+ channel Kvl.5. J. Physiol. 523:575-591. 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Single channel analysis reveals different  modes of  Kvl.5 gating behaviour regulated by changes of  external pH 4.1 Introduction Kvl .5 is a Shaker-related,  voltage-gated potassium channel encoded by the gene KCNA5. In the human heart it mediates the ultra-rapid delayed rectifier  current (IKur) involved in repolarizing the atrial action potential (Fedida et al., 1993; Feng et al., 1997). In a previous study, we reported that low external pH inhibits macroscopic Kvl.5 currents and causes a depolarizing shift  of  the conductance-voltage (g-V)  relationship without substantially affecting activation kinetics (Kehl et al., 2002). Based on structure-function  analyses, a histidine residue (H463) in the pore turret (S5-P linker) has been suggested to form  part of  the site to which external protons bind to produce these effects.  Current inhibition by external protons can be attenuated either by raising the external [K+] or by mutating an arginine residue near the outer pore mouth to valine (Kvl.5 R487V) (Kehl et al., 2002), which is equivalent to the T449V mutation in Shaker.  These latter manipulations also influence  an inactivation process involving the selectivity filter  which is best characterized in Shaker  channels where it is known as slow, P/C-type inactivation (Lopez-Barneo et al., 1993) but is often  referred  to below as outer pore inactivation. Together with the observation that the inactivation rate of  residual currents increases in a pH-dependent manner with a K D similar to that for  the current inhibition (Kehl et al., 2004; Steidl and Yool, 1999), these findings  suggested a possible link between the proton-induced inhibition of  Kvl.5 and outer pore inactivation. Pore occlusion as the basis for  the effect A version of  this chapter has been published. Kwan, D. C. H., Fedida, D., and Kehl, S. J. (2006) Single channel analysis reveals different  modes of  Kvl.5 gating behaviour regulated by changes of  external pH. Biophys. J. 90(2): 1212-1222. -has been considered but deemed unlikely given that Na + currents, reflecting  ion flux  through the inactivated pore, are little affected  by decreasing pH (Zhang et al., 2003). To get a clearer picture of  the mechanistic basis for  the current inhibition by external protons in Kvl .5, currents through single Kvl .5 channels were studied. Our results show that single channel current (/) is not affected  by changing the extracellular pH and indicate that a change of  P0 is the basis for  the effect.  Consistent with the acceleration of  inactivation of residual macroscopic currents at low pH, there was, in sweeps showing channel activity, a decrease of  the mean burst duration and an increase of  the mean interburst duration, suggesting a stabilization of  inactivation. Within bursts, channel gating between the open state and closed states outside of  the activation pathway was only slightly affected,  if  at all. The main cause of the pH-induced current inhibition of  Kvl.5 was an increased probability of  a gating mode where the channel was unavailable for  activation. 4.2 Materials and Methods 4.2.1 Cell preparation A stable mouse cell line, Itk,  expressing hKvl .5 channels at a low density was generated by transfection  with the cDNA of  the human potassium channel Kvl.5 (/?Kvl.5) subcloned in the gentamicin-resistant gene-containing pcDNA3 vector using methods described previously (Wang et al., 2000). Cells were maintained in minimum essential medium (MEM) supplemented with 10% fetal  bovine serum, 1% penicillin-streptomycin, and 1% gentamicin and incubated at 37°C in an atmosphere of  5% C02 in air. Cells were dissociated and plated onto cover slips for experimental use within 1-3 days. All tissue culture supplies were obtained from  Invitrogen (Burlington, Ontario, Canada). 4.2.2 Electrophysiology Single-channel current (/) and whole-cell current (7) were recorded in either the cell-attached or, in experiments where responses at two different  pHs were compared, the outside-out patch configuration.  For cell-attached recordings, the bath solution was assumed to depolarize the cell to 0 mV; it contained (in millimolar) 140 KC1, 3.5 NaCl, 2 CaCl2, 1 MgCl2, 5 glucose, 10 HEPES and was adjusted to pH 7.4 with NaOH; the patch pipette (external) solution contained 140 NaCl, 3.5 KC1, 2 CaCl2, 1 MgCl2, 5 glucose and was adjusted either to pH 7.4 with 10 HEPES/NaOH or to pH 6.4 with 10 MES/NaOH. For recordings from  an outside-out patch or in the whole-cell configuration,  the bath (external) solution contained 140 NaCl, 3.5 KC1, 2 CaCl2, 1 MgCl2, 5 glucose and was adjusted to pH 7.4 (10 HEPES/NaOH), 6.4 (10 MES/NaOH), or 5.9 (10 MES/NaOH); the patch pipette (internal) solution contained 130 KC1, 10 EGTA, 10 HEPES, 4.75 CaCl2, 1.38 MgCl2and was adjusted to pH 7.4 with KOH. The free  [Ca2+] and [Mg2+] were calculated to be 50 nM and 1 mM, respectively, using the MaxChelator program (Stanford University, Stanford,  CA, USA). In subsequent text the reference  to pH means the extracellular pH. All chemicals were purchased from  Sigma-Aldrich (Mississauga, ON, Canada). Voltage commands and current measurements were made with an EPC-7 patch clamp amplifier  connected to an ITC-18 digital interface  (Instrutech, Port Washington, NY, USA) and controlled, by Pulse software  (HEKA Electronik, Germany). Patch electrodes pulled from  thin-walled borosilicate glass (World Precision Instruments, FL, USA) had a resistance measured with recording solutions of  8-25 MQ. Current signals were low-pass filtered  at 3 kHz (-3dB, 8-pole Bessel, NPI Electronics, Tamm, Germany) and digitized at 10 kHz with a 16-bit A/D converter. Cells were held at -80 mV at rest. A junction potential of-4  mV was compensated in the voltage measurements in the outside-out patch configuration;  no junction potential correction was made for  data acquired in the cell-attached recording mode. 4.2.3 Data analysis Outside-out recordings were made at room temperature (20-25°C) and involved exclusively one-channel patches identified  as such by the absence of  overlapping open events in pH 7.4 solution at a potential, usually 100 mV, where the open probability was high. In cell-attached recordings it was not possible to change the pH of  the solution in the recording electrode and although patches containing one channel could be identified  unequivocally at pH 7.4 this was not the case at pH 6.4 where the open probability was lower. Single-channel records were analysed with TAC and TACFit software  (Bruxton, Seattle, WA, USA) after  digital filtering  at 1 kHz. Capacitive currents were removed by subtracting the average of  sweeps obtained at the same voltage that showed no channel activity (i.e.,  blank or "null" sweeps). Given a combined analogue and digital filter  frequency  of  950 Hz, the dead time of  the system was calculated to be 0.3 ms using the formula  0.253If  where/represents the overall combined analogue and digital filter  cut-off  frequency  in Hz (Hoshi et al., 1994). Half-amplitude  threshold analysis, with a rise time of  0.3 ms, was used to detect events and generate idealized records from  which dwell time histograms and ensemble time courses were constructed. Events with durations shorter than the dead time were censored. Although there was evidence for  a conductance substate in some traces (e.g.,  Figure 4.1), this phenomenon was not analysed in detail due to its relatively infrequent occurrence. For the analysis of  gating kinetics a burst was defined  as a group of  brief  open and closed events followed  by a closed interval of  20 ms or more. There was no correction for missed events in the analysis of  dwell time histograms that were fitted  to exponential functions. The choice of  the number of  components used for  fitting  was based on a maximum likelihood technique, in which the least number of  components with a significant  improvement was used (Saftenku  et al., 2001). A value of  twice the difference  in log-likelihood (i.e.,  2| LL, - LL21) being greater than the tf  value with a given degrees of  freedom  (v) is considered significant.  The number of  degrees of  freedom  is equal to the difference  in free  parameters between the models (Saftenku  et al., 2001). Averaged results are expressed as the mean ± SEM unless otherwise stated. Results for  fitting  histograms and the Hill equation are given as the mean ± SD. Statistical tests (Student's /-test, ANOVA) were performed  with Jmp In Software  (SAS Institute, Cary, NC, USA). A probability less than 0.05 was considered significant. A "runs analysis" was performed  (Gibbons, 1985) to check the randomness of  the occurrence of  sweeps in which the channel was available or unavailable (Horn and Vandenberg, 1984). Data were obtained from  one-channel cell-attached patches stepped to 100 mV for  150 ms with a cycle length of  3, 5, 10, or 15 seconds. A run (R)  is defined  as a sequence of  sweeps showing similar gating behaviour which, for  the purposes of  this paper, means either a sequence of  null sweeps in which the channel was in the unavailable mode (U)  or a sequence of  active (A) sweeps in which at least one open event was detected. When the number of  trials is > 40, a normalized statistical value (Z)  can be calculated by the equation, R - 2np(\  - p) Z = — 2*J~np(\  - p) ( 4 J ) where R is the number of  runs, n is the number of  trials or sweeps, and p is the probability of  the event. The expected number of  runs is 2np(l-p).  A smaller than expected number of  runs implies clustering and generates values of  Z greater than 0. Conversely, a value of  Z  less than 0 indicates a tendency to alternate between null and active sweeps (Horn et al., 1984). The value of  Z was compared to the normal distribution to determine the statistical significance;  a value greater than 1.64 indicated behaviour that was non-random and clustered (p  < 0.05, one-tail test (Plummer and Hess, 1991; Nilius, 1988)). Contingency tables (2x2) were constructed to determine whether, at pH 6.4, there was a correlation between a sweep ending in an inactivated or non-inactivated state and the following sweep being active or null. Chi-square {tf)  analyses were performed  using the equation, where f  and /^represent, respectively, the experimental and expected frequency  for  row i and column j (Zar, 1984). A £ value greater than 3.841 is considered significant  (p  < 0.05, v= 1). 4.3 Results Representative, consecutive unitary current traces recorded at pH 7.4 from  an outside-out patch during a 1-s depolarizing pulses at 100 mV with 3.5 mM K+0 are shown in Figure 4.1 A. As noted above, Kvl.5 exhibits conductance substate behaviour (marked by arrows) but these were not systematically analysed. As with ShakerlR  channels (Hoshi et al., 1994; Schoppa and Sigworth, 1998a), at this voltage and pH the latency to first  opening was consistently very brief (< 1 ms) and the channel rapidly flickered  between short-lived open and closed states. Save for  a few  traces in which the channel entered a long-lived closed state presumed to be due to outer pore inactivation, the open probability was high for  the duration of  the depolarization. At pH 7.4, null sweeps occurred rarely, e.g., Figure 4.4 A. Fitting the cumulated all-points amplitude histogram (Figure 4.1 A bottom panel) with a three-component Gaussian function  gave a single channel current of  1.7 ± 0.2 pA for  the main conductance state, which is comparable to previously published results for  /zKvl.5 (Fedida et al., 1993). The intermediate component is (fa-fa) 2 (4.2) <j due to a conductance substate and incomplete opening/closing events. Figure 4.1 B shows unitary currents from  the same patch as in Figure 4.1 A using the identical voltage protocol but after  switching to pH 6.4. As at pH 7.4, flickery  channel behaviour and conductance substates (arrows) were observed. There was no obvious difference  in the probability of  substates at pH 7.4 and 6.4. However, more sweeps showed the channel entering a long-lived closed state before  the end of  the voltage step, and the proportion of  null sweeps was higher. Fitting of  the all-points amplitude histogram at pH 6.4 gave a single channel current of 1.7 ± 0.2 pA. Figure 4.1 C and D shows representative current traces recorded from  another outside-out patch at pH 5.9 and during recovery at pH 7.4 using the same voltage protocol as in Figure 4.1 A. At pH 5.9, the number of  null sweeps was even higher; there was no evidence of  channel activity in eleven of  the twelve sweeps shown. The absence of  activity in the first  sweep suggests that depolarization-induced inactivation is unlikely to account for  the null sweeps. In addition, the series of  10 consecutive null sweeps observed over a period of—150  s is not likely to be due to a failure  to recover from  depolarization-induced inactivation since the recovery of  residual macroscopic currents from  inactivation at pH 5.4 with 3.5 mM K+0 is = 4 s (Kwan et al., 2004). This long period of  inactivity is not likely to be due to entry into a defunct  state since K+ was present in both the intracellular and extracellular solution and recovery did not require a long depolarization pulse (Loboda et al., 2001). Recovery responses were obtained after  switching the bath solution back to pH 7.4, demonstrating that the dramatic decrease of  channel availability at pH 5.9 was also not due to the spontaneous loss of  the channel from  the patch. Furthermore, the single channel current (/) did not change between pH 5.9 and pH 7.4 (Figure 4.1 C and D). A pH 7.4 B pH 6.4 pH 5.9 D Recovery (pH 7.4) ttWWWPWl'MW y^UMUi ij fWWIVilfiifW  _ _ M M I <... .1 >1 • i W I M L -flWWIWWPII i twwwfi i f l i iMJUflllMIUMi (P.i.WiUflww uMMUtttiMMi pWml^ lfBwlwll pWPWBWIntlWJI iai i i i iokM PflTOwWiHI IHWFIWilWIi IWWW^IIJip (PlnlllRllWf m m m m 2 p A 200 ms 6000 -» 4000 -O 2000 - J 12000 -8000 -4000 • i r 0 1 2 Amplitude (pA) i 20000 -15000-10000-u ^ d jL-j o - J i i—i—r 0 1 2 Amplitude (pA) 0 1 ' 2 Amplitude (pA) I 0 1 2 Amplitude (pA) Figure 4.1 The single channel current amplitude of  Kvl.5 does not change between pH 7.4, 6.4, and 5.9. A. Representative unitary current through a one-channel, outside-out patch at pH 7.4. Current traces were evoked by 1 s depolarizing pulses at 100 mV from  a holding potential of -80 mV every 15 s. Kvl.5 unitary currents show flickering  behaviour. The failure  of  the channel to open (e.g.  last trace in pH 7.4 column) was an infrequent  observation at this pH. The corresponding cumulative all-points histogram based on traces in A is shown at the bottom of  the panel. Fitting the histogram to a three-component Gaussian function  gave a value of  1.7 pA for the major conducting level. B. Unitary current from  the same outside-out patch using the same voltage protocol after  switching to external solution at pH 6.4. The open channel current does not change but there are more null sweeps. The all-points amplitude histogram shown at the bottom gave a single channel current of  1.7 pA for  the major conducting level. The higher proportion of  the non-conducting points reflects  the higher proportion of  null sweeps and the more frequent  termination of  active sweeps by a long-lived non-conducting state. C. Unitary current from  a different  outside-out patch using the same voltage protocol as in A but with an external solution at pH 5.9. Channel activity is observed in only one of  the twelve sweeps. This dramatic decrease of  channel activity was not due to the loss of  the channel from  the patch since recovery of  activity was obtained after  returning to pH 7.4 solution (D). The all-points histograms at the bottom of  panel C and D give a value of  2.0 pA for  the mean open channel current. Similar results were observed in 2 other patches. Together, these results show that decreasing external pH decreases the availability but not the single channel current of  Kvl.5. Analysis of  single channel currents over a range of  voltages confirmed  that the single channel conductance is not affected  by changing pH. Representative unitary currents recorded at 0 to 100 mV in 20 mV increments at pH 7.4 or 6.4 from  different  outside-out patches are shown in Figure 4.2 A. The graph in panel B shows the i-Vrelationship  based on data from  3 cells, derived from  all-points histograms as described for  Figure 4.1, at pH 7.4 (unfilled  circles) and pH 6.4 (filled  circles).  A line fitted  to the data gave a slope conductance (y  mean ± SD) of  11.8 ± 0.6 pS at pH 7.4 and 11.3 ± 0.8 pS at pH 6.4, which were not significantly  different.  This finding,  together with the data of  Figure 4.1, allowed us to reject the hypothesis that the reduction in macroscopic current by acidification  is due to the reduction of  the single channel conductance arising either by a change of  the permeation pathway or by occlusion of  the pore. Instead, the results of  Figure 4.1 and 4.2 indicate: 1) that the low pH-induced decrease of  macroscopic current must arise solely by an effect  on channel gating; and, 2) that depolarization-induced inactivation is unlikely to play a significant  role in the mechanism responsible for  the null sweeps. Macroscopic currents and the ensemble current behaviour generated from  idealized records, obtained in each case from  channels expressed in Itk cells, are shown for  comparison in Figure 4.3 A and B, respectively. Macroscopic currents evoked by a 1 s pulse to 100 mV at pH 7.4, 6.4 and 5.9 showed a pH-dependent decrease of  the peak current along with an increased rate of  inactivation (Figure 4.3 A; see inset of  normalized currents and figure  legend for  numerical values) as reported previously (Kehl et al., 2002). The reductions in peak currents are comparable to the reduction in macroscopic conductance measured from  Kvl.5 expressed in A pH 7.4 pH 6.4 B 2 .0 -, 100 mV 80 mV !  ^wififfli w.., 40 mv ^^ jmmmmm  fflV 0.5 -20 mv m m r n m r n m . 0 m V iww«mnnr<i wrrir mm |2 pA o.o 200 ms 20 40 60 80 100 Voltage (mV) Figure 4.2 Open channel current-voltage relationship showing the single channel conductance does not change with pH. A. Representative unitary currents at voltages between 0 and 100 mV in 20 mV increments at pH 7.4 and 6.4. All traces shown were digitally filtered  at 1 kHz. B. Fitting a line to the i-V  relationship at pH 7.4 (unfilled  circles)  and pH 6.4 (filled circles)  gave a slope conductance (mean ± SD) of  11.8 ± 0.6 pS and 11.3 ± 0.8 pS, respectively. These values for  the slope conductance were not significantly  different. HEK-293 cells with similar recording conditions (5 mM K+c; Figure 4.3 of  (Kehl et al., 2002)) and that measured from  the ensemble current (Figure 4.3 B; see figure  legend for  details). Furthermore, the time courses of  inactivation measured in the whole-cell configuration  at different  pH also agreed well with that measured from  the ensemble currents. This correlation shows that a change of  P0 nicely accounts for  the effect  of  pH on the amplitude and kinetics of macroscopic currents. Inspection of  the single channel responses reveals that this low pH-induced decrease of  PQ arises for  two reasons: 1) channel availability decreases; and, 2) the burst duration decreases. Of  these two actions, the influence  of  pH on channel availability was much greater and is considered first. 4.3.1 pH affects  channel availability Figure 4.3 C shows a plot of  channel availability, defined  as the proportion of  sweeps 4 nA B pH 5.9 0.2 P„ 200 ms 200 ms o Relative (l«) • , Availability A , Relative g m 0 , (HEK 293) Figure 4.3 Current behaviour at the macroscopic level and the ensemble average of  unitary current of  Kvl.5 at pH 7.4, 6.4, and 5.9 are qualitatively similar. A. Representative, superimposed macroscopic currents at pH 7.4 (top trace), pH 6.4 (middle trace), and pH 5.9 (bottom trace) evoked with a 1 s depolarizing pulse to 100 mV from  a holding potential of -80 mV. Peak current was reduced by about 22% at pH 6.4 and 81% at pH 5.9. Fitting the currents at pH 7.4, 6.4, and 5.9 to a single exponential function  gave mean inactivation time constants of  558 ± 75 ms, 556 ± 61 ms, and 346 ± 46 ms, respectively. Normalized currents are shown in the inset to better illustrate the acceleration of  inactivation. B. Ensemble open probability generated from  idealized single channel records at pH 7.4, 6.4, and 5.9. The ensemble behaviour reproduces the changes of  the peak amplitude and kinetics observed in the macroscopic currents. Compare to that at pH 7.4, the peak ensemble current was reduced by 34%) at pH 6.4 and 82% at pH 5.9. A single exponential fitted  to the ensembles gave time constants of  548 ms, 401 ms, and 197 ms for  pH 7.4, 6.4, and 5.9, respectively. Normalized ensembles are shown in the inset. C. A reduction in channel availability accounts for  the reduction of  peak macroscopic current by external H+. Channel availability (filled  circles)  agrees well with the normalized relative gmca recorded in HEK-293 cells (unfilled  triangles)  or the normalized peak macroscopic current in ltk~  (unfilled  circles).  Availability is defined  as the proportion of  sweeps with one or more open events. Data from  HEK-293 cells were obtained from  our previous study (Kehl et al., 2002). Briefly,  whole-cell currents were recorded from  a series of  300-ms depolarizing steps at -50 to +60 mV, and the instantaneous tail currents at -50 mV were analysed to give the relative gm a x values at different  pH. The composition of  the bath and pipette solutions were identical to that listed in Materials and Methods except the bath solution contained 5 mM K+ and 138.5 Na+. Fitting the whole-cell data to the Hill equation gave a p K H  of  6.2 ± 0.2 with a Hill coefficient  of  1.6 ± 0.4 (dashed  line) for  the reduction in relative gmax. Fitting channel availability to the Hill equation (solid  line) gave apK H  of  6.4 ± 0.2 with a Hill coefficient  of  1.9 ± 0.2. 139 with at least one open event, against pH (filled  circles).  Availability changed from  0.97 ± 0.02 at pH 7.4, to 0.64 ± 0.6 at pH 6.4 and to 0.17 ± 0.03 at pH 5.9 (n  = 3 -12 patches). Included for comparison in Figure 4.3 C is the relationship between pH and either the normalized pH 7.4 1.0 -, 0.8 0 . 6 -> 0.4 0.2 0 . 0 T 60 90 Sweep Index pH 6.4 ~r . 150 25 2 0 -15-10 5 -0 -D 25 -20-§ 15-\ t = 2.8 ± 0.4 sweeps o nn l 1 l 0 10 20 30 # of consecutive null sweeps 1 0 -5 0 T = 1.9 ± 0.3 sweeps i—1—i—1—r^—r 0 10 20 30 # of consecutive active sweeps Figure 4.4 Modal gating of  Kvl.5 at different  pHs. A. A diary plot of  Kvl.5 constructed by plotting the open probability per sweep for  150 consecutive sweeps depolarized at 100 mV for 150 ms every 3 s at pH 7.4 with a cell-attached patch. In this example, only one sweep (#79) shows mode U  gating. B. Diary plot from  another cell-attached patch using an identical protocol but at pH 6.4. Mode A (available) and mode U  (unavailable) gating appear in clusters, as suggested by runs analyses (see text). C. Frequency histogram of  the number of  consecutive sweeps showing mode U  gating at pH 6.4. The length of  runs with mode U  gating was pooled from  7 patches, and the resulting histogram was fitted  to a single exponential distribution to give a time constant of  2.8 sweeps (8.4 s). D. Frequency histogram of  the number of  consecutive sweeps with mode A gating at pH 6.4. Fitting the histogram to a single exponential distribution gave a time constant of  1.9 sweeps (5.7 s). macroscopic peak current at 100 mV for  Kvl .5 expressed in ltk~  cells (unfilled  circles) or the relative macroscopic conductance data taken from  our study using Kvl .5 in HEK-293 cells (;unfilled  triangles;  Kehl et al, 2002). The similarity of  these three relationships indicates that a change of  availability is the primary cause of  the inhibition of  Kvl .5 currents by extracellular protons and is consistent with our previous conclusion (Kehl et al, 2002) that the pH-induced increase of  the depolarization-induced inactivation rate alone is an insufficient  explanation. The decrease in availability may result from  an increase in the number of  random null sweeps or from  a modal gating scheme in which the null (or active) sweeps are clustered .together. To determine if  gating was modal, unitary currents through cell-attached patches were recorded using a pipette solution buffered  to pH 7.4 or 6.4, and the voltage protocol consisted of a 150 ms pulse at 100 mV applied at an interval of  3 s. The attached cell was depolarized by a high [K+] bath solution and its resting potential was assumed to be 0 mV. It was also assumed that the absence of  more than one (main conductance) open level indicated that the patch contained just one channel. A representative diary plot of  Pa per trace at pH 7.4 {panel  A) and 6.4 {panel  B) is shown in Figure 4.4. At pH 6.4, the diary plot indicates a clear tendency for alternating periods of  null and active traces and for  that reason a runs analysis was done (see Materials and Methods) to determine if  sweeps showing similar channel behaviour were clustered, i.e., that gating was modal. Sweeps were labelled as either unavailable (U)  for  null traces or available (A)  for  traces showing one or more openings of  the channel. Using Equation 4.1, the Z values were calculated to be 5.0 ± 1.3 (range 1.77-19.6; « = 6) at pH 7.4 and 6.3 ± 1.0 (range 2.85-9.17; n = 7) at pH 6.4. Both values indicate a significant  clustering (p < 0.001) of active and null sweeps. To estimate the average lifetimes  of  epochs of  U  and A sweeps, histograms were constructed from  the numbers of  consecutive sweeps with either type of  gating. Data at pH 7.4 or 6.4 were combined from  6 and 7 one-channel, cell-attached patches, respectively (Plummer and Hess, 1991) and represent a total of  1502 sweeps (1308 A sweeps, 194 U  sweeps) at pH 7.4 and 843 sweeps (385 A sweeps, 458 U  sweeps) at pH 6.4. Despite the large number of  sweeps recorded at pH 7.4, there were only 93 runs (49 in mode A, 44 in mode U)  and the histogram for mode A was inconclusive due to a large proportion of  very long runs (not shown). Fitting the histogram for  mode U  to a single exponential gave a time constant of  0.76 ± 0.06 sweeps (or 2.3 ± 0.2 s given a stimulus period of  3 s; not shown). On the other hand, 385 runs (194 in mode A, 191 in mode U)  were recorded with pH 6.4, and histograms for  mode U  and mode are shown in Figure 4.4 C and D, respectively. A log-likelihood test revealed the distribution was better fitted by a single exponential function  (difference  in log-likelihood < 0.2, not significant,  v=2) with a time constant of  2.8 ± 0.4 sweeps (8.4 ± 1.2 s) for  mode U  and 1.9 ± 0.3 sweeps (5.7 ± 0.9 s) for mode A. Together, these results suggest decreasing pH promotes mode U  gating while increasing pH favours  mode A gating. If,  for  simplicity, we assume that the transition between the two modes is first  order, i.e., U<r>  A k- \ then k,  and k_,  are equal to the reciprocal to the mean lifetime  of  U  and A, respectively. At pH 6.4, the estimates for  k,  and k_,  were 0.12 s"1 and 0.18 s'\  respectively. These numbers translate to a probability for  A of  0.4 at pH 6.4. This analysis was based on the assumptions that: 1) a channel could make at most one transition during a 3 s period; and, 2) the probability of switching between the two gating modes during a 150 ms pulse to 100 mV was negligible. The first  assumption is probably valid since changing the interpulse interval to either 5 or 10 s did not alter the outcome of  the runs analysis; however, when the interpulse interval changed to 15 s the grouping of  U  and A sweeps showed no statistical tendency to cluster (not shown). The second assumption seems valid for  the transition from  U to A because very few  traces (less than 1%) had a latency to first  opening longer than a few  ms. However, the long-lived closure sometimes observed before  the end of  a voltage pulse could be due to either a depolarization-induced inactivation or a transition to mode U. An important question regarding mode U  is whether it simply reflects  depolarization-induced inactivation. With a 1 s depolarization to 100 mV at pH 6.4 there was, compared to pH 7.4, a higher probability of  entering a long-lived closed state (Figure 4.1 B) but this was not invariably linked to the channel being unavailable on the next sweep evoked 15 s later. In Figure 4.1 B, for  example, five  of  eleven active sweeps terminate in a long-lived closed state, which is presumed to reflect  depolarization-induced inactivation, and four  of  those five  sweeps are followed  by an active sweep. Statistical analysis of  a 2x2 contingency table (sweep ending in inactivation versus sweep active at end) x (next sweep null versus next sweep active) showed there was no correlation between a pulse ending in an inactivated state.and the next sweep being blank (27 pairs, j 2 value = 2.41,/? > 0.1). With shorter depolarizations (150 ms), such as those used for  Figure 4.4 B, the probability of  inactivating before  the end of  the pulse was low, as indicated by the similarity of  the P0 per active trace at either pH, but clustering of  null sweeps persisted. Similarly, runs analysis at pH 6.4 with 20 ms depolarizing pulses to 100 mV delivered at 0.33 Hz (not shown) showed clustering of  null sweeps (Z= 12.0 ± 3.5; n = 6; not shown). These results indicate that mode U  gating is not simply due to depolarization-induced inactivation. This is consistent with our previous finding  that decreasing external pH has no effect  on the maximum gating charge (Q max) mobilized during activation (Kehl et al., 2002), in contrast to what is expected were the channels in the depolarization-induced C-type inactivated ' state. The gating charge data also imply that mode U  gating includes voltage-sensitive transitions between several non-conducting states. However, the data do not allow us to say whether transitions between mode U  and the depolarization-induced inactivated state are possible. 4.3.2 pH does not substantially affect  intraburst behaviour As noted above, inspection of  channel behaviour during active sweeps at either pH 7.4 or 6.4 revealed bursts comprised of  brief  closures and brief  openings. Dwell time histograms of open (conducting) and closed (non-conducting) events within bursts at pH 7.4 and pH 6.4 are shown in Figure 4.6 and are based on depolarizations lasting for  up to 180 s (Figure 4.5) to prevent the censoring of  a long-lived, non-conducting state that usually occurred with 1 s depolarizations to 100 mV, particularly at pH 6.4 (e.g.,  Figure 4.1 B). The open duration histogram at pH 7.4 (Figure 4.6 A) was fitted  by the sum of  two exponentials. The slower component of  the frequency  distribution had a mean open time (rs) of  1.5 ± 0.1 ms (n  = 8 patches) and represented the more frequently  observed open event (a s = 0.70 ± 0.04). The faster component of  the distribution represented 0.3 ± 0.04 of  the open events and had a mean duration (Tj) of  0.34 ± 0.02 ms, which is probably an underestimate given the limited frequency  response of  the system. At pH 6.4 the open duration histogram was also biexponential (Figure 4.6 Q. The slower component had a longer mean open time (rs) of  1.4 ± 0.1 ms and represented approximately 0.80 ± 0.04 of  the events. The less frequent  (a f=  0.20 ± 0.04), faster  component had a mean dwell time (rj)  of  0.35 ± 0.05 ms (n  = 7 patches). Neither the mean dwell times nor the proportion of  time spent in these two open states changed significantly  between pH 7.4 and pH 7.4 m*mm mmmimmm^jj » 1 s pH 6.4 M 1 i m M  , II 1 1 s Figure 4.5 Decreasing external pH to 6.4 decreases the mean burst length and increases the apparent interburst duration. A. Representative unitary currents at pH 7.4 in a cell-attached patch. High K+0 bath solution was assumed to depolarize the cell to 0 mV; the pipette solution contained 3.5 mM K+. Contiguous traces of  the first  80 s of  activity at 100 mV are shown. Bursts of  flickering channel behaviour were bracketed by gaps longer than 20 ms (see Materials and Methods). B. Representative contiguous traces from  another cell-attached patch but at pH 6.4. The within-burst behaviour was only slightly changed, the mean burst duration was decreased and the apparent interburst duration was increased. Both traces were digitally filtered  at 1 kHz. 6.4 (p  > 0.05; ANOVA). Panels B and D of  Figure 4.6 plot, at pH 7.4 and 6.4, respectively, the frequency distribution of  the closed events during a long depolarization to 100 mV. At pH 7.4 the closed duration histogram was fitted  to the sum of  four  exponentials but we focus  initially on the three fastest  components that largely reflect  gating behaviour within a burst. A critical time of  20 ms was used as the minimum gap length that signified  the end of  a burst (defined  below). The time constants (and amplitudes) for  the fastest,  intermediate-fast  and slow-fast  components were 0.22 ± 0.1 ms (a f  = 0.63 ± 0.02), 0.42 ± 0.03 ms (a,_ f  = 0.35 ± 0.02), and 2.6 ± 0.2 ms (a s.f  = 0.02 ± 0.003) (n  = 8 patches). The corresponding values at pH 6.4 were 0.17 ± 0.04 ms (a f  =0.81 ± 0.03), 0.45 ± 0.05 ms {a,_ f=  0.18 ±0.03), and 2.4 ± 0.3 ms (a s_f=  0.01 ± 0.002) (n  = 7 patches). The fastest  closed component had a time constant shorter than the dead time of  our system and consequently its mean lifetime  should be taken as a rough estimate only. Although the mean dwell time of  each of  the three fastest  components of  the closed duration histograms did not A Open Durations B Closed Durations Figure 4.6 Extracellular acidification  does not significantly  affect  gating transitions within bursts. Unitary current from  cell-attached patches recorded at pH 7.4 and 6.4 with 2- or 3-minute depolarizing pulses to 100 mV were idealized using a half-amplitude  method. A. The open duration histogram obtained from  the current trace shown in Figure 4.5 A was fitted  to a biexponential distribution to give time constants of  0.5 ms and 2.6 ms, and an area of  0.30 and 0.70, respectively. B. The closed duration histogram obtained at pH 7.4 was fitted  to a four-component exponential distribution, and the fastest  three time constants were 0.2, 0.5, and 4.8 ms, with a respective area of  0.63, 0.36, and 0.02. The slowest component (time constant 0.48 s) is thought to represent one or more inactivated states. It has a relative area less than 1% of  the total closed events. C. The open duration histogram generated from  the current trace shown in Figure 4.5 B at pH 6.4. The fast  and slow time constants are 0.5 and 2.1 ms with an area of  0.2 and 0.8, respectively. D. The closed duration histogram at pH 6.4 was fitted  to a 4-component exponential distribution. The three fastest  components have time constants 0.2, 0.4, and 2.6 ms and an area 0.81, 0.18, and 0.01, respectively. The slowest component (time constant 5.5 s) represents less than 1% of  the total non-conducting events. change significantly  between pH 7.4 and pH 6.4 (p  >  0.05; ANOVA), their relative proportions did change. Compare to that at pH 7.4, the relative proportion of  the fastest  component increased significantly  (p <0.01) at pH 6.4 while the intermediate-fast  component decreased significantly {p  < 0.01) at pH 6.4; however, the slow-fast  component was unchanged (p  > 0.1). These results showed that intraburst gating behaviour was not dramatically affected  by changing pH from  7.4 to 6.4. 4.3.3 pH affects  the burst- and interburst duration A burst of  openings is conventionally defined  as a series of  openings separated by gaps that are less than a critical length (t crjl) (Colquhoun and Sigworth, 1995). A value for  tcn, was determined by numerically solving the equation, exp(-/OT/rs/) = 1- exp(-/cr,/r5) where r t / i s the time constant for  the slow-fast  component during a burst and r5 is the time constant for  the slowest component in the closed duration histogram (Figure 4.6). Using conservative estimates for  rJ./and rs of  3.5 and 500 ms respectively, tcn, was set to 20 ms for  either pH. Despite pooling the data from  patches at pH 7.4 (n = 8) or pH 6.4 in = 7) the frequency  of  events remained low and therefore  it was not possible to obtain a reliable estimate of  the ..number of  interburst non-conducting states. Additionally, at pH 6.4 the P0 was low and, although overlapping channel openings were never observed in the data that were analysed, it was not possible to be certain that there was only one channel in the patch, thus making a meaningful  analysis of  the interburst data problematic. An analysis of  the mean burst duration was more straightforward  because overlapping openings were never observed during bursts at either pH. By dividing the accumulated burst length by the number of  bursts observed, the mean burst duration was determined to be 1.7 s at pH 7.4 and 0.73 s at pH 6.4. The data of  Figure 4.5 also show that the slow, non-conducting state at either pH is non-absorbing. Compared to the pH 7.4 data, the probability of  the slow, non-conducting state was higher at pH 6.4 and it is more stable. 4.4 Discussion Whether Kvl .5 channels are expressed in HEK-293 cells (Kehl et al., 2002) or in ltk~ cells, as in this study, extracellular acidification  has two major effects  on Kvl.5 macroscopic current: 1) the peak current amplitude decreases, and, 2) the inactivation rate of  residual currents increases. Macroscopic current, /, is equal to NPJ  or, in an expanded form,  NP ay(V-E K)  where N is the channel density, P0 is the open probability which has voltage and time dependence, y is the single channel conductance, and EK  is the reversal potential. Analysis of  one-channel patches showed that decreasing pH did not affect  i at pHs between 7.4 and 5.9 and over a range of voltages (Figure 4.1 and 4.2) indicating that y had not changed. This provided direct evidence that the decline of  macroscopic, currents is not due to occlusion of  the open pore or a change of the permeation pathway. A similar conclusion has been reached for  the effect  of  external protons on macroscopic ShakerlR  current (Starkus et al.',  2003). Ensemble data constructed from  idealized traces (Figure 4.3 B) replicated the main features  of  macroscopic currents indicating that the observed changes of  single channel behaviour can account for  most of  the previously reported effects.  One effect  of  low pH, a rightward shift  of  the g-V  relationship (Kehl et al., 2002) that is presumably due to screening and/or binding to surface  charges (Trapani and Korn, 2003), was not apparent at the single channel level because of  the strong depolarization that was typically used. The main effect  of acidification  was to decrease P0: 1) by decreasing channel availability as shown by an increased proportion of  blank or null sweeps; and, 2) by decreasing the average burst duration during active sweeps. Of  these two changes, the decrease of  channel availability was the primary cause of  the decrease of  the peak macroscopic conductance. This was illustrated by the overlap of  the availability-pH curve derived from  single channel analysis and the gmax-pH curves derived from' the analysis of  macroscopic currents (Figure 4.3 C). Diary plots and the outcome of  runs analyses of  the channel behaviour at pH 6.4 (Figure 4.4) showed that null and active sweeps were clustered and consequently that the behaviour likely represented different  sets or modes of gating. The three criteria typically used to define  modal gating were met (Nilius, 1988). First, the two distinct kinetic behaviours were consistently observed in one-channel patches, thus ruling out the possibility of  two populations of  channels. Second, the probability of  a given gating mode could be experimentally manipulated, in this case by changing pH. And, third, the rate for the transition between the two modes was slow, as shown by clustering in diary plots and by the absence of  clear evidence of  a switch from  mode U  to mode A during 1 s depolarizations. Active sweeps at pH 6.4 and 5.9 frequently  ended with a long, censored closed state (Figure 4.1 B) that we assume is due primarily to depolarization-induced inactivation, but we cannot preclude the possibility of  transitions to mode U. Three lines of  evidence lead us to reject a model in which depolarization-induced inactivation accounted for  the null sweeps. These are, first,  that mode U  gating observed at low pH is not dependent on prior channel opening (first  sweeps in Figure 4.1 B and C). The mean burst length, which is a reflection  of  the rate of  depolarization-induced inactivation, is much longer than the dead time of  our system so that a failure  to detect very brief  open events is an unlikely explanation for  the null sweeps. Second, clustering of  null sweeps was observed even with very brief  (20 ms) depolarizations where the probability of  depolarization-induced inactivation was very low. Third, in a previous study of  macroscopic currents the decline of  the peak current was evident on the first  sweep after  a 2 min period in which the potential was held continuously at -80 mV while the pH was changed (Kehl et al., 2002). These observations are inconsistent with a model in which null sweeps resulted from  the occupancy of  a long-lived inactivated state entered during a depolarizing pulse. The lumping together of  two distinct gating behaviours into U  and A modes, and the assignment of  a first  order reaction scheme for  transitions between these two modes is very likely an oversimplification  of  a more complex gating behaviour. The details of  states and the kinetics of  transitions within a gating mode, as well as the exact connectivity between states in the two modes, will be required to develop a full  understanding of  the kinetic processes involved. A more detailed scheme of  mode A would include several closed states, closed-inactivated states, an inactivated state, an open state as well as closed states outside of  the activation pathway (Olcese et al., 1997; Zagotta et al., 1994b; Kurata et al., 2004). Much less is known about the states traversed in mode U.  However, save for  a rightward shift  of  the Q-V  curve, gating currents are unchanged at low pH indicating that voltage sensor movement is more-or-less intact in channels gating in mode U. Analysis of  active sweeps at pH 7.4 or pH 6.4 suggests that in mode A the opening and closing transitions within a burst were largely pH-insensitive (Figure 4.6). At either pH, Kvl .5 channel activity is characterized by rapid flickering  that, especially at 100 mV, most likely reflects  transitions between short-lived open and closed (i.e.,  non-inactivated) states outside of the normal activation pathway (Schoppa and Sigworth, 1998b; Zagotta et al., 1994a). There was evidence for  two states in the open duration histogram but the faster  of  these two was near the dead time of  the system and is therefore  equivocal. Aside from  this second open state, channel gating within a burst, including the existence of  the three fastest  components in the closed duration histogram, is very much as described for  ShakerlR  channels (Schoppa and Sigworth, 1998b). The slowest component of  the closed duration histogram represented a state that terminated a burst and was attributed to inactivation. However, because of  the relatively small number of  these long non-conducting events, we cannot be certain of  either the number of inactivated states or their mean dwell times. To address the simpler question of  how a change of pH affected  the equilibrium between bursting behaviour and gaps between bursts, channel activity was studied during minutes-long depolarizations to 100 mV (Figure 4.5). Decreasing pH from  pH 7.4 to 6.4 decreased the burst duration and appeared to increase the interburst interval which can account for  the faster  inactivation rate of  macroscopic and ensemble currents (Figure 4.3, A and B) as well as the slower rate of  recovery from  inactivation at low pH (Kwan et al., 2004). Extracellular protons increase the depolarization-induced inactivation rate in ShakerlR (Starkus et al., 2003), N-terminal deleted Kvl.4 (Claydon et al., 2002), rat Kvl.5 (Steidl and Yool, 1999) and human Kvl.5 (Kehl et al., 2004). This enhancement of  the inactivation rate is attenuated by the T449V mutation in ShakerlR  and by the homologous mutation, R487V, in Kvl .5 and as such is consistent with a connection to outer pore inactivation. However, despite the fact  that depolarization-induced inactivation both in ShakerlR  and Kvl.5 is accelerated by decreasing pH, there are several important, unexplained differences  in the response of  these two structurally related channels to extracellular acidification.  Protonation of  H463 appears to be the primary event that triggers both a change of  availability and the acceleration of  depolarization-induced inactivation in Kvl.5 since both effects  are substantially reduced in Kvl.5 H463Q (Kehl et al, 2002). In contrast, in ShakerlR  the acceleration of  depolarization-induced inactivation has been provisionally attributed to protonation of  the aspartate residue in the GYGD sequence of  the selectivity filter.  The most striking disparity is that acidification  decreases channel availability (increases mode Ugating)  in Kvl.5 but not in ShakerlR  or Kvl .4. Interestingly, however, mutations of  the threonine residue at position 449 in Shaker  (Lopez-Barneo et al., 1993) or its positional equivalent in Kvl.4 (Pardo et al, 1992) can produce currents that collapse in K+-free external medium, an effect  that has been attributed to a change of  availability due to closed-state inactivation (Pardo et al., 1992). We have not yet assessed the consequences of  mutations of R487 on channel availability at pH 7.4 in Kvl .5 but have found  that mutations of  residues at or near the putative proton-binding site (H463) in the pore turret can produce currents at pH 7.4 that show fast  inactivation and have a K+0-sensitivity similar to that observed both for  ShakerlR mutants and for  wt Kvl.5 channels at low pH0 (Eduljee et al., 2004; Kehl et al., 2002). Although a faster  rate of  depolarization-induced inactivation is associated with a K+0-sensitive change of availability in ShakerlR  T449 mutants, this is not a consistent correlation. For example, with wt ShakerlR  decreasing the external pH can dramatically speed inactivation but does not appear to affect  channel availability (Starkus et al., 2003), and, conversely, the inhibition of  Kvl.5 current by divalent cations such as Ni2+, Co2+ and Zn2+ is correlated with a substantial decrease of  channel availability that is associated with relatively small changes of  depolarization-induced inactivation rate (Kwan et al., 2004). This indicates that "closed-state inactivation" and "open-state inactivation" do not always show parallel changes. An important question that remains to be answered is whether the decreased availability or mode U  gating we observe at low pH in Kvl .5 is due to a constriction of  the outer pore mouth, as has been proposed for  P/C-type inactivation. Typically, P/C-type inactivation is assumed to be strongly coupled to channel activation such that it occurs from  the open state or perhaps from  one or more closed states accessed outside of  the activation pathway. If,  as suggested by the effects of  K+0 and the R487V mutation on the inhibition of  Kvl.5 current at low pH, the decreased channel availability does indeed reflect  outer pore inactivation, this would mean that this process becomes uncoupled from  activation. Evidence obtained from  the non-conducting mutant ShakerlR  W434F for  "permanent" or resting P-type inactivation provides some support for  this idea (Yang et al., 1997). Acknowledgement This work was supported by a grant from  the Canadian Institutes of  Health Research to S.J.K. and D.F. D.C.H.K. received trainee awards from  the Michael Smith Foundation for  Health Research and the Natural Sciences and Engineering Research Council of  Canada. 4.5 References Claydon, T. W., M. R. Boyett, A. 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Depolarization of  Kvl .5 channels in acidic, K+-free external medium evokes a slowly rising phase of  current. Biophys. J. 86:534a. (Abstr.) Kurata, H. T., Z. Wang, and D. Fedida. 2004. NH2-terminal inactivation peptide binding to C-type-inactivated Kv channels. J. Gen. Physiol. 123:505-520. Kwan, D. C., C. Eduljee, L. Lee, S. Zhang, D. Fedida, and S. J. Kehl. 2004. The external K+ concentration and mutations in the outer pore mouth affect  the inhibition of  Kvl.5 current by Ni2+. Biophys. J. 86:2238-2250. Loboda, A., A. Melishchuk, and C. Armstrong. 2001. Dilated and defunct  K channels in the absence of  K+. Biophys. J. 80:2704-2714. Lopez-Barneo, J., T. Hoshi, S. H. Heinemann, and R. W. Aldrich. 1993. Effects  of  external cations and mutations in the pore region on C-type inactivation of  Shaker  potassium channels. " Receptors. Channels 1:61-71. Nilius, B. 1988. Modal gating behavior of  cardiac sodium channels in cell-free  membrane patches. Biophys. J. 53:857-862. Olcese, R., R. Latorre, L. Toro, F. Bezanilla, and E. Stefani.  1997. Correlation between charge movement and ionic current during slow inactivation in Shaker  K+ channels. J. Gen. Physiol. 110:579-589. Pardo, L. A., S. H. Heinemann, H. Terlau, U. Ludewig, C. Lorra, O. Pongs, and W. Stuhmer. 1992. Extracellular K+ specifically  modulates a rat brain K+ channel. Proc. Natl. Acad. Sci. U. S. A. 89:2466-2470. Plummer, M. R. and P. Hess. 1991. Reversible uncoupling of  inactivation in N-type calcium channels. Nature 351:657-659. _ Saftenku,  E., A. J. Williams, and R. Sitsapesan. 2001. Markovian models of  low and high activity levels of  cardiac ryanodine receptors. Biophys. J. 80:2727-2741., Schoppa, N. E. and F. J. Sigworth. 1998a. Activation of  Shaker  potassium channels. I. Characterization of  voltage-dependent transitions. J. Gen. Physiol. 111:271-294. Schoppa, N. E. and F. J. Sigworth. 1998b. Activation of  Shaker  potassium channels. III. An activation gating model for  wild-type and V2 mutant channels. J. Gen. Physiol. 111:313-342. Starkus, J. G., Z. Varga, R. Schonherr, and S. H. Heinemann. 2003. Mechanisms of  the inhibition of  Shaker  potassium channels by protons. Pflugers  Arch. 447:44-54. Steidl, J. V. and A. J. Yool. 1999. Differential  sensitivity of  voltage-gated potassium channels Kvl.5 and Kvl.2 to acidic pH and molecular identification  of  pH sensor. Mol. Pharmacol. 55:812-820. Trapani, J. G. and S. J. Korn.~2003. Effect  of  external pH on activation of  the Kvl .5 potassium channel. Biophys. J. 84:195-204. Wang, Z., X. Zhang, and D. Fedida. 2000. Regulation of  transient Na+ conductance by intra- and extracellular K+ in the human delayed rectifier  K+ channel Kvl.5. J. Physiol. 523:575-591. Yang, Y., Y. Yan, and F. J. Sigworth. 1997. How does the W434F mutation block current in Shaker  potassium channels? J. Gen. Physiol. 109:779-789. Zagotta, W. N., T. Hoshi, and R. W. Aldrich. 1994a. Shaker  potassium channel gating. Ill: Evaluation of  kinetic models for  activation. J. Gen. Physiol. 103:321-362. Zagotta, W. N., T. Hoshi, J. Dittman, and R. W. Aldrich. 1994b. Shaker  potassium channel gating. II: Transitions in the activation pathway. J. Gen. Physiol. 103:279-319. Zar, J. H. 1984. Biostatistical Analysis. 2nd Ed. Prentice-Hall. Upper Saddle River, New Jersey. Zhang, S., H. T. Kurata, S. J. Kehl, and D. Fedida. 2003. Rapid induction of  P/C-type inactivation is the mechanism for  acid-induced K+ current inhibition. J. Gen. Physiol. 121:215-225. 5. The microscopic changes in Kvl.5 slow inactivation gating caused by external K+ and H+ 5.1 Introduction Increasing external K+ antagonizes the inhibition of  macroscopic current caused by H+ in Kvl .5 (Kehl et al., 2002). While the mechanism by which external H+ inhibits Kvl.5 current was investigated at the single channel level in Chapter 4 (Kwan et al., 2006), the effects  of external K+ on this inhibition were not studied previously. With a fixed  [K"]0 (3.5 mM), both the > single channel conductance (between 0 and 100 mV) and the mean dwell times within bursts were unaltered by external H+. Instead, H+0 increased the number of  sweeps showing no channel activity (null sweeps). This behaviour was modelled as a shift  in the mode of  gating, in which mode A gating was proposed to result in normal gating behaviour (i.e.,  with open channel activity), and mode Ugating  resulted in channels that failed  to open (i.e.,  null sweeps). Based on the idea that external H+ inhibits Kvl.5 current by promoting mode {/gating, external K+ is hypothesized to antagonize the effect  of  H+ by inhibiting the promotion to mode {/gating. In addition to the aforementioned  effect,  removing all external K+, either at low extracellular pH or in the presence of  external Ni2+, Kvl.5 current evoked by long depolarizing pulses (> 1 s) showed an unexpected "slow rising phase" instead of  a time-dependent decay (i.e.,  slow inactivation) (Kwan et al., 2004). The mechanism underlying this unusual behaviour remains uncertain, but this phenomenon appears to result from  channels coming out of  a non-conducting state that may be related to slow inactivation. Therefore,  a detailed understanding of  slow inactivation in Kvl.5 is needed to answer these questions. To investigate the molecular mechanisms of  external K+ modulation of  H+-induced effects and the slow rising phase, two different  experimental approaches were used. In Part I, the mechanism by which external K+ modulates Kvl.5 current at a fixed  pH was examined. Unitary currents were recorded at pH 6.4 with 0, 1, 3.5, and 20 mM external K+ in the cell-attached configuration.  By using long depolarizing pulses (> 6 s), some sweeps were seen to have long first  latencies to opening, which was thought to represent channels switching from  mode U  back to mode A. With increased external [K+], the proportion of  sweeps starting in mode A was increased, which was consistent with an inhibition of  mode U  gating. In Part II, the slow rising phase of  current was studied using an external solution in which the external [K+] was maintained at 0 mM while varying external pH from  7.4 to 5.9. When the sweeps showing long latency were summed together, a prominent slow rising phase was seen. The result suggested the slow rising phase resulted from  a recovery from  mode U gating. In addition, the mean long first latency correlated with the mean dwell time of  a depolarization-induced inactivated state. Together, these results suggest that a slow inactivation process underlies mode U  gating. 5.2 Materials and Methods 5.2.1 Cell preparation A stable mouse cell line ltk~  expressing Kvl.5 was made from  the cDNA of  Kvl.5 subcloned into the gentamicin-resistant gene-containing pcDNA3 vector as described previously (Wang e/ al., 2000). Cells were maintained in minimum essential medium (MEM) supplemented with 10% fetal  bovine serum, 1% penicillin-streptomycin, and 1% gentamicin and incubated at 37°C ( in an atmosphere of  5% CO, in air. Cells were dissociated enzymatically and were plated onto coverslips for  experimental use within 1-3 days. All tissue culture supplies were obtained from Invitrogen (Burlington, Ontario, Canada). 5.2.2 Electrophysiology Whole-cell (macroscopic) and unitary (microscopic) currents were recorded using standard voltage clamp techniques in the whole-cell and cell-attached configurations,  respectively. For whole-cell recording, cells were bathed in nominally K+-free  bath (external) solution containing (in millimolar) 143.5 NaCl, 2 CaCl2, 1 MgCl2, and 5 glucose adjusted to pH 7.4 (10 HEPES/NaOH), 6.9 (10 HEPES/NaOH), 6.4 (10 MES/NaOH), or 5.9 (10 MES/NaOH), or K+-containing external solutions made by substituting equimolar amounts of  KC1 for  NaCl at pH 6.4. The patch pipettes contained Ca2+- and Mg2+-free  (internal) solution with 130 KC1, 10 EGTA, 10 HEPES adjusted to pH 7.4 with KOH. For cell-attached single channel recording, the bath solution contained 140 KC1, 3.5 NaCl, 2 CaCl2, 1 MgCl2, 5 glucose, and 10 HEPES buffered  to pH 7.4 and was assumed to depolarize the cell to 0 mV. The pipettes for  cell-attached recordings contained either nominally K+-free  (extracellular) solution with 143.5 NaCl, 2 CaCl2, 1 MgCl2, and 5 glucose adjusted to pH 7.4 (10 HEPES/NaOH), 6.9 (10 HEPES/NaOH), 6.4 (10 MES/NaOH), or 5.9 (10 MES/NaOH), or K+-containing solution made by substituting an equimolar amount of  KC1 for  NaCl at pH 6.4. In subsequent text, references  to K+ and pH refers to the extracellular [K+] and extracellular pH. All chemicals were purchased from  Sigma-Aldrich (Mississauga, Ontario, Canada). Voltage commands and current measurements were made at room temperature (20-25 °C) with an EPC-7 patch clamp amplifier  connected to an ITC-18 digital interface  (Instrutech, Port Washington, New York) and controlled by Pulse software  (HEKA Electronik, Lambrecht, Germany). Patch electrodes pulled from  thin-walled borosilicate glass (World Precision Instruments, Sarasota, Florida) had a resistance of  1-3 MQ for  whole-cell experiments and 8-15 MQ for  cell-attached single channel recordings when measured in recording solutions. Cells were held at -80 mV, and a liquid junction potential of-4  mV (or 4 mV) was compensated in all voltage measurements in the whole-cell (or cell-attached) recording modes. Current signals were low-pass filtered  at 3 kHz (-3dB, 8-pole Bessel, NPI Electronics, Tamm, Germany) and digitized at either 5 or 10 kHz. For analysis, unitary currents were further  filtered  digitally at 1 kHz which gave an effective  filtering  frequency  of  950 Hz. 5.2.3 Data analysis Whole-cell macroscopic currents were analysed with Igor Pro 5.0.4 (WaveMetrics, Lake Oswego, Oregon).- The (fast)  activation time course was estimated by fitting  currents within the first  20 ms between 50% and 100% of  the peak current to a single exponential function. Inactivation kinetics were measured by fitting  the current decay with a double exponential function.  The time course of  the slow rising phase was measured by-fitting  the current from 200 ms after  the start of  the pulse to the end of  the pulse with double exponential functions  to avoid an initial "hook" of  current that was observed in some recordings {e.g.,  Figure 5.7 A, pH 6.4), Single channel currents recorded by voltage steps at +100 mV from  a holding potential of -80 mV were first  corrected for  capacitive currents using blank (null) sweeps recorded from  the same patch and then idealized in TAC (Bruxton, Seattle, Washington) using a half-amplitude threshold criterion with a rise time of  0.3 ms for  event detection (Kwan et al., 2006). Only patches with a single (major) level of  event were used for  all analyses except for  the first  latency histograms, in which all unequivocal latencies to first  opening in multi-channel patches were also included (see below). Conductance substates were observed, but this phenomenon was not analysed in detail. Single channel current amplitude was determined from  the difference  in amplitude between the two major peaks in the all-points histogram that was fitted  to a Gaussian distribution function  with three components (see Figure 4.7). Given a combined filtering frequency  of  950 Hz, the dead time of  the system was estimated to be 0.3 ms (Hoshi et al., 1994), and events.shorter than the dead time were censored. Based on the analysis in Chapter 4, a , 20-ms critical time (t cril) was implemented for  determining the termination of  bursts (Kwan et al., 2006). No correction was made for  missed events in the analysis of  duration histograms. Duration (dwell time) histograms were fitted  to exponential distribution functions  with multiple components in TACFit (Bruxton, Seattle, Washington) and in Igor Pro. The number of components was determined by a maximum likelihood technique, in which the least number of components with a significant  improvement was used (Saftenku  et al., 2001). A significant improvement was achieved if,  given the log likelihood values of  the two models (LL,  and LL2), a value of  (2\LL, - LL2\ ) was greater than the £ value with a degree of  freedom  (v) equal to the difference  in free  parameters between the two models. Averaged results were expressed as mean ± SEM (standard error of  the mean) unless otherwise stated, and the time constants for  the histograms are given as mean ± SD (standard deviation). Statistical tests (Student's /-test, ANOVA) were performed  with Jmp In Software  (SAS Institute, Cary, North Carolina). The correlation coefficient  (r)  and its standard error (s r) were calculated as described previously (Zar, 1984). A probability of  less than 0.05 was considered significant. 5.3 Results Part  I  Mechanism  for  the relief  of  It-induced  inhibition  by K +a 5.3.1 External K+ antagonizes the H+-induced Kvl.5 current inhibition in ltk~  cells Based on the analysis of  tail current behaviour of  Kvl.5 in HEK-293 cells, it was shown that external K" antagonizes the macroscopic current inhibition induced by external IT (Chapter 2). Figure 5.1 A shows superimposed representative current traces evoked by 6 s depolarizing pulses at +60 mV with 0, 1, 3.5, and 20 mM external K+ at pH 6.4 from  a mouse Itk  cell expressing Kvl .5. As noted in Chapter 3, with 0 mM K+, Kvl .5 showed a slow rising phase at pH 6.4 instead of  a time-dependent decay. This slow rising phase was not evident with 1 mM or higher concentrations of  K+. The basis for  this slow rising phase is considered in more detail in the second part of  this chapter. By increasing external [K+], current amplitude is increased, and this result can be seen more clearly by expanding the first  100 ms of  the current traces (Figure 5.1 B). During the first  100 ms, Kvl.5 reached a peak within the first  20 ms (Ip20ms). Figure 5.1 C shows the relative I p 2 0 m s with different  [K+] normalized with respect to that with 0 mM K+. With 1, 3.5, and 20 mM K+, the peak current had an average fold-increase  of  3.68 ± 0.14, 4.14 ± 0.07, and 5.64 ±0.61, respectively (p  < 0.01), compared to that with 0 mM K+. This enhancement of  current caused by increasing [K+]0 was similar to that shown in Figure 2.4. These values did not take into account the changes in driving force  or single channel conductance and thus may underestimate the relief of  the pH-dependent inhibition by external K+. Figure 5!1 D shows the current traces normalized to their respective I p 2 0 m s , an approach that emphasizes the slow rising phase at 0 mM K+. The current traces with 0 mM K+ (at pH 6.4) were fitted  to a double exponential function,  which gave a mean time constant ( t s l o w r ise) of  1.44 ± 0.12 s (n  = 11; Figure 5.1 E) for  the slow rising phase and a slower component ( r = 7.2 ± 0.6 s) for  the decay phase. In contrast, the current traces with 1, 3.5, and 20 mM K+ show a multi-component time-dependent decay (i.e.,  slow inactivation) which can be fit  to a double exponential function.  The values for  rfas l and rs/0M, were 0.35 ± 0.07 s and 1.87 ± 0.33 s with pH 6.4 D 5 nA B E 2.5-1 2 . 0 -1 . 5 -1.0 0.5 0 . 0 - 1 50 ms ^ 4 1 0-6 N 15 0.4 ° 0.2 OK ' 1 K ' 3.5 K 20 K 1.0—1 0 . 8 -o 0 . 6 -% o f  0.4 0.2-1 0.0 • Steady state • Fast comp. • Slow comp. :Uim 1 K m 3.5 K 20 K H E 1.5 OK ' 1 K ' 3.5 K ' 20 K ' OK ' 1 K 1 3.5 K 20 K Figure 5.1 Effects  of  external K+ on the macroscopic current through Kvl.5 expressed in ltk~  cells at pH 6.4. Kvl .5 currents were recorded using 6 s depolarizing pulses to +60 mV with 0, 1, 3.5, and 20 mM external K+ at pH 6.4 to study the changes in current amplitude and gating kinetics. A. Representative Kvl.5 currents from  the same cell are superimposed to show the relief  of  current inhibition at pH 6.4 by external K+. Note that the current recorded at pH 6.4 with 0 mM K+ has a slow rising phase as opposed to the prominent current decay observed with higher [K+]0. B. The traces in A are shown on an expanded time scale. The difference  in current amplitude and the activation kinetics can be seen more clearly, but the slow rising phase with 0 mM K+ was not evident on this time scale. C. The relief  of  current inhibition was quantified by.calculating the relative peak current amplitude during the first  20 ms (Ip 2 0 m s) with respect to that during the first  20 ms at 0 mM K+. The relative current was 3.68, 4.14, and 5.64 with 1, 3.5, and 20 mM K+, respectively D. Current traces were normalized to their respective I p20ms to allow a comparison of  the current kinetics. This approach emphasizes the slow rising phase observed with 0 mM K+, which has a mean time constant of  1.44 ± 0.12 s (n  = 11). Inactivation kinetics were affected  only slightly by changing external K+ between 1 and 20 mM. E. Current traces were fitted  to double exponential functions,  and the fast  (r fas t; black bars) and slow (r,/oM,; grey bars) time constants were 0.35 ± 0.07 s and 1.87 ± 0.33 s with 1 mM K+ [n = 5), 0.20 ± 0.03 s and 1.41 ± 0.27 s with 3.5 mM K+ (n = 5), and 0.35 ± 0.03 s and 2.08 ± 0.45 s with 20 mM K+ (n  = 4), respectively. Only T f m, of  3.5 mM K+ was significantly  different  from  that with 1 and 20 mM K+. Included in this panel is the mean time constant for  the slow rising phase (T slownse; hatched  bar) for  comparison. F.  The proportions of the steady-state (empty  bars), fast  (black  bars), and slow (grey  bars) components were unaltered by changing external K+. Their respective values were 0.30 ± 0.02, 0.19 ± 0.02, and 0.51 ± 0.03 with 1 mM K+ (n  = 5), 0.20 ± 0.01, 0.15 ± 0.01, and 0.65 ± 0.01 with 3.5 mM K+ (.n = 5), and 0.23 ± 0.02, 0.24 ± 0.05, and 0.53 ± 0.04 with 20 mM K+ (n  = 4). G. The current at the end of  the 6s pulse (I6s) was normalized to its corresponding I p 2 0 m s . With 0 mM K+, I6s was larger than 1 (1.22  ± 0.05; n = 12) due to the slow rising phase. The values for  the relative 4 with 1, 3.5, and 20 mM K+ were 0.32 ± 0.02 (n  = 5), 0.21 ± 0.01 (n  = 5), and 0.26 ± 0.02 (n = 4). H.  External K+ did not change the activation kinetics significantly.  The activation kinetics were estimated by fitting  current between 50% and 100%) of  l p 2 0 m s within the first  20 ms of  the 6 s pulses to a single exponential function  to obtain the activation time constant ( racl), which was 1.5 ± 0.1 ms (n = 12), 1.8 ± 0.2 ms (n  = 5), 1.9 ± 0.3 ms (n  = 5), and 1.8 ± 0.1 ms (n  = 4) for  the currents with 0, 1, 3.5, and 20 mM K+, respectively (not significant; ANOVA; p = 0.28). The error bars in each panel represent the standard error of  the mean (SEM). 1 mM K+ (n  = 5), 0.20 ± 0.03 s and 1.41 ± 0.27 s with 3.5 mM K+ (n  = 5), and 0.34 ± 0.03 s and 2.08 ± 0.45 s with 20 mM K+ (n  = 4), respectively (Figure 5.^ 1 E). Notice that ts,owrjse is very similar to vslow with 3.5 mM K+. The average rf m, with 3.5 mM K+ was significantly  different from  that with 1 and 20 mM K+, while the values of  rfa s, with 1 and 20 mM K+ were not significantly  different.  The values of  r s h w with 1, 3.5, and 20 mM K+ were also not significantly different  from  each other. The proportion of  each of  the components did not show a clear trend except that the proportion of  the slow component was dominant, and the proportions of  all three components with 1 and 20 mM IC were not significantly  different  (Figure 5.1 F\ see figure legend for  values). Together, these results show that slow inactivation is somewhat enhanced with 3.5 mM K+0, or that a progression out of  the slow rising phase contributed to the apparently slower inactivation with 1 mM K+0. Otherwise, external K+ has a very limited effect  on slow inactivation (Chen et al., 1997). Figure 5.1 G shows the current amplitudes at the end of  6 s depolarizing pulses to +60 mV at different  [K+ Jo with respect to their corresponding Ip 2om s . With 0 mM K+, the current was potentiated to 1.22 ± 0.05, reflecting  the contribution of  the slow rising phase. With 1, 3.5, and 20 mM K+, the current amplitude was reduced to 0.32 ± 0.2, 0.21 ± 0.01, and 0.26 ± 0.02, respectively. Figure 5.1 H  shows the activation time constants (rflC /) with different  [K+], The activation time constant, as measured by fitting  the last 50% of  the rising phase within the first 20 ms of  the current traces, was 1.5 + 0.1, 1.8 + 0.2, 1.9 + 0,3, and 1.8 + 0.1 ms with 0, 1,3.5, and 20 mM K+, respectively (not significantly  different;  ANOVA; p = 0.28). Together, these results are consistent with the findings  presented in Chapters 2 and 3 and suggest that the relief by external K+ of  the H+-induced macroscopic Kvl.5 current inhibition observed in Itk  cells and HEK-293 cells is similar. Since the current inhibition induced by H+ is due to a promotion of mode U  gating, external K+ may relieve this inhibition by antagonizing this promotion to mode U (i.e.,  by promoting mode A gating). 5.3.2 External K+ increases the mean PQ but not the unitary current amplitude The relief  of  H+-induced macroscopic current inhibition by K+ may result from  an increase in open probability (P 0) or from  an increase in the single current amplitude (I  = N-P 0-i). To determine the mechanistic basis for  this relief  of  inhibition, unitary currents were recorded in the cell-attached configuration  from  mouse Itk  cells expressing Kvl.5 with depolarizing pulses lasting 6, 31, or 61 seconds. A 6 s pulse was used so that a comparison with the macroscopic current could be made. In contrast, the 31 s and 61 s pulses were usedto increase the number of reopening events in each sweep and to observe the transition out of  mode U.  Figure 5.2 A-D / shows representative unitary current traces evoked by 31 s depolarizing pulses to +100 mV (pulsed every 60 s) from  different  cell-attached patches with 0, 1, 3.5, and 20 mM K+ at pH 6.4. The traces show that increasing K+ causes an apparent increase in P0 but no change of  the unitary current amplitude (/). To quantify  P0, unitary current traces were idealized using a half-amplitude  technique, and the mean PQ within the first  20 ms (hatched  bars), 6 s pulses (black  bars), and 31 s pulses (grey bars) with 0, 1,3.5, and 20 mM K+ was calculated and shown in Figure 5.2 E. The mean P0 within the first  20 ms gave an estimate of  the proportion of  channels starting in mode U. However, the capacitive currents of  some of  the sweeps had a complex waveform  and thus could not be compensated completely. Consequently, the onset of  current could not be determined unequivocally, and the PQ within the first  2 ms was a rough estimate. Given this uncertainty, an additional 10% error was added to the mean P0 within the first  20 ms. Using this criterion, increasing external K+ significantly  increased P0 within the first  20 ms from  0 to 20 mM K+ (but not between the P0 with 1 and 3.5 mM K+). This result is qualitatively similar to that shown in Figure 5.1 B. Interestingly, the P0 during the first  20 ms was not significantly  different  from  that with 6 s pulses with 0 mM K+. This result reflects  the relative I6s for  the slow rising phase as shown in Figure 5.1 G. For the 6 s pulses (applied every 30 s), the mean P0 was significantly  different  for  each pair except for  that between 0 and 3.5 mM K+. The P0 of  6 s pulses with 1 mM K+ was significantly higher than with 3.5 mM K+, which was consistent with a slower slow inactivation with 1 mM 100 mVr -80 mV-l pH 6.4 B 0 K 5 s 2pA|( 1 K+ 200 ms J U 5S : PAITHMW, , 3.5 K+ 200 ms 5 s 2pA| D AlimiflJlHihfciUMi  i illM^dffc  lilhiitilii^ ili U UiBtfMfcrfdMtaL J f f l B 20 K+ 200 ms I I 5 s 200 ms 0.6-, 0.5 • n 0.4 -_Q £ Q_ c 0.3 • 01 a. O £ 0 2 ' a) s 0.1 • o.o • za First 20ms • 6s pulses • 31 s pulses 1 Q I a 1 Q I. i OK 1 K 3.5 K 20 K OK 1 K 3.5 K 20 K Figure 5.2 Unitary current traces and the mean open probability with 0,1,3.5, and 20 mM external K+ at pH 6.4. Unitary current traces with 0, 1, 3.5, and 20 mM K+ were recorded from different  cell-attached patches using 6 s, 31 s, or 61 s depolarizing pulses to 100 mV. Representative traces (short latency only) lasting 31 s are shown in the upper panels in A-D. Selected 1 s segments of  the current traces (horizontal  bars) are expanded in the lower panel to better show the intraburst behaviour at different  [K+]0. E. The mean open probability (P 0) during the first  20 ms of  depolarizing pulses (hatched  bars), the entire 6 s pulses (black  bars), and the entire 31 s pulses (grey  bars) was 0.10 ± 0.01, 0.13 ± 0.01, and 0.06 ± 0.01 with 0 mM K+, 0.34 ± 0.01, 0.16 ± 0.01, and 0.06 ± 0.01 with 1 mM K+, 0.29 ± 0.01, 0.11 ± 0.01, and 0.07 ± 0.01 with 3.5 mM K+, and 0.55 ± 0.01, 0.29 ± 0.02, and 0.17 ± 0.02 with 20 mM K+. These data show that each increment of  [K+]0 significantly  increased the mean open probability. F.  Single channel current amplitude is unaltered by external K+. Mean single channel current amplitude was determined by fitting  the all-points histograms derived from  individual unitary current sweeps to a Gaussian distribution with 3 exponential components and calculating the difference  between the two major components. The single current amplitudes were 1.5 ± 0.1 pA with 0 mM K+ and 1.6 ±0.1 pA with 1, 3.5, and 20 mM K+. Error bars represent the standard error of  the mean (SEM). K+ as shown in Figure 5.1 A. On the other hand, the mean P0 during 31 s pulses with 0, 1, and 3.5 mM K+ was not significantly  different  from  each other; only that with 20 mM K+ was significantly  higher than the rest. Together, these results show that increasing external K+ changes the mean P( } mainly within the first  6 s, whereas the effect  of  K+ on P0 is diminished with longer pulses. The unitary current amplitude at +100 mV with 0, 1, 3.5, and 20 mM K+ at pH 6.4 was determined by plotting the all-points amplitude histograms (see Methods; data not shown). It was found  that the mean current amplitude was not significantly  different  with varying [K+] between 0 and 20 mM (Figure 5.2 F).  It is unclear why the single current amplitude did not change with the change of  driving force.  Nonetheless, the result shows the relief  of  current inhibition by K+ is due to an increase in P0 rather than an increase in the single channel current amplitude. 5.3.3 External K+ does not change the intraburst behaviour of  Kvl.5 dramatically The increase in mean P0 could result from  changes in the intraburst behaviour (i.e.,  an increase in the dwell time of  the open state(s) and/or a reduction in the dwell time of  the closed states), in the bursting behaviour (increased burst length and/or reduction in gap (interburst) duration), and/or an increase of  the probability of  mode A gating. The lower trace from  each panel in Figure 5.2 A-D shows a selected section of  the upper trace with channel activities on an expanded time scale as indicated by the horizontal bars. The intraburst behaviour at each [K+] tested appears to be very similar, suggesting that a change in the intraburst behaviour is not responsible for  the K+-induced current relief.  Figure 5.3 A shows representative open duration histograms for  0, 1, 3.5, and 20 mM K+ from  4 different  cell-attached patches. Each of  the histograms was fitted  to a double exponential distribution function  with time constants r 0 / and t0s for  the fast  and slow components, respectively. Figure 5.3 B shows the averaged r a / and r 0 v at each of  the [K+]0 tested. The values of  r 0 /were very close to the dead time of  the recording system and thus cannot be compared with any degree of  confidence.  On the other hand, r 0 1 did not show a clear trend with external [K+], except that r 0 s with 0 mM K+ was slightly larger than the others (significant;  ANOVA; p < 0.05). As shown in Figure 5.3 C, both the relative proportion of  r a / and r 0 s did not change significantly  with [K+], These results show the open states are not dramatically affected  by external K+. Similarly, the closed states within bursts were not affected  by external K+. Figure 5.3 D shows representative closed duration histograms for  0, 1, 3.5, and 20 mM K+ at pH 6.4. Each histogram was fitted  to a 5-component exponential distribution function.  Among the 5 time constants, r c „ r C 2 , and r C J were smaller than the critical time (tcn, = 20 ms; red line) and thus taken to represent the "mean durations of  non-conducting states within a burst. On the other hand, r C 4 and r 0 were larger than tcr i, thus representing the mean durations of  inactivated states. On this basis, rC 4 and t C 5 were treated separately from  the other three components and are . considered in greater detail below. The mean dwell times and the proportion for  t c i , t C2, and rC3 with 0, 1, 3.5, and 20 mM K+ were summarized in Figure 5.3 E and F,  respectively. Again, the values of  r C J are too small to be compared unequivocally. Both the mean closed times and the proportion for  each of  these components did not change significantly  with external [K+], Together, the data of  Figure 5.3 show that both the open times and closed times within bursts were at most modestly affected,  if  at all, by changing external [K+], 0 mM K 1 mM K D 0 mM K 1 mM K 2.5 n 2.0 • 1.5-1 . 0 -0.5 0.0 = 0.3 ms i c 2 = 1.1ms 3000 -* = 5.7 ms c „ „ c'3 2 2000-t c „ = 46ms « 4000 § 1000-n l 0.1ms 1ms 10 ms 0.1 Duration 3.5 mM K+ 0.1 ms 1 ms 10 ms 0.1 s 1s Duration 0.1 ms 1 ms 10 ms 0.1 s 1s Duration 20 mM K 3.5 mM K 20 mM K ~l I I 0.1 ms 1 ms 10 ms 0.1 s Duration T 0.1 ms 1 ms 10 ms 0.1 s 1s Duration rh r±| 1+1 1 K 3.5 K 20 K c 1 . 0 0 . 8 -| 0 .6-o Q. e 0.4 CL 0 . 2 -0.0 1 1 1 K 3.5 K 20 K 6 • 5 • 4 -1.3-" 2 -1 -0- •Q =Q j j m I K 3.5 K 20 K Figure 5.3 External K+ does not alter gating within bursts at pH 6.4. A. Representative open duration histograms from  four  different  cell-attached patches with 0, 1, 3.5, and 20 mM K+ are shown. Each open duration histogram was fitted  to a distribution with the sum of  2 exponential components ( r 0 /and t 0 v). B. Both xof  (black  bars) and x0s (grey  bars) with 0, 1, 3.5, and 20 mM K+ are shown, and the respective values were 0.4 ±0.1 ms and 1.9 ± 0.1 ms with 0 mM K+ (n  = 16 patches), 0.3 ± 0.1 ms and 1.4 ± 0.1 ms with 1 mM K+ (n  = 7 patches), 0.2 ± 0.1 ms and 1.4 ± 0.1 ms with 3.5 K+ (n  = 14 patches), and 0.2 ± 0.1 ms and 1.5 ± 0.1 ms with 20 mM K+ (n  = 7 patches). Only the value of  t 0 s with 0 mM K+ was significantly  different  from  that with 1 and 3.5 mM K+. C. Changing [K+]0 did not affect  the proportion of  the fast  (black  bars) and slow (grey  bars) components, which were 0.19 ± 0.02 and 0.81 ± 0.03 with 0 K + ( « = 16), 0.24 ± 0.04 and 0.76 ± 0.04 with 1 mM K+ (n  = 7), 0.29 ± 0.03 and 0.71 ± 0.03 with 3.5 mM K+ (n  = 14), and 0.26 ± 0.03 and 0.74 ± 0.03 with 20 mM K+ (n  = 7). D. Representative closed duration histograms were generated from  idealized unitary current traces recorded with 0, 1, 3.5, and 20 mM K+ from  4 different  cell-attached patches. Each closed duration histogram was fitted  to a 5-component exponential distribution function.  The two slowest components had mean durations that were longer than the critical time (20 ms; red line) and thus are taken to represent gating activities occurring outside of  bursts. E. Increasing [K+]0 did not change significantly  the three fastest  components (r c /, black  bars; t C 2, grey bars; and r C 3 , hatched  bars) of  the closed duration histograms. The values of  r c / , zC 2 , and t c j were 0.3 ± 0.1 ms, 0.7 ± 0.1 ms, and 3.9 ± 0.4 ms with 0 mM K+ (n  = 16), 0.3 ± 0.1 ms, 0.8 ± 0.1ms, and 3.7 ± 0.4 ms with 1 mM K+ (n  = 7), 0.2 ± 0.1 ms, 0.6 ± 0.1 ms, and 2.9 ± 0.3 ms with 3.5 mM K+ (n  = 14), and 0.2 ± 0.1 ms, 0.8 ± 0.1 ms, and 3.5 ± 0.6 ms with 20 mM K+ (n  = 7), respectively. F.  Similarly, the proportions of  these three components were also not significantly  different.  The proportion of  rCI  (black  bars), vC2 (grey  bars), and rC3 (hatched  bars) were 0.82 ± 0.02, 0.5 ± 0.02, and 0.03 ± 0.01 with 0 mM K+ (n  = 16), 0.85 ± 0.03, 0.13 ± 0.03, and 0.01 ± 0.01 with 1 mM K+ (n  = 7), 0.85 ±0.02, 0.13 ± 0.02, and 0.02 ± 0.01 with 3.5 mM K+ (n  = 14), and 0.89 ± 0.02, 0.10 ± 0.02, and 0.01 ± 0.01 with 20 mM K+ (n  = 7). Error bars represent the standard error of  the mean (SEM). 5.3.4 External K+ promotes mode A gating Thus far,  external K+ has been shown not to affect  either the unitary current amplitude or the intraburst behaviour. This implies that external K+ antagonizes the H+-induced current inhibition by changing the burst behaviour (burst length and/or interburst duration) and/or channel availability (the proportion of  mode A gating). Mode U  gating was defined  in Chapter 4 as a failure  to open during depolarizing pulses shorter or equal to 1 s. With longer depolarizing pulses, the probability of  observing a transition from  mode U  to mode A was expected to increase. For this reason, depolarizing pulses lasting for  6, 31, and 61 seconds were used. As expected, the proportion of  null sweeps in these long pulses was reduced, and with 0 mM K+0 many sweeps showed delayed opening (long first  latency; Figure 5.8). These delayed openings were assumed not to result from  missed brief  opening events. This was supported by the analysis of  burst duration and the probability of  observing bursts with duration shorter than the time resolution of  the recording system (see below). First latency data were obtained from  idealized unitary current traces recorded at pH 6.4 with 0, 1, 3.5, and 20 mM K+ by measuring the time between the start of  the depolarizing pulse and the first  apparent opening event. With multi-channel patches, the first  latency value for  channel (level) x (where x > 1) was included only if  it occurred within the first  burst of  channel opening (x  - 1). The duration histograms for  first  latency with 0, 1, 3.5, and 20 mM K+ are plotted in Figure 5.4 A. All four  duration histograms show a bimodal distribution, with a peak at around 1 ms and another peak in the range of  seconds. However, as with measuring P0, a maximum estimated error of  2 ms was added to the error of  the latency values due to imperfect compensation of  the capacitive currents. Given an error as large as 2 ms, the fast  component is only an estimate. In addition, no correction of  first  latency for  filter  time delay was performed. Normally, the shape of  the probability density function  for  first  latency is a skewed bell-shaped curve and deviates from  an exponential function  especially in the shorter duration, domain. However, the probability density function  can be well approximated by an exponential distribution with two components for  all durations longer than the mean short first  latency. . Therefore,  the first  latency histograms were fitted  to a double exponential distribution to estimate the mean short latency ( r s l ) and mean long latency (r /L). The values for  rSL and rLL are shown in Figure 5.4 C and E, respectively. Values for  rSL were around 1 to 1.5 ms and did not appear to change significantly  with external K+. This is consistent with the lack of  an effect  of  external K+ on the macroscopic activation kinetics shown in Figure 5.1 H.  The mean long first  latency ( t u ) increased from  2.9 ± 0.4 s with 0 mM K+ to 4.0 ± 0.7 s and 10.3 ± 3.1 s with 1 and 3.5 mM K+. The value of  rLL with 20 mM K+ had a large standard deviation due to the small number of  long latencies (22 events) with this [K+] and was therefore  omitted. Since vLL was well over 1 s, the probability of  seeing sweeps coming out from  mode U  was very low with a 1 s pulse; hence, these sweeps would have appeared as "null" sweeps in Chapter 4. To be consistent with the analysis performed  in Chapter 4 and the definition  of  mode U  (unavailable) gating, the channels with a long first  latency were assumed to have shifted  from  mode U  to mode A gating during the depolarizing pulses. Alternative models are considered (see Discussion), but the above interpretation was favoured  for  its simplicity. 0 mM K+ i 1 ms 10 ms 0.1 s Gap Length 1 mM K l L= 1.0 ± 0.1 ms ,= 4.0 ± 0.7 s 0,1 ms 1 ms 10 ms 0.1 s 1s 10 s Latency to first  opening 3.5 mM K+ = 1.4 ± 0.1 ms = 10.3 ±3.1 s 1 0 -o-0.1 ms 1 ms 10 ms 0.1 s 1s 10 s Latency to first  opening 20 mM K i L = 0.9 ±0.1 ms , = 13.9 ± 11.6S 1 ms 1 ms 10 ms Latency to - f — i r 0 . 1 S 1 8 10 first opening 1 I I T 0.1 ms 1 ms 10 ms 0.1 s 1s Gap Length "i r 0.1 ms 1 ms 10 ms 0.1 s 1 s Gap Length ms 10 ms 0.1 s Gap Length 8 40-OK 1 " 1 K ' 3.5 K 20 K nh [tl rfl 1 K 3.5 K 20 K o J q Figure 5.4 External K+ causes parallel changes in both the mean long first  latency and the mean long gap at pH 6.4. A. First latency histograms at pH 6.4 with 0, 1, 3.5, and 20 mM K+ were generated by measuring the first  latency values from  idealized unitary current traces recorded at 100 mV from  different  cell-attached patches. All four  histograms show bimodal distributions and were fitted  to a double exponential distribution function  to obtain the mean short ( t s /) and long (x u) latencies, which are presented as bar graphs in panel C and E, respectively. B. Gap (interburst) duration histograms were generated from  closed events longer than 20 ms. Each of  the four  gap duration histograms was fitted  to a distribution representing the sum of  three exponential components. The mean duration of  the fastest  component ( r C 3 ) for  each of  the four  histograms was shorter than 20 ms and thus assumed to represent the "tail" of  the slow closed state (C3; see Figure 5.3 D). The data in A and B are presented on the same time scale to emphasize the similarity between xLL and xI G. Values for  rSG and xLG at different  pH are shown in panel D and E (grey  bars), respectively. C. Values of  xs/  were plotted against [K+], and they were not significantly  different  (a maximum error of  2 ms was assumed). The fitted  values of  xSL were 1.0 ms for  0, 1, and 20 mM K+ and 1.4 ms for  3.5 mM K+. D. Values of  xC3, (black bars) and xSG (grey  bars) are shown against pH. The dash line represents the critical time set for this study (20 ms). Values of  r S G changed from  36 ± 6 ms with 0 mM K+ to 59 ± 9 ms, 76 ± 9 ms, and 78 ± 7 ms with 1, 3.5, and 20 mM K+. E. Values of  xLL (black  bars) and xlG (grey  bars) were plotted against [K+], No significant  difference  between the two was detected at any pH. The values for  xLL were 2.9 ± 0.4 s, 4.0 ± 0.7 s, and 10.3 ± 3.1 s with 0, 1, and 3.5 mM K+, respectively, and the values for  xLG were 2.5 ± 0.2 s, 3.0 ± 0.2 s, and 5.8 ± 0.3 s with 0, 1, and 3.5 mM K+, respectively. The error for  xLL with 20 mM K+ was too large and thus omitted. All error bars represent the standard deviation of  the fit. To test the hypothesis that sweeps with a long (first)  latency represented channels shifting from  mode U  to mode A during the depolarizing step, the following  analysis was done. In Chapter 4, the reduction in current by external FT was attributed to the reduction in availability; that is, a promotion of  mode U  gating. For this study, sweeps obtained from  one-channel, cell-attached recordings were divided into three groups based on their first  latency. Sweeps in group 5" had a first  latency shorter than 20 ms; sweeps in group L had a first  latency > 20 ms; and, sweeps in group N  did not show any opening during the entire depolarizing pulse (up to 31 s). The use of  a 20-ms criterion for  group S  was based on the fact  that the first  peak of  the first latency histogram occurred at approximately 2 ms, and a value 10 times longer should clearly separate the short latencies from  the long latencies, given that the mean long latency was in seconds. Figure 5.5 A shows the proportion of  group S, L, and Vat different  [K+]0. From 0 mM to 20 mM K+, the proportion of  sweeps in group S  was increased and the proportion of  sweeps in group L and N  was decreased. If  both group S  and L were combined as "mode A" gating, the availability with 0 mM K+ would be 0.55. Yet, from  Chapter 2, the residual current is predicted to be around 0.2, a number much closer to the proportion of  sweeps in group 5 alone (0.26). In Chapter 4, the proportion of  sweeps in mode A at pH 6.4 with 3.5 mM K+ was calculated to be approximately 0.60, which was comparable to the proportion of  sweeps in group S  (0.63). The current inhibition observed under macroscopic current correlated with the proportion of  sweeps in group S  alone much better than with the combined proportion of  sweeps in group S  and L. These correlations suggest that sweeps belonging to either group L or N  contribute to the current inhibition by H+ and that sweeps belonging to group L start in mode U. In Figure 5.5 A, the proportion of  sweeps in each group was calculated from  all sweeps pooled from  different  patches to compensate for  a very low number of  sweeps obtained from some patches. For that reason, no standard error is given. However, with such a large number of cumulated events (> 150 events in total for  each treatment), the data were considered to be B 1 . 0 - 1 0 . 8 -0 . 6 -0.4 -• Group S (First latency < 20 ms) • Group L (First latency > 20 ms) • Group N  (Null sweep) 0.2 -• n 1.0-1 0 . 8 -0.6 -• Short Gaps (SG) • Long Gaps (LG) 20 K Figure 5.5 External K+ promotes mode A and short gaps between bursts. A. Unitary current traces evoked by depolarizations lasting up to 31 s were categorized into group S  (first  latency < 20 ms), L (first  latency > 20 ms), or N  (null sweeps). Sweeps in group S  were considered to undergo mode A (normal) gating, sweeps in group L were assumed to shift  from  mode U  to mode A during depolarizing pulses, whereas sweeps in group N  were assumed to remain in mode U. The proportion of  sweeps in group S  increased from  0.27 with 0 mM K+ to 0.63, 0.57, and 0.84 with 1, 3.5, and 20 mM K+. Conversely, the proportion of  sweeps starting in mode U  (group L and N)  decreased from  0.74 (0.30 and 0.44) with 0 mM K+ to 0.36 (0.20 and 0.16) with 1 mM K+, 0.42 (0.24 and 0.18) with 3.5 mM K+, and 0.17 (0.08 and 0.09) with 20 mM K+. B. The proportion of  long and short gaps between bursts was different  only with 20 mM K+0. The proportion of  short gaps was 0.48 ± 0.09, 0.48 ± 0.08, 0.46 ± 0.05, and 0.63 ± 0.05 with 0, 1, 3.5, and 20 mM K+, respectively. The proportion of  long gaps was 0.52 ± 0.03, 0.52 ± 0.03, 0.54 ± 0.02, and 0.37 ± 0.02 with 0, 1, 3.5, and 20 mM K+, respectively. The error bars represent the standard error of  the mean (SEM). normally distributed and hence a 5% error was used to determine significance.  Using this criterion, external K+ from  0 to 20 mM significantly  increases the proportion of  mode A (/. e., group S);  there was no significant  change of  the probability of  mode A between 1 and 3.5 mM K+. This result is consistent with the hypothesis that external K+ inhibits mode U  gating. 5.3.5 External K+ slows the onset but not the recovery from  slow inactivation By increasing the number of  long-lived non-conducting events, the use of  long depolarizing steps (6, 31, and 61 seconds) allowed a better characterization of  the microscopic rate of  the onset of,  and recovery from  slow inactivation. In Figure 5.3 D, each of  the four  closed duration histograms showed two components ( r C v , and rC5) that were longer than the critical time (t cnl) of 20 ms as determined in Chapter 4 (Kwan et al., 2006). Since these two components represent < 0.1% of  all the closed events, the resolution was not very good with individual patches. To obtain a better estimate of  the dwell times, all the closed events with duration longer than 20 ms were binned together from  the 7-16 single-channel cell-attached patches recorded with 0, 1, 3.5, and 20 mM K+ to generate their respective gap (interburst) duration histograms (Figure 5.4 B). Each of  the gap duration histograms was fitted  to an exponential distribution function  with three components. The mean duration for  the fastest  component (rCJ ,) of  each of  the histograms was shorter than 20 ms and is assumed to represent the "tail" of  the longest closed state (i.e.,  rcr; see Figure 5.3 D), whereas the other two components (r S G and rLG) were assumed to represent the dwell times of  two inactivated states. Both rC 3, (black  bars) and tsg (mean short gap; grey bars) are shown in Figure 5.4 D. Surprisingly, the value of  rSG increased significantly  from  0 to 3.5 and from  0 to 20 mM K+ (ANOVA; p < 0.05). This result suggests that external K+ increases the dwell time of  a fast  component of  slow inactivation. Similarly, the value of  r 1 G with 3.5 mM K+ was significantly  different  from  the others, but the values of  vLG with 0 and 1 mM K+ were not significantly  different  from  each other. Interestingly, with each [K+]0 tested the values of  vLL and rL G were not significantly  different.  Moreover, rLL and rL G correlate significantly  with each other (r  = 0.93; p < 0.05). This can also be seen by comparing the histograms in Figure 5.4 A and B, in which the duration axes span the same time scale. This approach emphasizes the correlation between the mean (peak) long latency and the mean (peak) long gap. This result suggests that the long first  latency is related to a transition out of  an inactivated state. The proportion of  the short (r SG) and long ( r / c ) gap are shown in Figure 5.5 B to be around 0.5 for  0, 1, and 3.5 mM K+, but the proportion of  short gaps appeared to dominate (0.63 ± 0.05) with 20 mM K+. Together, the above results suggest that recovery from  inactivation may at best be slowed slightly by external K+. ( Panels A-D of  Figure 5.6 show the burst duration histogram with 0, 1, 3.5, and 20 mM external K+, respectively. Surprisingly, given that the macroscopic inactivation time course contains multiple components, each histogram was well fitted  to a single exponential distribution function.  Figure 5.6 E shows the mean burst duration with each of  the [K+]0 tested. A burst was defined,  as in Chapter 4, as a collection of  openings terminated by a closing event longer than tcir, (20 ms). The mean burst length changed from  0.42 ± 0.01 s and 0.44 ± 0.01 s with 0 and 1 mM K+ to 0.37 ± 0.01 s and 0.55 ± 0.01 s with 3.5 and 20 mM K+. The significant  reduction in mean burst length with 3.5 mM K+ is consistent with an acceleration of  slow inactivation observed at the macroscopic level. The above result shows that external K+ causes a small but significant slowing of  the macroscopic slow inactivation kinetics. Using the shortest mean burst duration measured (0.37 s), the proportion of  bursts shorter than 2 ms was -0.5%. That is, with uncertainties in the first  2 ms of  the pulses, at most 0.5% of the long latency sweeps may have grouped inappropriately (i.e.,  in group L instaed of  group 5). On this basis, missed events are not likely to account for  the long first  latencies. i Part  II  Molecular  mechanism for  the slow rising  phase at low pH  with  0 mM  K +0 In this section, the effects  of  changing external pH in 0 mM K+0 are examined in relation to the slow rising phase. As discussed in connection with Figure 5.1 A, there is a prominent slow 0 mM K B 1 mM K 0.1 ms 1 ms 10 ms 0.1 s 1 s Burst duration 0.1 ms 1 ms 10 ms 0.1 s 1 ! Burst duration 10 s •2 0.4 -3.5 mM K D 20 mM K i i i 0.1ms 1ms 10 ms 0.1s 1s 10 s Burst duration OK 1 K 3.5 K 20 K 0.1ms 1ms 10 ms 0.1s 1s 10 s Burst duration Figure 5.6 At pH 6.4 increasing [K+]0 increases the burst duration. Burst duration histograms with 0, 1, 3.5, and 20 mM K+ are shown m.A-D. A burst was defined  as a series of openings terminated by a closing event longer than 20 ms. Each burst duration histogram was fitted  to a single exponential distribution function.  E. The time constants for  the burst duration histograms were 0.42 ± 0.01 s, 0.44 ± 0.01 s, 0.37 ± 0.01 s, and 0.55 ± 0.01 s for  0, 1, 3.5, and 20 mM K+, respectively. The values were significantly  different  from  each other but constituted only a modest effect. rising phase of  current during a 6 s pulse to +60 mV at pH 6.4 that is not evident with 1 mM and higher [K+]0. To investigate this slow rising phase of  current, an approach similar to that employed in Part I was used. Macroscopic and single channel currents were analysed to deduce the microscopic changes that underlie the slow rising phase. 5.3.6 Kvl.5 shows a slow rising phase at low pH with K+-free  solution To characterize the slow rising phase at different  pHs, Kvl.5 currents were recorded with 6 s depolarizing pulses to +60 mV at pH 7.4, 6.9, 6.4, and 5.9 in nominally K+-free  solutions. Representative traces at pH 7.4, 6.4, and 5.9 are shown superimposed in Figure 5.7 A. Consistent with previous findings  (Chen et al., 1997), Kvl.5 current during the 6 s pulse at pH 7.4 inactivated with a time course that was well fitted  by a double exponential function  (see figure  legend for  time constants). After  switching to bath solution at pH 6.4 or 5.9, currents through Kvl .5 were strongly reduced as reported in Chapters 2 and 4. Figure 5.7 B shows the relative peak current.within the first  20 ms ( I p 2 0 , J with respect to that at pH 7.4, which was decreased to 64 ± 8% at pH 6.9, 13 ± 1% at pH 6.4, and 3.1 ± 0.4% at pH 5.9, and this reduction in peak current is comparable to the data obtained with HEK-293 cells (Figure 2.3; 0 mM K+0). As in HEK-293 cells (not shown), currents at both pH 6.4 and 5.9 showed a slow rising phase. The mean time constants for  this slow rising phase ( r s h w r i s e ) at pH 6.4 and 5.9 were 1.44 ± 0.12 s (  n = 11) and 0.47 ± 0.08 s {n  = 7), respectively, and were significantly  different  (ANOVA; p < 0.001). Compared to Ip 2 0 m s , the maximum current amplitude (/m m) at pH 6.4 and 5.9 was potentiated by 1.32 ± 0.04 and 1.47 ± 0.17-fold  due to the slow rising phase (Figure 5.7 C, black  bars). In contrast, since no potentiation was observed at pH 7.4 and 6.9, the maximum current occurred within the first  20 ms. Figure 5.7 C also shows the relative current amplitude at the end of  the 6 s pulse (4 , grey bars) relative to I p 2 0 m s . At pH 7.4 and 6.9, the current amplitudes were reduced to 0.52 ± 0.03 and 0.60 ± 0.03 of  their respective Ip 2 0 m s due to slow inactivation. Conversely, the relative 4 at pH 6.4 and 5.9 was potentiated to 1.26 ± 0.05 and 1.31 ± 0.16 due to the slow rising phase. As reported in Chapter 2, decreasing pH increased the time constant for  activation (racl). Values of  r ac t were obtained by fitting  current traces between 50% and 100% of  1 2 0 m s within the first  20 ms. Figure 5.7 D shows the increase in raa with decreasing pH, and the values were significantly  different  (except between pH 7.4 and 6.9; see figure  legend for  details). This slowing of  the activation kinetics may be due to a rightward gating shift  related to screening of 2 nA [K+]0 = 0 mM ^ 7 . 4 pH 6.4 pH 5.9 B 1 .0 - i 1 s 7.4 6.9 6.4 5.9 PH 2 . 0 - i p, 20ms <u 1 .0 -<r D 3.0 - i 2 . 5 -2 . 0 -7.4 6.9 6.4 5.9 pH 7.4 6.9 6.4 5.9 PH Figure 5.7 Kvl.5 current shows a slow rising phase at low pH with 0 mM external K+. Whole-cell Kvl.5 currents at pH 7.4, 6.4, and 5.9 with 0 external K+ were recorded with voltage commands to +60 mV for  6 s from  a holding potential of  -80 mV. A. Representative Kvl.5 current obtained from  the same cell. A prominent current inhibition, along with a slow rising phase, can be seen in the current traces recorded at both pH 6.4 and 5.9. B. To quantify  the current inhibition with decreasing external pH, the peak current within the first  20 ms of  the depolarizing pulse ( I p 2 0 m w a s normalized with respect to that at pH 7.4. The relative I p 2 0 m s was reduced to 64 ± 8%, 13 ± 1%, and 3 ± 4% at pH 6.9, 6.4, and 5.9, respectively. C. The relative peak current within the 6 s pulse (J mcJI Px2 0 m^  black  bars) and the relative current at the end of  the 6 s pulse (I 6/I P:2oms> g^  bars) at different  pHs are compared. Values at different  pHs were normalized with respect to individual I p 2 0 m s within the group. The slow rising phase at pH 6.4 and 5.9 results in a maximum current that is larger than I p 2 0 m s by 32 ± 4% and 47 ± 17%, respectively. In contrast, l 6JIP t 2 0 m s at pH 7.4 and 6.9 were reduced to 0.52 ± 0.3 and 0.60 ± 0.3, whereas at pH 6.4 and 5.9, the relative I6 s had increased to 1.26 ± 0.05 and 1.31 ± 0.16, respectively. D. External [H+] increases the activation time constant (r acl). By fitting  the current traces between 50% and 100% of  I p20ms to single exponential function,  xact at different  pHs was measured, and it increases from  1.0 + 0.1 ms at pH 7.4 and 0.9 + 0.1 ms at pH 6.9 to 1.5 + 0.1 and 2.3 ± 0.4 ms at pH 6.4 and 5.9, respectively. Error bars represent the standard error of  the mean (SEM). surface  charges by protons (Deutsch and Lee, 1989; Kehl et al., 2002; Trapani and Korn, 2003). To obtain an insight into the mechanistic basis for  the slow rising phase of  current in 0 mM K+0, unitary currents from  Kvl .5 were recorded at different  pHs in the cell-attached mode, and the data were pooled to determine the mean first  latencies, mean gap durations, and mean burst length. 5.3.7 External H+ promotes group L (long first  latency) and N  (null) behaviour As for  the study of  the effect  of  different  [K+]0 (Part I), unitary currents were recorded from cell-attached patches with depolarizing pulses lasting up to 61 s in order to study the microscopic inactivation kinetics. Representative current traces from  different  cell-attached patches evoked by 31 s depolarizing pulses to +100 mV from  -80 mV at pH 7.4, 6.9, 6.4, and 5.9 are shown in Figure 5.8 A-D, respectively. In each panel, the top trace shows a sweep with a short latency (i.e.,  group S),  and the bottom trace shows a sweep with a long latency (i.e.,  group L). Both the intraburst behaviour and single channel current amplitude were shown in Chapter 4 to be virtually unaffected  by decreasing external pH with 3.5 mM K+0, and a similar effect  was observed with 0 mM K+0 (data not shown). From idealized unitary current traces, P0 was determined at different  pHs. Values for  P0 calculated from  the 6 s and 31 s sweeps at different pHs are shown in Figure 5.8 E. Both the 6 s and 31 s pulses showed a.reduction in mean P0 with decreasing external pH, with the 6 s pulses showing a higher P0. The reduction in PQ with 31 s pulses suggests the existence of  a long-lived non-conducting state. Similar to the results obtained when changing [K+]0 at pH 6.4, unitary current traces recorded at different  external pHs with 0 [K+]0 showed three distinct gating behaviours and were divided into group S  (first  latency < 20 ms), L (first  latency > 20 ms), and N  (no channel opening). [K ] = 0 mM 100 mV | -80 mvJ pH 7.4 2 pA pulses 31s pulses Figure 5.8 Unitary current, mean open probability, and the proportion of  sweeps in groups S, L, and N  at different  pHs. Representative unitary current traces recorded from  different  cell-attached patches with 0 mM K+ 0 at +100 mV for  31 s at pH 7.4 (A),  6.9 (B),  6.4 (Q,  and 5.9 (D) are shown. In each panel, the top trace shows a representative sweep with a short latency, whereas the bottom trace shows a sweep from  another patch with long latency. Current amplitudes were unaltered with pH (-1.6 pA at 100 mV in each case). E. Decreasing external pH decreased the mean open probability. Mean open probability (PQ) during 6 s (black  bars) and 31 s (grey  bars) pulses at different  pHs are compared in the bar graph. Error bars represent the standard error of  the mean (SEM). F.  Decreasing external pH also reduced the proportion of sweeps in group S.  Unitary current traces lasting up to 31 s were divided into groups S, L, and N (see Figure 5.5). Group S  contains traces with a short (< 20 ms) latency to first  opening; group L contains traces with long (> 20 ms) latency to first  opening; and, group N  contains traces without opening during a depolarizing pulse lasting up to 31 s. The proportion of  group S, L, and N  were 0.66, 0.13, and 0.21 at pH 7.4, 0.54, 0.28, and 0.18 at pH 6.9, 0.27, 0.30, and 0.43 at pH 6.4, and 0.04, 0.38, and 0.58 at pH 5.9. Figure 5.8 F  shows the proportion of  sweeps in the three groups at the 4 different  pHs. The proportion of  sweeps in group S  decreased significantly  from  0.66 at pH 7.4 to 0.54, 0.27, and 0.04 at pH 6.9, 6.4, and 5.9, respectively. In other words, if  sweeps with long first  latency were assumed to be in mode U  at the start of  the depolarizing pulse, then the proportion of  sweeps starting with mode Ugating  (i.e.,  group L and N)  was increased with external H+. This reduction -in the proportion of  group S  was qualitatively similar to the reduction in availability reported in Chapter 4 (Kwan et al., 2006), which is consistent with the above interpretation of  group L behaviour. 5.3.8 The mean long first  latency correlates significantly  with the mean long gap From the idealized traces, duration histograms for  first  latency, gaps between bursts, and burst lengths at different  pHs were plotted to determine the changes the bursting behaviour. Figure 5.9 A shows the first  latency histograms at pH 7.4, 6.9, 6.4, and 5.9. As in Figure 5.4 A, the histograms fit  well to a double exponential distribution function  except at pH 7.4, for  which very few  long latencies (17 in total) were observed; therefore,  the long latency (rLL) at pH 7.4 is only an estimate. Again, the mean short latency (r sz) corresponds to sweeps in group 5, and the mean long latency (r I L) represents the mean delay in opening of  sweeps in group L. The histograms showed a shift  in the relative distribution towards the longer component with decreasing pH as shown in Figure 5.8 F.  The pH-dependent changes in rs/  and rL[  are shown in Figure 5.9 C and E, respectively. The value of  rSL increased from  0.4 ms at pH 7.4 to 1.0 ms at pH 6.9, 1.0 ms at pH 6.4, and 1.6 at pH 5.9. However, given an estimated maximum error of  2 ms (see Figure 5.4 A), the changes in r s / with pH cannot be resolved unambiguously. The gap (interburst) duration histograms at different  pHs are shown in Figure 5.9 B. As in Figure 5.4 B, the gap duration histograms were best fitted  with an exponential distribution function  with 3 components, except at pH 5.9 where the data were best fitted  with a double pH 7.4 pH 6.9 pH 6.4 pH 5.9 = 0.35 ± 0.02 ms = 2.35 ± 1.3 s 2 0 -15 -10 5 -0 0.1 ms 1 ms 10 ms 0.1 s 1s Latency to first  opening nJ-W Ij 0.1 ms 1 ms 10 ms 0.1 s 1s Latency to first  opening 0.1 ms 1 ms 10 ms 0.1 s 1s Latency to first  opening 0.1 ms 1 ms 10 ms 0.1 s- 1 s Latency to first  opening n i r 0.1 ms 1 ms 10 ms 0.1 s 1s Gap Length 0.1 ms 1 ms 10 ms 0.1 s Gap Length i 1 r 0.1 ms 1 ms 10 ms 0.1 s Gap Length "i r 0.1 ms 1 ms 10 ms 0.1 s 1s Gap Length Figure 5.9 The mean long first  latency correlates with the mean long gap between bursts. A. First latency histograms are generated from  idealized unitary currents recorded at pH 7.4, 6.9, 6.4, and 5.9 with depolarizing pulses at 100 mV. All four  histograms show bimodal distributions and were fitted  to a double exponential distribution function  giving the mean short (x SL) and long (T U)  first  latencies. Values of  RS/  and X U  were summarized in panel C and E, respectively. B. Gaps (interbursts) were defined  as closing events longer than 20 ms in the idealized traces and were binned to generate the gap duration histograms. Histograms for  pH 7.4, 6.9, and 6.4 were best fitted  to distributions representing the sum of  3 exponential components, and the histogram for  pH 5.9 was fitted  to a distribution comprising the sum of  2 exponential components. The fastest  component (rCJ.) for  all four  histograms has a mean value smaller than 20 ms and was assumed to represent the "tail" of  a long closed state (C 3, see Figure 5.3 D). The data in A and B are presented on the same time scale to emphasize the similarity between xLL and xLG. Values for xSG and X]Q  at different  pHs are shown in panel D and E, respectively. C. Values of  T s/, plotted against pH, increased from  0.4 ± 0.1 ms at pH 7.4 to 1.0 ± 0.1, 1.0 ± 0.1, and 1.6 ± 0.3 ms at pH 6.9, 6.4, and 5.9, respectively. D. Values of  xC3, (black  bars) and xSG {grey  bars) at different  pHs were summarized. The dashed line represents the critical time (20 ms). Values of  the mean short gap (xSG) changed from  70.7 ± 10 and 71.1 ± 6 ms at pH 7.4 and pH 6.9 to 36.2 ± 6 ms at pH 6.4. The histogram for  pH 5.9 does not show a component corresponding to short gap. E. Comparison of  the mean long first  latency and the mean long gap. Values of  xLL (black  bars) and xLG (grey  bars) are shown against pH; no significant  difference  was detected between the two values at each pH tested, and xLL and xLG correlated significantly  (r = 0.99 ± 0.08; p < 0.01). The values for  xLL were 2.35 ± 1.3 s, 2.78 ± 0.35 s, 2.87 ± 0.39 s, and 4.53 ± 0.36 s at pH 7.4, 6.9, 6.4, and 5.9, respectively. The values for  vLC were 1.76 ± 0.19 s, 1.77 ± 0.12 s, 2.54 ± 0.18 s, and 4.44 ± 0.40 s at pH 7.4, 6.9, 6.4, and 5.9, respectively. F.  The proportion of  short and long gaps at different  pHs. The proportion of  short gaps was reduced from  0.64 at pH 7.4 to 0.59 and 0.48 at pH 6.9 and 6.4, respectively. Short gaps were not detected at pH 5.9. All error bars represent the standard deviation from  the fit  to the histograms. exponential distribution function.  The fastest  component ( r C 3 ) in all four  histograms had a mean dwell time shorter than 20 ms and was interpreted as the "tail" of  the distribution of  a long closed state. The other 2 components (r S G and vLG) are shown in Figure 5.9 D and E, respectively. With decreasing pH, rSG decreased significantly  from  71 ± 9 ms at pH 7.4 and 71 ± 6 ms at pH 6.9 to 36 ± 6 ms at pH 6.4, suggesting that decreasing external pH reduces the dwell time of  a "fast" inactivated state. This result was unexpected as it appears to contradict the acceleration of  slow inactivation observed in Kvl .5 as discussed in Chapter 4 (Kwan et al., 2006). On the other hand, theproportion of  rSG was reduced with pH as well (Figure 5.9 F).  The proportion of  rSG decreased from  0.64 at pH 7.4 and 0.59 at pH 6.9 to 0.48 at pH 6.4 and 0 at pH 5.9. That is, the overall time spent in all inactivated states may still be increased, which is consistent with the reduction in mean P0 with decreasing external pH (Figure 5.8 E). The absence of  vSG at pH 5.9 is interesting since the macroscopic inactivation time course at pH 5.9 with 3.5 mM external K+ is well fitted  by a single exponential function  (Kehl et al., 2004; Kwan et al., 2006). Figure 5.9 E shows the comparison between rLL and T lg at different  pHs. Both rLL and r / c increase with decreasing pH, from  2.3 ± 1.3 s and 1.8 ± 0.2 s at pH 7.4, to 2.8 ± 0.4 s and 1.8 ± 0.1 s at pH 6.9, 2.9 ± 0.4 and 2.5 ± 0.2 s at pH 6.4, and 4.5 ± 0.4 and 4.4 ± 0.4 at pH 5.9, respectively. The values of  rLL and rI G were not significantly  different  at each of  the pHs tested. Moreover, they correlated significantly  (r  = 0.99 ± 0.08; p < 0.01). These results show an intriguing correlation between the dwell time of  the non-conducting states prior to channel opening and that of  an inactivated state visited after  channel opening. To test this idea that the long first  latency gives rise to the slow rising phase shown in the macroscopic current, ensemble traces were generated from  all the 6 s sweeps with long latency (group L) at either pH 6.4 (Top  row; n = 135 sweeps) or 5.9 (.Bottom row, n = 389 sweeps) and are shown in Figure 5.10. Indeed, the ensemble traces show a slow rising phase which, when fitted  to a single exponential function,  has a time constant of  1.10 s at pH 6.4 and 0.252 s at pH 5.9. These values are comparable to that shown in Figure 5.7 (1.52 s at pH 6.4 and 0.36 at pH 5.9). However, when the sweeps from  groups S  and L were added, the resulting ensemble traces did not show the slow rising phase (data not shown). It is uncertain how this discrepancy occurs (but see Discussion). pH 6.4 Figure 5.10 Ensemble of idealized unitary current traces with long first  latency at pH 6.4 and 5.9. The ensemble traces from  all the unitary current traces with delayed (> 20 ms) first opening (/. e., group S  traces) at pH 6.4 (top)  and 5.9 (bottom)  are shown. The ensemble traces show a slow rising phase, and the red lines represent the best fit single exponential function  to the data. The slow rising phase at pH 6.4 has a time course slower than that at pH 5.9, and the time constant at each pH is comparable to that for  the macroscopic current. The dotted lines represent zero current. In addition to the dwell time in the inactivated states that are related to the microscopic rate of  recovery from  inactivation, the burst duration, which at 100 mV is related to the microscopic rate of  inactivation, was also determined. Figure 5.11 A-D shows the burst duration histograms at pH 7.4, 6.9, 6.4, and 5.9, respectively. As in Figure 5.6, all four  duration histograms show a single component. However, whereas changing [K+]0 at pH 6.4 showed an inconsistent trend (Part I), reducing pH0 at a fixed  [K+]0 (0 mM) decreased the mean burst duration. The mean burst duration decreased from  0.53 ± 0.02 s at pH 7.4 to 0.40 ± 0.01 s at pH 6.9, 0.42 ± 0.01 s at pH 6.4, and 0.22 ± 0.01 s at pH 5.9 (Figure 5.11 E). The mean burst length at pH 6.9 was not significantly  different  from  that at pH 6.4. However, from  pH 7.4 to pH 5.9, the mean burst pH 7.4 B pH 6.9 2 4 0 0 .6 - i I ms 1 ms 10 ms 0.1 s 1s Burst duration (s) 10s 0.1 ms 1 ms 10 ms 0.1 s 1s Burst duration (s) 10s pH 6.4 D pH 5.9 I ms 1 ms 10 ms 0.1 s 1s Burst duration (s) 10s 0.1 ms 1 ms 10 ms 0.1 s 1s Burst duration (s) Figure 5.11 Mean burst durations of  Kvl.5 at different  external pH with 0 external K+. Histograms plotting burst duration at pH 7.4 (A),  6.9 (B),  6.4 (C), and 5.9 (D)  were fitted  to a single exponential distribution function  with the indicated time constant. A burst is defined  as a series of  openings terminated by a closing event longer than 20 ms. The plot of  the time constants versus pH0 in E show that the burst length decreased with decreasing pH. The values of  the mean burst lengths were 0.53 ± 0.02 s, 0.40 ± 0.01 s, 0.42 ± 0.01 s, and 0.23 ± 0.01 s at pH 7.4, 6.9, 6.4, and 5.9, respectively. The mean burst durations at pH 6.9 and 6.4 were not significantly  different;  otherwise, all values are significantly  different  from  each other. Error bars represent standard deviation. duration was significantly  decreased, which was consistent with an acceleration of  slow (P/C-type) inactivation observed at the macroscopic level. A single component in the burst duration histograms is consistent with the hypothesis that slow inactivation proceeds in a sequential manner, which is similar to the model proposed by Loots and Isacoff  (1998). Alternatively, Kvl .5 may inactivate through multiple pathways with similar transition rates such that the peaks cannot be resolved unambiguously. 5.4 Discussion In this study, unitary currents through Kvl.5 were recorded using long depolarizing pulses either at a fixed  pH (6.4) with varying [K+]0 (0, 1,3.5, and 20 mM) or with a fixed  K+0 (0 mM) at various pH0 (pH 7.4, 6.9, 6.4, and 5.9) to determine the changes in microscopic gating (Figure 5.2 to 5.6) and the mechanism leading to the slow rising phase at pH 6.4 and 5.9 (Figure 5.8 to 5.11). The mean dwell times for  the open and closed states within bursts were virtually unaltered by external K+. That is, the transitions between the open state(s) and the "closed" states outside the normal activation pathway were largely independent of  external K+ and H+. Instead, external K+ decreased the probability of  mode U  gating (Figures 5.5 A and 5.8 F),  which was evident as an increased proportion of  sweeps with short latency (< 20 ms, group S)  and a reduction in the proportion of  sweeps either with long latency (> 20 ms, group L) or without open channel activity (group N).  The sweeps with long latency-were interpreted as channels switching from mode U  to mode A, and the null sweeps represented channels that stayed in mode U  or entered a more stable inactivated state from  mode U.  The mean long first  latency ( T u) correlated significantly  with the mean long gap ( r i G ) (Figures 5.4 E and 5.9 E), which suggested that the delayed opening is related to the dwell time in an inactivated state. This result is consistent with the idea that channels in mode U  gating have undergone a slow inactivation process either at rest or from  states visited during activation. The ensemble traces generated from  all 6 s sweeps in group L at either pH 6.4 and 5.9 with 0 mM K+0 resembled the slow rising phase observed at the macroscopic level (Figure 5.10), which suggested that recovery from  mode U  could be the basis for  the slow rising phase of  current observed at low pH. That is, the slow rising phase may reflect  the time dependence for  recovery from  one or more inactivated states. Together, these data show that mode A and mode U  are linked by at least a common inactivated state and by a common closed state at rest (see Scheme 5.1 below). -5.4.1 External K+ and mode U  gating Similar to the classical slow (P/C-type) inactivation in Shaker  (Lopez-Barneo et al., 1993), mode U  gating is inhibited by external K+. The molecular basis for  this inhibition is unknown; however, if  mode U  gating results from  an outer pore inactivation process as proposed, a "foot-in-door" mechanism may also be involved. Compared to its effect  on Kvl.5, external K+ plays a more direct role in regulating current amplitude in some other channels. Removing external K+ has been shown to result in a partial or complete "conductance collapse" in Kvl .4, the large conductance calcium-activated potassium channel (BK), and the plant inward rectifier  K channel KAT1 (Hertel et al., 2005; Pardo et al., 1992; Vergara et al., 1999). In some ShakerlR  mutants and Kvl .3, reducing external K+ was proposed to inhibit current by decreasing channel availability due to inactivation occurring in closed state(s) (Jager et al., 1998; Lopez-Barneo et al., 1993). This reduction in channel availability has clear parallels with the external H+-induced current inhibition in Kvl.5, as seen in this and previous chapters. However, it is uncertain why this regulation of  channel availability by external K+ is pH-dependent in Kvl .5 but not, apparently, in Shaker  (Lopez-Barneo et al., 1993; Starkus et al., 2003). This question may be answered by swapping the outer pore of  Kvl .5 with Shaker  with progressively larger regions to determine if  the outer pore is involved. An unresolved issue during the testing of  external K+ was the unaltered single channel amplitude (Figure 5.3), which was expected to decrease with increasing [K+]0 due to the reduction in driving force.  We cannot provide a satisfactory  explanation for  this result. In Kv2.1, the residue K356 was proposed to regulate the single channel conductance by adopting different  conformations  (Trapani et al., 2006); however, whether a similar regulation of  the single channel conductance exists in Kvl .5 is uncertain. Additional experiments are needed to see whether the rectification  property of  Kvl .5 changes with K+0, as our data suggest the outward current is K+0-independent. In any case, the results in Figure 5.2 F  shows the single channel current amplitude was not significantly  increased by K+0. 5.4.2 Mode U  or not mode U? ( In Chapter 4, mode U  gating was proposed to explain the current inhibition induced by increasing external [FT], The use of  a model incorporating two different  gating modes was justified  since the properties of  the H+-induced current inhibition satisfied  the criteria of  modal gating (Nilius, 1988). First, the two modes had distinct gating kinetics that could consistently be observed in single-channel patches. Second, the proportion of  the two modes of  gating in Kvl .5 could clearly be manipulated by changing external H+. Third, the two modes occurred in different  clusters with 150-ms test pulses repeated up to every 10 s. In this study, the first  two criteria were clearly met. In addition, a transition from  mode U  to mode A could be clearly seen from  the sweeps with long first  latency, which was consistent with the criterion of  a distinct gating behaviour in each mode. Therefore,  the use of  a gating model incorporating two modes is still consistent with the data shown in this chapter. An alternative interpretation for  the long latencies is that the sweeps in group L may represent channels in another mode of  gating. However, this view of  a modal gating is more complex (3 modes instead of  2) but provided no additional insights into the.gating of  Kvl.5; therefore,  this 3-mode model is not considered further. 5.4.3 Mode U  gating and slow inactivation . One of  the key observations in this Chapter is the correlation between vLG and rLL. The simplest structural interpretation of  this result is that the inactivation gate is closed when a channel is in mode U,  such that the channel remains non-conducting even after  S4 is activated and the activation gate is presumed to be open. This view of  mode U  is engendered in the gating model shown in Scheme 5.1, which is offered  as a preliminary model of  Kvl.5 gating at low pH and is adopted from  gating models for  Shaker  (Olcese et al., 1997) and Kvl.5 (Kurata et al., 2001). This model was proposed to explain closed-state inactivation in the truncated form  of Kvl.5. In Scheme 5.1, the top row represents mode gating, whereas the bottom row represents mode U  gating, and an inactivation process is represented by the downward arrows. That is, during mode A gating (top  row), a channel can reach the open state (O')  through the normal closed states (CO  to C4), resulting in a burst of  openings. (O'  in this model represents the composite of  states within a burst including the non-conducting states outside of  the activation pathway and the open states (Hoshi et al., 1994)). A burst of  openings terminates when the channel enters an inactivated state (I). In contrast, at low pH, a closed inactivation process puts the channel in the closed-inactivated state (10)  in the bottom row (mode U),  and upon depolarization, the channel progresses through the closed-inactivated states (10  to 14) to the inactivated state (I)  that is shared between mode U  and mode A. With a sufficiently  long depolarizing pulse, a channel may recover from  state / back to O', which results in a delayed opening at the single channel .level or the slow rising phase of  current at the macroscopic level. It is possible that a channel may switch to the other row during activation (i.e.,  CI  * II,  etc.), but the probability for  these transitions is assumed to be very low based on the transition rates between mode A and mode U  from  both resting state and activated states; however, the rates for the vertical transitions from  the pre-open states have not been determined explicitly from  this study. _ r:i - _ r.7 - - r.d  " < r i » n ^14 Scheme 5.1 A gating scheme proposed for  the modal gating observed in Kvl.5. The top and bottom row represents mode A and mode U  gating, respectively. Increasing [H+]0 promotes the transition to the bottom row, whereas increasing [K+]0 promotes the upward transition. The open state (O') represent the composite of  all the open and closed states within a burst. The inactivated state (I) also represents multiple inactivation processes. This gating scheme is adopted from Olcese et al. (1997) and Kurata et al. (2001). Scheme 5.1 is a slight departure from  early models of  modal gating where no specific reference  was given in relating how the states in each mode were connected to those in other modes (Hess et al., 1984). However, in models of  Ca2+-mediated inactivation of  the L-type Ca2" channel, two modes, Mode Normal and Mode Ca, were connected explicitly through all of  the states in the activation pathway (Shirokov et al., 1998; Tanskanen et al., 2005). In particular, Mode Ca consisted of  Ca2+-inactivated states with a very low probability of  visiting the open state in Mode Ca (0Ca), whereas gating in Mode Normal resulted in bursts of  openings. In this light, Schenie 5.1 is considered to be a reasonable first  approach to describing the modal gating in Kvl.5 at low pH. Although Scheme 5.1 can explain the closed-inactivation of  Kvl.5, it does not replicate all of  the details for  the depolarization-induced inactivation. For example, state I in this model is an oversimplification  of  the inactivation process, which is shown to have at least two components, I SG and I LG, with mean dwell times rS G and rLG, respectively. Since the inactivated state I SG does not show a corresponding component in the first  latency histogram, it is likely not connected directly to mode U  (i.e.,  the bottom row in Scheme 5.1). Moreover, the burst duration histogram contains only a single exponential component, which suggests there may only be a single inactivation process that terminates a burst. Alternatively, there may be multiple processes with similar kinetics that can terminate a burst. Unfortunately,  the connection between the inactivated states and the states within bursts cannot be determined unequivocally from  the limited data presented here. In addition, the assumption that rLG represents the dwell time of  one inactivated state is possibly an oversimplification;  that is, rLL may also represent the mean duration spent in more than one state. 5.4.4 Molecular basis for  the slow rising phase Based on our interpretation of  mode U  gating, the slow rising phase is assumed to reflect recovery from  inactivation. The concept of  recovery (from  inactivation) during depolarization may at first  seem unusual, but this is an innate ability of  Kvl .5 channel even under normal physiological conditions. For example, at pH 7.4 the steady-state current is about 50% with a 6 s pulse (Figure 5.7), which suggests the inactivated state is non-absorbing and inactivated channels can recover from  an inactivated state during a depolarizing pulse. It is possible that, given a large number of  inactivated channels at the start of  a depolarizing pulse (e.g.,  low pH with 0 mM K+0) , a significant  number of  them recoverto the open state to give rise to an apparent slow rising phase. However, the data presented here do not exclude other possibilities, such as slow activation, as the basis for  this slow rising phase. A number of  unresolved issues presented in this chapter warrant caution in our preliminary interpretation of  the molecular basis for  the slow rising phase. For example, although the ensemble of  traces in group L shows a slow rising phase, the ensemble average of  group S  and L sweeps did not show a slow rising phase (data not shown). It is unclear what underlies this discrepancy. The use of  different  voltages between the macroscopic data and the single channel data has been ruled out as a possibility, as unitary currents recorded from  60-mV or 100-mV did not show a substantial difference  that would result in the absence of  the slow rising phase (data not shown). One possibility is that in the cell-attached mode, fewer  channels show group L behaviour (or recovery from  inactivation). In whole-cell recordings, increasing the apparent rate of  slow inactivation by increasing intracellular Mg2+ and/or Ca2+ can inhibit the slow rising phase (data not shown). It is possible that the extent of  channels recovering from  either mode U  gating or from  inactivated states was smaller than that going into an inactivated state with cell-attached patches. This is similar to the absence of  slow rising phase at pH 7.4 with 0 mM K+ even though both group S  and N  sweeps were present; that is, the proportion of  group S  was too small to show the slow rising phase. To minimize the variability, both unitary current and macroscopic current should be recorded in the outside-out configuration  so that a direct comparison can be made. Another issue with the ensemble traces shown in Figure 5.10 was the apparent discrepancy between the mean long first  latency and the time course of  the slow rising phase. The mean long latency ( r u ) became longer with decreasing pH, yet the time course of  the slow rising phase was faster  at pH 5.9 compared to that at pH 6.4. A preliminary explanation for  the difference between tL l and the time constant of  the slow rising phase is the possible involvement of  the burst duration during the slow rising phase. The slow rising phase may be interpreted as an interplay between channels inactivating and channels coming out from  a non-conducting state. At pH 5.9, the burst duration was short (Figure 5.11 D and E) and T ll was relatively long; therefore,  only a small proportion of  channels with relatively short latency contributed to the rising phase, while the channels with longer latencies made up the quasi-steady-state current. In contrast, in the case of  pH 6.4, the mean burst duration was longer and the mean latency was shorter. Upon depolarization, a larger proportion of  channels, even those with longer first latencies, contributed the rising phase before  channels opening with a shorter first  latency started to close. Therefore,  the time course of  the slow rising phase at pH 6.4 was slower than that at pH 5.9. In an extreme hypothetical situation, in which the burst length is much longer than the mean (long) first  latency, the time constant for  the slow rising phase would be the same as that of  the •"N first  latency (i.e.,  the ensemble becomes the cumulative first  latency graph). Together, it is believed that the correlation between first  latency and recovery from  inactivation is genuine, and that recovery from  a slow inactivation process underlies the slow rising phase observed at low pH. To summarize, the mechanism by which external K+ antagonizes the H+-induced current inhibition in Kvl.5 was shown to result from  an inhibition of  mode U  gating. With long depolarizing pulses (> 1 s), some sweeps (group L) showed delayed opening that was interpreted to reflect  channels "recovering" from  mode U  gating into mode A. Furthermore, this long latency correlated with a depolarization-induced inactivated state (long gap), which suggested a common inactivated state visited by both mode Uand  mode A. This delayed opening may also result in the slow rising phase at low pH with 0 external K+, suggesting a link between recovery from  slow inactivation and the slow rising phase. Acknowledgement This study was supported by a grant from  the Canadian Institutes of  Health Research to S.J.Kehl and D.Fedida. We thank Fifi  Chiu for  preparing the cells. D.C.H.K. was in receipt of  a Doctoral Trainee Award from  the Michael Smith Foundation for  Health Research. 5.5 References Chen, F. S., D. Steele, and D. Fedida. 1997. Allosteric effects  of  permeating cations on gating currents during K+channel deactivation. J. Gen. Physiol. 110:87-100. Deutsch, C. and S. C. Lee. 1989. Modulation of  K+ currents in human lymphocytes by pH. J. Physiol. 413:399-413. Hertel, B., F. Horvath, B. Wodala, A. Hurst, A. Moroni, and G. Thiel. 2005. KAT1 inactivates at sub-millimolar concentrations of  external potassium. J. Exp. Bot. 56:3103-3110. Hess, P., J. B. Lansman, and R. W. Tsien. 1984. Different  modes of  Ca channel gating behaviour favoured  by dihydropyridine Ca agonists and antagonists. Nature 311:538-544. Hoshi, T., W. N. Zagotta, and R. W. Aldrich. 1994. Shaker  potassium channel gating. I: Transitions near the open state. J. Gen. Physiol. 103:249-278. Jager, H., H. Rauer, A. N. Nguyen, J. Aiyar, K. G. Chandy, and S. Grissmer. 1998. Regulation of mammalian Shaker-related  K+ channels: evidence for  non-conducting closed and non-conducting inactivated states. J. Physiol. 506:291-301. Kehl, S. J., C. Eduljee, D. C. Kwan, S. Zhang, and D. Fedida. 2002. Molecular determinants of the inhibition of  human Kvl.5 potassium currents by external protons and Zn2+. J. Physiol. 541:9-24. Kehl, S. J., S. Zhaiig, and D. Fedida. 2004. Depolarization of  Kvl.5 channels in acidic, K+-free external medium evokes a slowly rising phase of  current. Biophys. J. 86:534a. (Abstr.) Kurata, H. T., G. S. Soon, and D. Fedida. 2001. Altered state dependence of  C-type inactivation in the long and short forms  of  human Kvl.5. J. Gen. Physiol. 118:315-332. Kwan, D. C., C. Eduljee, L. Lee, S. Zhang, D. Fedida, and S. J. Kehl. 2004. The external K+ concentration and mutations in the outer pore mouth affect  the inhibition of  Kvl.5 current by Ni2+. Biophys, J. 86:2238-2250. Kwan, D. C., D. Fedida, and S. J. Kehl. 2006. Single channel analysis reveals different  modes of Kvl .5 gating behavior regulated by changes of  external pH. Biophys. J. 90:1212-1222. Loots, E. and E. Y. Isacoff.  1998. Protein rearrangements underlying slow inactivation of  the Shaker  K+ channel. J. Gen. Physiol. 112:377-389. Lopez-Barneo, J., T. Hoshi, S. H. Heinemann, and R. W. Aldrich. 1993. Effects  of  external cations and mutations in the pore region on C-type inactivation of  Shaker  potassium channels. Receptors. Channels 1:61-71. Nilius, B. 1988. Modal gating behavior of  cardiac sodium channels in cell-free  membrane patches. Biophys. J. 53:857-862. Olcese, R., R. Latorre, L. Toro, F. Bezanilla, and E. Stefani.  1997. Correlation between charge movement and ionic current during slow inactivation in Shaker  K+ channels. J. Gen. Physiol. 110:579-589. Pardo, L. A., S. H. Heinemann, H. Terlau, U. Ludewig, C. Lorra, O. Pongs, and W. Stuhmer. 1992. Extracellular K+ specifically  modulates a rat brain K+ channel. Proc. Natl. Acad. Sci. U. S. A. 89:2466-2470. Saftenku,  E., A. J. Williams, and R. Sitsapesan. 2001. Markovian models of  low and high activity levels of  cardiac ryanodine receptors. Biophys. J. 80:2727-2741. Shirokov, R., G. Ferreira, J. Yi, and E. Rios. 1998. Inactivation of  gating currents of  L-type calcium channels. Specific  role of  the alpha2-delta subunit. J. Gen. Physiol. 111:807-823. Starkus, J. G., Z. Varga, R. Schonherr, and S. H. Heinemann. 2003. Mechanisms of  the inhibition of  Shaker  potassium channels by protons. Pflugers  Arch. 447:44-54. Tanskanen, A. J., J. L. Greenstein, B. O'Rourke, and R. L. Winslow. 2005. The role of  stochastic and modal gating of  cardiac L-type Ca2+ channels on early after-depolarizations.  Biophys. J. 88:85-95. Trapani, J. G., P. Andalib, J. F. Consiglio, and S. J. Korn. 2006. Control of  single channel conductance in the outer vestibule of  the Kv2.1 potassium channel. J. Gen. Physiol. 128:231-246. Trapani, J. G. and S. J. Korn. 2003. Effect  of  external pH on activation of  the Kvl.5 potassium channel. Biophys. J. 84:195-204. Vergara, C., O. Alvarez, and R. Latorre. 1999. Localization of  the K+ lock-in and the Ba2+ binding sites in a voltage-gated calcium-modulated channel. Implications for  survival of  K+ permeability. J. Gen. Physiol. 114:365-376. Wang, Z., X. Zhang, and D. Fedida. 2000. Regulation of  transient Na+ conductance by intra- and extracellular K+ in the human delayed rectifier  K+ channel Kvl.5. J. Physiol. 523:575-591. Zar, J. H. 1984. Biost'atistical Analysis. 2 Ed. Prentice-Hall. Upper Saddle River, New Jersey. 6. Slow inactivation in Kvl.5 The molecular mechanism for  the external H+-, Zn2"-, and Ni2"-induced inhibition of human Kvl.5 (/zKvl.5, or simply Kvl.5) current has been investigated. Some of  the changes in the biophysical properties of  Kvl.5 induced by these ions are listed in Table 6.1, and the major findings  that relate to slow inactivation are summarized and discussed in the following  section. 6.1 Mechanism for  the external cation-induced current inhibition In a previous study, Kvl.5 current was shown to be inhibited by external Zn2+, and this inhibition was antagonized allosterically by external K+ (Zhang et al., 2001b). In search of  the binding site for  Zn2+ that mediates this inhibition, a histidine residue (H463) in the turret came to our attention (Steidl and Yool, 1999). When Kvl .5 currents were recorded at low pH, a prominent inhibition was observed, and we showed that this inhibition was also antagonized by external K+ in an allosteric fashion  (Figures 2.2-2.4). The similarities between the effects  of  H+ and Zn2+ suggested that the mechanism by which these ions inhibited Kvl .5 was similar and possibly involved the same binding site. To test further  whether H463 mediated the Zn2+- and H+-induced current inhibition in /zKvl.5, H463 was mutated to glutamine (H463Q), as the equivalent mutation in rKvl .5 (H452Q) was found  to reduce H+-sensitivity (Steidl and Yool, 1999). As expected, this mutation attenuated current inhibition induced by these ions and shifted  the concentration-response curves for  both H+ and Zn2+ to the right (Figure 2.6). Other ligands for  histidine, such as Ni2+, were also > shown to inhibit Kvl .5 (Chapter 3) and this effect  was similarly attenuated by the H463Q mutation. These results were consistent with H463 being the binding site for  H+, Zn2+, and Ni2+ to inhibit Kvl.5. Table 6.1 Comparison of  the effects  of  external Zn2+, Ni2+, and H+ in Kvl.5. Listed below are some of  the biophysical parameters measured either in control condition (pH 7.4, 3.5 mM K+0) or with Zn2+, H+, or Ni2+. Values for  V, / 2 and slope factor  (k)  were obtained by fitting  the corresponding date to a Boltzmann function.  The time constants for  activation (Tacl lvanon), deactivation (Tdeacl jvat l0n), inactivation (Tmacl), and recovery (T recover )) were measured by fitting  the data with single exponential functions  (see the Method sections in Chapter 2 and 3 for  detail). Availability is defined  as the proportion of  sweeps (duration < 1 s) showing channel activity. Control Zn2* H+ Ni2' Macroscopic current properties Activation -10.2 ± 0.4 mV +21.1 ±0.7 mV (1 mM)2 +3 ± 4 mV (pH 6.4) 0.4 ± 0.9 mV (0.25 mM) slope factor  (k) 6.8 ±0.4 mV 9.4 ±0.7 mV (1 mM)2 9.2 ± 0.3 (pH 6.4) No change (0.25 mM) ^activation 1.76 ms (+50 mV) 16.9 ms (1 mM; +50 mV)! 2.3 ±0.4 ms (pH 5.9; +60 mV) No change (0.25 mM)' Deactivation ^deactivation 15.1 ms (-40 mV) 9.1 ms (-40 mV; 1 mM)2 11.0 ± 0.4 ms (-50 mV; pH 5.9) No change (0.25 mM) Current inhibition K d 69 nM (0 mM 1CJ2 153 ± 13 nM (pK„ -6.8; 0 mM K*0) 150 ± 10 |iM (0 mM K'J Hill coefficient 0.892 1.5 ±0.2 1.3 ± 0.1 IC-dependence (K K) -500 nM2 680 ± 90 nM -1.0 mM (est.) Inactivation W , (+50 mV) 2.63 ±0:11 s(5 mM n 2.14 ± 0.16 s (1 mM; 5 mM K+0)2 1.19 ± 0.04 s(pH 6.4; 5 mM K+„) 1.71 ± 0.07 s (1 mM; 3.5 mM K+0) Tre'coven-  ("80 ITlV) 4.3 s 27.7. ±2.1 s(2 mM)2 4.2 ±0.1 s (pH 5.4) 23.5 ±2.1 s (2 mM) Gating current properties Relative Q m a x 1 • 0.85 (1 mM)1 1 (pH 5.5) 1 (1 mM) . K . A 0 O . : ) -2.2 mV 60.2 ± 1.5 mV (1 mM)1 48.9 ± 1.2 mV (pH 5.4) 2.2 + 0.8 mV (1 mM) Single channel properties • / Single channel current / conductance I.7 pA (+100 mV)/ II.8 pS ND3 I.7 pA (+100 mV)/ II.3 pS 1.6 pA (+100 mV)/ ND3 Availability 0.97 NDJ 0.18 (pH 5.9) 0.3 (0.5 mM) ' From Zhang et al. (2001 a) 2From Zhang et al. (2001b) 'Not Determined After  showing the binding site for  these cations, the next step was to deduce the mechanism for  the current inhibition. Based on the KcsA structure, the distance between H463 / o and the central axis of  the pore was -14-16 A, which was probably too far  for  H+ to occlude the conducting pore directly (Aiyar et al., 1995; Doyle et al., 1998). Therefore,  we surmised that H+, and possibly Zn2+, probably did not cause a fast  block in Kvl .5. Conversely, the inhibition induced by Zn2+, H+, and Ni2+ showed a property that was similar to the classical slow (P/C-type) inactivation, namely, inhibition by external K+. We tested this idea by measuring the current inhibition in the mutant in which the outer pore residue R487 was changed to valine (R487V), as the homologous mutation in Shaker  (T449V) was shown to inhibit slow inactivation (Lopez-Barneo et al., 1993). Indeed, the current inhibition was dramatically antagonized by this mutation (Figure 2.8 and 3.5), which suggested that a slow inactivation process was involved in the cation-induced current inhibition. A number of  findings  support a conclusion that this slow inactivation process proceeded from  resting closed states rather than from  depolarized/activated states. First, the current inhibition induced by FT or Ni2+ was "instantaneous;" that is, the extent of  inhibition remained the same after  a complete solution exchange to that at low pH or with Ni2+ (Figures 2.9 and 3.3). Second, at concentrations that produced roughly the same amount of  inhibition, the kinetics of the onset of,  and recovery from,  slow inactivation were still much faster  at low pH than with metal divalent cations (Figures 3.4 and 3.9; Table 6.1). This result was inconsistent with the hypothesis that the current inhibition resulted from  channels that failed  to recover from  an inactivated state in the interval between pulses. Third, the maximum gating charge movement (Qmax) w a s unaltered by H+ (Figure 2.10) and Ni2+ (Figure 3.7), which suggested that the activation machinery was largely unaffected  by these ions even under conditions where a large proportion of  channels were unable to conduct. Based on these results, it was hypothesized in Chapter 3 that the binding of  cations to H463 promoted a closed-inactivation process which resulted in a reduction in the number of  channels available for  activation and hence a reduction in the macroscopic current amplitude. A similar hypothesis of  reduction in channel availability was proposed to explain the reduction in current amplitude with lowering external K+ in fast-inactivating Shaker  mutants (Lopez-Barneo et al., 1993) and in Kvl.4 (Pardo et al., 1992). To gain some mechanistic insights into the cation-induced current inhibition, unitary currents were studied. The above hypothesis of  a reduction of  channel availability predicted that the number of  blank (null) sweeps should be increased. This prediction was confirmed  both with Ni2+ (Figure 3.6) and with H+ (Figures 4.1 and 4.2). In addition, the proportion of  null sweeps correlated well with the extent of  current inhibition (Figure 4.3). At the same time, the single channel current amplitude was unaltered by either Ni2+ or H+. The latter result directly refuted the hypothesis that H+ and the divalent cations inhibit macroscopic Kvl.5 current by a "fast block" mechanism. Furthermore, the unaltered gating behaviour within bursts (Figure 4.6) is inconsistent with the hypothesis that H+0 acts as an "intermediate" block (increased flickering within a burst and prolonged burst length). Together, the single channel results show that FT, and possibly Zn2+ and Ni2+, act as "slow blockers" that modify  channel gating. The change in the gating behaviour induced by H+0 was proposed to be a promotion of Kvl .5 from  an available mode (mode A) of  gating to an unavailable mode (mode U)  of  gating. During mode A gating, a channel can open and conduct current (giving rise to active sweeps), whereas during mode U  gating, a channel remains non-conducting despite normal gating charge movement (resulting in null sweeps). In other words, mode U  gating was suggested to result from  the previously proposed closed-inactivation process which underlies the cation-induced current inhibition. Based on this modal gating model, conditions that attenuate the H+-induced current inhibition, such as increasing external [K+], were predicted toinhibit mode U  gating. This was confirmed  by an increase in the proportion of  sweeps in mode A with increasing [K+] (Figure 5.5). By using long depolarizing pulses (> 6 s), some channels were seen to switch from mode f/back  to mode A, which appeared as sweeps showing long first  latencies. That is, the long first  latencies represented the duration that a channel spent in one or more states of  mode U during a depolarizing pulse. The mean long latency was found  to correlate with the mean long gap duration between bursts, and the latter value was taken to represent the mean dwell time in some inactivated states during a depolarizing pulse. Together, these results suggest mode A and mode U  are connected through some depolarization-induced inactivated states, and support the hypothesis that a closed-inactivation process with many of  the properties of  slow inactivation underlies mode U  gating. 6.2 Implications of  slow inactivation gating For the remainder of  this chapter, the term slow inactivation is used to describe the process by which the outer pore becomes constricted, without regard to the particular state from which this process occurs. Conversely, the term depolarization-induced inactivation is used to describe the inactivation process that proceeds from  any states within in a normal burst of openings during a depolarizing pulse. These terms are so defined  in hope of  avoiding any confusion  in the following  discussion of  the possible mechanism for  the cation-induced current inhibition and mode U  gating. 6.2.1 Mode U  gating versus U-type inactivation Based on the proposed mechanism for  mode U  gating, it is worthwhile to compare it with other known closed-inactivation processes. U-type inactivation is proposed to occur from  closed states, and it is defined  by a number of  biophysical properties (Klemic et al., 2001). The most obvious one is the "U-shaped" inactivation curve, from  which the term "U-type" is derived. However, U-type inactivation seems to have little in common with mode U  gating. For example, a U-shaped inactivation curve is not observed with the current inhibition induced either by H+ (Figure 2.2), Ni2+ (Figure 3.1), or Zn2+ (Zhang et al., 2001b) in Kvl.5, suggesting that U-type inactivation was not involved. In addition, although it can occur in truncated Kvl .5 (Kurata et al., 2002), U-type inactivation has never been shown with full-length  Kvl.5; conversely, mode U is shown in wt Kvl.5. A second property of  U-type inactivation is its sensitivity to the holding potential (Klemic et al., 2001); that is, by changing the holding potential from  -80 to -30 mV, more channels dwell in the 'pre-open" states from  which the transition to U-type inactivated state is faster,  and hence more channels are inactivated. However, most experiments performed  with regard to the H+-induced current inhibition were done at a holding potential of-80  mV, yet mode \ U  was consistently observed. In addition, at pH 5.9 with 0 mM K+0 raising the holding potential to -30 mV had no effect  on the H+-induced current inhibition (personal communication, Dr. S. J. Kehl). Both of  these results are inconsistent with an involvement of  U-type inactivation. A third property of  U-type inactivation is the "reversed" sensitivity to external TEA+ and K+, both of which promote U-type inactivation (Klemic et al., 1998; Klemic et al., 2001; Kurata et al., / 2005). However, mode U  gating was inhibited  by K+0 (Figure 5.5). A fourth  property is the "excessive cumulative inactivation," which arises because repetitive pulsing results in a greater ' extent of  inactivation (Aldrich et al., 1983; Klemic et al., 2001). Repetitive pulsing does not increase the extent of  the H+-induced current inhibition (Figure 2.9). Together, the properties of the cation-induced current inhibition are inconsistent with that of  U-type inactivation and point to a distinct form  of  closed-inactivation process in mode U  gating. Is this closed-state inactivation related to slow (P/C-type) inactivation? 6.2.2 Slow inactivation from  closed states The concept of  an inactivation process proceeding from  the resting closed state is not new. It has been used to explain the reduction in channel availability in various Kv channels (Lopez-Barneo et al., 1993; Pardo et al., 1992; Steidl and Yool, 1999; Teisseyre and Mozrzymas, 2006; Yang et al., 1997), and a P-type inactivation process may be involved. P-type inactivation was originally proposed to be distinct from  C-type inactivation based on the finding  that external TEA+ increased the current amplitude in addition to causing a slowing of  the time-dependent decay in the chimeric Kv2.1-3.1 V369S mutant (De Biasi et al., 1993). In that report, a stabilization of  transitions from  an inactivated state to both the resting closed state and the open state was proposed to account for  the paradoxical potentiation of  the peak current by low concentrations of  TEA+. That is, some channels proceeded directly from  the closed state to the inactivated state, and the recovery of  these channels into the open state gave rise to the potentiation. This is similar to our hypothesis that the potentiation of  current during the slow rising phase of  Kvl .5 current at low pH with 0 mM K+0 (Figure 5.7) is due to channels recovering from  mode U  gating and/or an inactivation process. Another possible example of  P-type inactivation proceeding from  closed state is the Shaker  W434F mutant (Yang et al., 1997). With more subunits expressing phenylalanine at position 434 in a concatameric construct, the rate of  inactivation became faster,  and when all the W434s were mutated to phenylalanine, the channel was predominantly inactivated. However, the W434F homomeric mutant was shown to have normal gating charge movement (Perozo et al., 1993), which suggests the mutant either inactivates from  closed states or the depolarization-induced inactivation rate exceeds the resolution of  the recording system. In the ShakerlR 434WFWF concatamer, active sweeps and null sweeps were seen to be clustered. In addition, the intraburst behaviour was unaltered by the mutations. The above observations were ( surprisingly similar to that shown for  the H+-induced inhibition presented in this dissertation (Figures 2.10, 4.3, 4.4 and 4.6). This parallelism argues for  a similar mechanism underlying both the non-conducting W434F mutation and the cation-induced current inhibition; that is, a P-type • inactivation process. However, as conceded by Yang et al. (1997), we cannot rule out the possibility of  ultra-short bursts that are not resolved because of  the limited response time of  our recording system. The model of  a slow inactivation process involved in the cation-induced current inhibition is very attractive based on the comparison between our data and previous studies. However, this interpretation appears to be inconsistent with some of  the properties ascribed to mode U  gating and depolarization-induced slow inactivation. The most obvious difference  between the two is • their K+-dependence (Chen et al., 1997). In fact,  compared to the depolarization-induced inactivation, mode U  gating is more similar to the "typical" slow (P/C-type) inactivation; that is, mode U  gating is inhibited by external K+ and (probably) by the R487V mutation, whereas the depolarization-induced inactivation is relatively insensitive to both K+0 and the R487V mutation. In addition, the K D for  the K+ effect  on slow inactivation in Shaker  (1 mM) estimated by Baukrowitz and Yellen (1996) was very similar to the K D for  the K+ effect  on the current inhibition induced by Zn2+, H+, and Ni2+ (0.5-1 mM; Figures 2.3 and 3.2). We do not yet have a satisfactory  explanation for  why external K+ and R487V do not inhibit the depolarization-induced inactivation in Kvl .5 under physiological conditions. However, external K" did not attenuate slow inactivation in the ShakerlR  434WWWW concatamer (the residue at position 434 was tryptophan in each of  the four  subunits) expressed mXenopus  oocytes (Yang et al., 1997). This phenomenon was proposed to result from  K+ accumulation at the pore mouth. Yet, the current traces recorded with 0 and 1 mM K+ consistently show significant  differences  (e.g., Figure 5.1), suggested that the accumulation of  K+ at the outer pore was minimal and hence could not be the reason for  the K+-insensitivity. Similarly, the Shaker  T449V mutant expressed in HEK-293 cells, as opposed to Xenopus  oocytes, was found  to have an inactivation rate similar to that of  wt Shaker  (Holmgren et al., 1996). These results suggest the regulation of  slow inactivation may be more complicated than once thought, and these phenomena may warrant additional attention in the future. 6.2.3 Possible mechanism for  closed state inactivation A number of  results shown in this dissertation and in other studies are inconsistent with the dogma that slow inactivation is coupled to the open state. That is, closed state inactivation may arise by an uncoupling of  the inactivation machinery from  the activation apparatus. This idea is consistent with the result that the kinetics of  the ow-gating charge and Qmax are unaltered by external H+ (Figures 2.10 and 3.7), which suggests S4, and possibly the activation gate, is functional  even in channels exhibiting mode U  gating. This view of  a possible uncoupling between the activation and inactivation machineries is supported by the uncoupling of  gating charges and recovery from  slow inactivation in Kvl .5 reported by Wang & Fedida (2002), in which the kinetics of  gating charge recovery is faster  than the kinetics of  recovery from  slow inactivation, suggesting, at least during recovery, that the movement of  the gating charges (and the activation gate?) can be uncoupled from  the movement of  the slow inactivation gate. As mentioned, P-type inactivation has been suggested to proceed from  closed states, at least in mutated channels (De Biasi et al., 1993; Yang et al., 1997). Furthermore, it should be noted that depolarization is not required for  Kvl.5 to be.in mode U(i.e.,  closed-inactivated; Figures 2.9 and 4.1), which suggested that this inactivated state did not result from  channels failing  to recover from  slow (P/C-type) inactivation as in the case in Kvl .4 at low pH. However, the physical basis for  the coupling between slow inactivation and activation is currently unknown, so a coherent model cannot be constructed. Given its role in activation, S4 is expected to play a role in the coupling, but the details remain unknown. 6.2.4 Possible interactions between H463 and the outer pore Compared to the physical coupling between inactivation and activation, the interaction between the turret and slow inactivation is clearer. At the very least, the binding of  H+ or other cations to H463 appears to result in a conformational  change involving R487. From this relationship, the most obvious question is how these two residues might interact. A direct electrostatic interaction between H463 and R487 has been considered in Chapter 2, but based on the finding  that reducing ionic strength has no effect  on the extend of  current inhibition (not shown), a direct electrostatic interaction between the two residues has been deemed unlikely. From the KcsA structure (Doyle et al., 1998), the distance between the a-carbons (Ca) of  Q58 o (homologous to H463) and Y78 (homologous to R487) is approximately 8 A, but their side o chains are much further  apart (~12 A) and point in different  directions. Therefore,  a direct interaction between H463 and R487 is not favoured. Based on the KcsA structure, H463 may interact with two regions. The first  one is the pore helix, which has been implicated in slow inactivation, with the best example being Shaker W434 (or Kvl .5 W472). As mentioned, the effect  of  the W434F mutation on Shaker  is very similar to the effect  of  H+ on Kvl .5, which raises the possibility that a similar mechanism is involved. The side chain of  W434 is proposed to form  a "hydrophobic cuff'  that interacts with the side chains of  the tyrosine residue in the selectivity filter  (Y76) thus holding it in the open conformation  (Doyle et al., 1998). Changing this tryptophan residue to phenylalanine removed this interaction, and the channel became permanently P-type inactivated (Yang et al., 1997). Similarly, any movement of  the pore helix may also weaken the wt W67-Y78 interaction. Since the pore helix is likely to move as a rigid rod (as with other a-helical structures), any movement along the length of  the pore helix may result in significant  movement at W67. In Shaker, mutating the aspartate residue at position 431 (at the top of  the pore helix) to asparagine (D43 IN) accelerates slow inactivation slightly (Loots and Isacoff,  2000). When mapped onto the KcsA structure, this residue is approximately 1 "turn" away from  W434 and is proposed to interact with the turret residue at position 424 (Loots and Isacoff,  2000), which is homologous to Kvl.5 T462. In addition, the side chain of  this residue (D469 in Kvl .5) is about 6 A away from  that of Kvl.5 H463. The charge on H463 may interact with that on D469, and such interaction may affect  the tilting/orientation of  the pore helix at the N-terminal end, resulting in the breaking of  a hydrophobic interaction between W472 and Y483 and hence P-type inactivation. Another region with which H463 may interact is a short stretch of  polypeptide between the selectivity filter  and the start of  the P-S6 linker. Based on the KcsA structure, the side chain of  H463 is only ~5 A away from  the side chain of  a methionine (M486) between a conserved aspartate (D485) and R487: In Shaker,  mutation of  M448 (homologous to Kvl .5 M486) to cysteine has been shown to accelerate slow inactivation in the presence of  Cd2+ (Liu et al., 1996). M448, along with T449 and P450, has been suggested to move during slow inactivation (Liu et al., 1996), and it is possible that a conformational  changes of  any one of  these residues could affect  the orientation of  the side chain of  the other two residues. In addition, the side chains of these residues seem to be inaccessible to MTS reagents applied from  the external milieu during slow inactivation, and Liu et al. (1996) envisioned the side chains of  T449 may "flip"  and become exposed to the external milieu. In a similar vein, the side chain of  D80 in KcsA (homologous to Kvl .5 D485 or Shaker  D447) has indeed been shown to "flip"  or rotate about 180°, and it has been proposed that a large rotation about the polypeptide backbone of  the residues in this outer pore region is possible. It is conceivable that a protonated Kvl .5 H463 may affect  the conformational  change of  M486 or other neighbouring residues and thus accelerate slow inactivation. 6.2.5 Gating within bursts and depolarization-induced inactivation Besides mode U  and closed state inactivation, the states within bursts and the depolarization induced inactivated states were also studied. The single channel analyses in both Chapters 4 and 5 have consistently shown that Kvl .5 has 2 open states and 3 non-conducting states within bursts. In Shaker,  only 1 open state was observed, but 3 non-conducting states were found  within bursts (Hoshi et al., 1994). It is uncertain if  the presence of  two functional  open states in Kvl .5 is significant,  as both of  these states appear to have the same conductance. It is possible the open state with shorter dwell time may be an artifact  arising from  the flickery behaviour of  Kvl .5. The mean dwell time for  the three non-conducting states were similar between Kvl .5 and Shaker,  suggesting their pore structures are very similar. Unfortunately,  no physical basis for  the "closed" states outside the normal activation pathway is known, and the connections between these states and the open state(s) are unknown. It is also unclear whether these closed states are involved during depolarization-induced inactivation, and the study of  these state may require additional structural information  and/or some methods to separate these state from  one another. To my knowledge, no report has investigated depolarization-induced inactivation in detail at the single channel level; therefore,  a direct comparison between the results found  in this study and others is not possible. Two depolarization-induced inactivated states (I SG and I / G) were identified  from  the gap duration histograms shown in Chapter 5 (Figures 5.4 and 5.9). In addition, given a mean dwell time of  ~4 s for  I lG, another inactivated state (/;V) should be included to account for  the null sweeps lasting for  more than 60 seconds. If  mode U  gating results from  a process other than a failure  of  recovery from  inactivation, I N  is likely not connected directly to any closed-inactivated states (10-14).  However, how I N  is connected to the other inactivated states is unclear. These three inactivated states may represent three different  steps in slow (P/C-type) inactivation. For example, since mode £/gating was only seen to connect to I LG, this state may represent a P-type inactivated state. If  this is the case, the inactivated state I N  may represent a C-type inactivated state, since the time course of  C-type inactivation, as judged by charge immobilization, is slower than P-type inactivation (Loots and Isacoff,  2000). However, it is uncertain what process underlies I SG. In summary, the binding of  H+ and other cations to H463 promotes a closed state inactivation resembling P-type inactivation. This effect  may result from  a conformational  change of  the pore helix and/or the outer pore that uncouples the inactivation machinery from  the activation apparatus. However, several issues with regard to the coupling between activation and slow inactivation have not yet been resolved. More direct structural information  on the conformational  changes during closed-state and open state inactivation is needed to show how if they are related. 6.3 Comparison of  H+-induced effects  in Kvl.5 and in other channels In addition to Kvl.5, a number of  Kv channels are sensitive to changes of  the extracellular pH. The binding sites for  H+0 in these channels are mainly in the pore domain versus the turret region. Comparing the effects  of  H+ between that in Kvl.5 and that in other channels may provide addition insights into the modulation of  slow inactivation and other biophysical processes. 6.3.1 External H+ modulation of  Shaker One of  the standards in the study of  Kv channels, especially in the context of  slow inactivation, is Shaker,  and in particular, the N-type /nactivation removed ShakerlR  channel. In ShakerlR,  the current amplitude was reduced by 25% at pH 5.0 in 2.5 mM K+0 (pK H  = 4.7), an effect  that coincided with an acceleration of  slow inactivation (the time constants were ~1.64 s at pH 7.4 and ~190 ms at pH 5.5) (Starkus et al, 2003). The inhibition of  current at low pH in ShakerlR  was roughly 1.5 orders of  magnitude less sensitive to external H+ than wt Kvl .5 but similar to that of  the H463Q (pK H  = 5.3) and R487V (pK H  = 4.6) mutants. A comparison of  the turret of  Kvl .5 and ShakerlR  reveals that the site homologous to Kvl .5 H463 in Shaker  is F425, which does not constitute a H+ binding site. There are four  acidic residues (E418 and E422 in the turret, D431 in the pore helix, and D447 in the outer pore mouth) near the external surface  that might act as H+ binding sites, and Starkus et al. (2003) proposed the outer pore aspartate (D447) residue as the H+ binding site as this residue was proposed to modulate slow inactivation (Olcese et al., 1997). In contrast to our result with Kvl .5, the authors proposed an acceleration of  slow inactivation was primarily involved in the inhibition of  peak current. This idea was supported by the finding  that the effects  of  H+ in ShakerlR  were antagonized but not eliminated by the outer pore mutation T449V (R487V in Kv 1.5). The results of  the H+-induced current inhibition in ShakerlR  might be relevant to our findings  in the Kvl.5 H463Q and R487V mutants. Neither of  these mutations eliminated the H+-induced current inhibition but instead shifted  the concentration-response curve to the right (Figures 2.6 and 2.8). This latter result suggests a second, lower-affinity,  binding site for  H+ exists in Kvl .5. Given a similar sensitivity to external FT in ShakerlR  and in these Kvl .5 mutants, a common mechanism may be involved in both channels. This "second" mechanism may be studied in the Kvl .5 mutants at low pH by recording their unitary currents. 6.3.2 External H+ modulation of  Kvl.4 The Shaker-related  Kvl .4 channel, which exhibits intrinsic N-type inactivation, also shows an external H+-dependent current inhibition (Claydon et al., 2000; Claydon et al., 2002; Ishii et al., 2001). Alignment of  the Kvl .5 and the Kvl .4 sequence (Figure 1.3 A) shows the turret, the P-loop, and the P-S6 linker of  the two channels are very similar. There is a histidine residue (H508) at the site homologous to Kvl .5 H463, and a positively-charged lysine residue (K532) is located at the site homologous to Kvl .5 R487. The close resemblance of  these two channels, at least around the pore mouth, suggests that Kvl.4 might behave like Kvl.5. Indeed, as with Kvl.5, mutating H508 to glutamine (H508Q) or K532 to tyrosine (K532Y) antagonizes the H+-induced current inhibition (Claydon et al., 2000). However, the effects  of  H+ in Kvl.4 differ  from  those in Kvl .5 in that the current inhibition was frequency  dependent, arising in large part because of  a slowing of  recovery from  inactivation (Claydon et al., 2000). These results suggested, in contrast to those in Kvl .5, that activation of  the channel was required for  the current inhibition by H+0. Furthermore, in the N-terminal deleted Kvl.4 mutant, lowering external H+ accelerated slow inactivation without causing a significant  current inhibition (Claydon et al., 2002), and when either the H508Q, H508C, H508E, or K532Q mutation was introduced into the N-terminal deleted mutants, the time course of  slow inactivation became insensitive to external pH between pH 8.5 and 6.5. These results suggested that even though H508 could act as a pH sensor and K532 was probably a component of  the effector  mechanism, binding of  H+ to H508 did not result in an closed-inactivation dependent current inhibition, at least at a pFl > 6.5. It is uncertain whether reducing extracellular pH further  (e.g.  5.5) will induce any changes in the inactivation kinetics as in Kvl .5 or ShakerlR.  Together, the H+-induced current inhibition in Kvl.4 was mechanistically very different  from  that in Kvl.5, and the molecular basis for  these surprising differences  remains unknown. 6.3.3 External H+ modulation of  KCNQ2/KCNQ3 The heteromeric KCNQ2/KCNQ3 channel is a voltage-gated channel that is proposed to be the molecular correlate of  the neuronal M-current (Wang et al., 1998). It does not show slow inactivation, but external FT inhibits the channel with apK H  of  6.68 (Prole et al., 2003), and this inhibition showed some similarities to that of  Kvl.5. First, increasing external K+ antagonizes the H+-induced current inhibition. Second, the single channel conductance is unaltered by changing external pH. These results are consistent with FT modulating a gating process rather than directly occluding the pore. However, the gating changes in KCNQ2/KCNQ3 were due to a decrease of  the mean open time and the presence of  a long-lived closed state, in contrast to the unaltered intraburst behaviour observed in Kvl .5 (Figure 4.6). Interestingly, when the basic lysine residue at position 260 in the turret of  KCNQ3 was mutated to glutamine (K260Q), the current inhibition was enhanced. A similar result was also observed with the KCNQ2 H260Q (homologous to Kvl .5 H463Q) mutation. Together, these results are consistent with H+ modulating an outer pore process that makes the channel non-conducting; however, whether this process is similar to the slow (P/C-type) inactivation in other Kv channels is uncertain, if  not unlikely, given that this channel does not normally show slow inactivation. Nevertheless, an involvement of  the turret in the pH effect  is clearly illustrated. In summary, external IT can inhibit many Kv channels, and also affect  the rate of  slow inactivation through its interaction with the turret and/or the outer pore. While the mechanism for  the current inhibition may be different  in some of  these channels compared to that proposed ( for  Kvl.5, it nevertheless underscores a role for  the turret in the regulation of  channel function. 6.4 Physiological significance  and future  directions External FT, Zn2+, Ni2+, and some other divalent cations have been shown to inhibit macroscopic Kvl.5 current by binding to the histidine residue (H463) in the turret region. This current reduction, at least that caused by external FT, has a number of  implications for  cardiac pathophysiology. During ischemic heart attack or acidosis, extracellular pH may drop to a value as low as 6.0 (Rehncrona, 1985). At this pH, Kvl .5, which mediates the ultra-rapid delayed rectifier  current (IKur) in atrial myocytes (Feng et al., 1997; Feng et al., 1998) that is responsible for  repolarizing the action potential (Snyders, 1999), is inhibited. If  no other channels or transporters are being affected,  a direct consequence of  acidosis would be a reduction in the total repolarizing current and a prolongation the action potential, which may decrease the risk of  atrial fibrillation  (Eun et al., 2005). However, during ischemic heart attack, K+ may accumulate in the extracellular fluid,  and this elevated external K+ would be expected to antagonize the HMnduced current inhibition of  Kvl .5, diminishing the anti-atrial fibrillation  effect  induced by H+0. As suggested by Trapani and Korn (2003), even with a small change in pH, Kvl.5 currents may also be reduced by H+0 because of  a rightward gating shift  of  the g-V  curve. Of  course, with the presence of  other channels and transporters in the native tissue, the interplay between the membrane currents makes a prediction of  the functional  outcome more complicated. Given the important role of  Kvl.5 in atrial repolarization, therapeutic agents targeting Kvl.5 for  treating atrial arrhythmia are currently in development (Brendel and Peukert, 2003; Peukert et al., 2003). In light of  this, the results presented here have provided one possible mechanism for  modulating Kvl .5, in which H+ has been proposed to shift  the equilibrium from  a normal mode of  gating towards an "unavailable" mode of  gating that may relate to a slow inactivation process occurring from  closed states. This mechanism for  modulating Kvl .5 availability may be exploited pharmacologically in the rational design of  drugs. Several new questions arise from  this report. The inactivation process is suggested to be similar to the depolarization-induced inactivation; however, whether they reflect  the same conformational  changes in the outer pore and/or selectivity filter  is still unclear. If  mode U gating involves conformational  changes similar to those occurring in slow (P/C-type) inactivation, the outer pore would be expected to be constricted most of  the time at low pH (P 0 < 0.05 at pH 5.9 with 0 mM K+0). This hypothesis could be tested either by voltage clamp \ fluorimetry  with rapid perfusion  to determine if  a movement of  the turret and/or the outer pore occurs when pH0 is lowered. It can also be tested by "trapping" experiments at low pH using Cd2+ with cysteine mutants (Yellen et al., 1994) or with Ba2+ (Harris et al., 1998). For example, external Cd2+ has a higher affinity  for  the Shaker  T449C mutant when the channel-is in an inactivated state. If  a similar observation can be made in Kvl.5 with the R487C mutant, perfusion  with solution at pH 5.9 in 0 mM K+0 and suitable concentration of  Cd2+ should stabilize most channels in the inactivated state, such that even when the bath solution is switched back to the control solution (pH 7.4 with 0 mM K+0 and 0 mM Cd2+), the current amplitude would still be very small. Conversely, if  such a constriction of  the outer pore is absent during mode U  gating, the current amplitude would remain unchanged before  and after  the Cd2+ treatment. In this chapter, the voltage sensor movement is proposed to be uncoupled from  the inactivation machinery during closed-state inactivation, which predicts the activation gate may be open despite the closing of  the inactivation gate. This hypothesis may be tested using electron pair resonance (EPR) similar to the shown in KcsA (Cordero-Morales et al., 2006), in which the relative distance between the labelled residues can be determined. Alternatively, channels may be reconstituted in lipid bilayer such that conditions on both sides of  the membrane can be varied, and an open channel blocker (e.g.  internal TEA+) can be tested for  trapping in the central cavity at low pH with short depolarizing pulses. Similarly, auxiliary P subunit with N-terminal ball (e.g.  Kv pi. l) can also be used to determine the accessibility of  the N-terminal ball to the central cavity by comparing the recovery rate after  switching from  low pH to pH 7.4. However, this last experiment may be complicated by interactions between the N-terminal ball and slow inactivation. Slow inactivation is an important biophysical property of  Kv channels. By regulating the total repolarizing current through a reduction in the number of  channels available for  activation, slow inactivation can play a role not only in neuronal action potentials but also in cardiac action potentials which last for  several hundreds of  milliseconds. This dissertation has provided a number of  findings  regarding the modulation of  slow inactivation, and these results can lead to a better understanding of  the molecular mechanisms underlying this intrinsic process of  Kv and other voltage-gated ion channels. 6.5 References Aiyar, J., J. M. Withka, J. P. Rizzi, D. H. Singleton, G. C. Andrews, W. Lin, J. Boyd, D. C. Hanson, M. Simon, and B. Dethlefs.  1995. Topology of  the pore-region of  a K+ channel revealed by the NMR-derived structures of  scorpion toxins. Neuron 15:1169-1181. Aldrich, R. W., D. P. Corey, and C. F. Stevens. 1983. A reinterpretation of  mammalian sodium channel gating based on single channel recording. Nature 306:436-441. Baukrowitz, T. and G. Yellen. 1996. Two functionally  distinct subsites for  the binding of internal blockers to the pore of  voltage-activated K+ channels. Proc. Natl. Acad. Sci. U. S. A. 93:13357-13361. Brendel, J. and S. Peukert. 2003. Blockers of  the Kvl .5 channel for  the treatment of  atrial arrhythmias. Curr. Med Chem. Cardiovasc. Hematol. Agents 1:273-287. Chen, F. S., D. Steele, and D. Fedida. 1997. Allosteric effects  of  permeating cations on gating currents during K+ channel deactivation. J. Gen. Physiol. 110:87-100. Claydon, T. W., M. R. Boyett, A. Sivaprasadarao, K. Ishii, J. M. Owen, H. A. O'Beirne, R. Leach, K. Komukai, and C. H. Orchard. 2000. Inhibition of  the K+ channel Kvl.4 by acidosis: protonation of  an extracellular histidine slows the recovery from  N-type inactivation. J. Physiol. 526:253-264. Claydon, T. W., M. R. Boyett, A. Sivaprasadarao, and C. H. Orchard. 2002. Two pore residues mediate acidosis-induced enhancement of  C-type inactivation of  the Kvl.4 K+ channel. Am. J. Physiol. 283:C1114-C1121. Cordero-Morales, J. F., L. G. Cuello, Y. Zhao, V. Jogini, D. M. Cortes, B. Roux, and E. Perozo. 2006. Molecular determinants of  gating at the potassium-channel selectivity filter.  Nat. Struct. Mol. Biol. 13:311-318. De Biasi, M., H. A. Hartmann, J. A. Drewe, M. Taglialatela, A. M. Brown, and G. E. Kirsch. 1993. Inactivation determined by a single site in K+ pores. Pflugers  Arch. 422:354-363. Doyle, D. A., C. J. Morais, R. A. Pfiietzner,  A. Kuo, J. M. Gulbis, S. L. Cohen, B. T. Chait, and R. MacKinnon. 1998. The structure of  the potassium channel: molecular basis of  K+ conduction and selectivity. Science 280:69-77. Eun, J. S., J. A. Park, B. H. Choi, S. K. Cho, D. K. Kim, and Y. G. Kwak. 2005. Effects  of oxypeucedanin on /zKvl.5 and action potential duration. Biol. Pharm. Bull. 28:657-660. Feng, J., B. Wible, G. R. Li, Z. Wang, and S. Nattel. 1997. Antisense oligodeoxynucleotides directed against Kvl.5 mRNA specifically  inhibit ultrarapid delayed rectifier  K+ current in cultured adult human atrial myocytes. Circ. Res. 80:572-579. Feng, J., D. Xu, Z. Wang, and S. Nattel. 1998. Ultrarapid delayed rectifier  current inactivation in human atrial myocytes: properties and consequences. Am. J. Physiol. 275:H1717-H1725. Harris, R. E., H. P. Larsson, and E. Y. Isacoff.  1998. A permanent ion binding site located between two gates of  the Shaker  K+ channel. Biophys. J. 74:1808-1820. Holmgren, M., M. E. Jurman, and G. Yellen. 1996. N-type inactivation and the S4-S5 region of the Shaker  K+ channel. J. Gen. Physiol. 108:195-206. Hoshi, T., W. N. Zagotta, and R. W. Aldrich. 1994. Shaker  potassium channel gating. I: Transitions near the open state. J. Gen. Physiol. 103:249-278. Ishii, K., K. Nunoki, T. Yamagishi, H. Okada, and N. Taira. 2001. Differential  sensitivity of Kvl .4, Kvl .2, and their tandem channel to acidic pH: involvement of  a histidine residue in high sensitivity to acidic pH. J. Pharmacol. Exp. Ther. 296:405-411. Klemic, K. G., G. E. Kirsch, and S. W. Jones. 2001. U-type inactivation of  Kv3.1 and Shaker potassium channels. Biophys. J. 81:814-826. Klemic, K. G., C. C. Shieh, G. E. Kirsch, and S. W. Jones. 1998. Inactivation of  Kv2.1 potassium channels. Biophys. J. 74:1779-1789. Kurata, H. T., K. W. Doerksen, J. R. Eldstrom, S. Rezazadeh, and D. Fedida. 2005. Separation of P/C- and U-type inactivation pathways in Kvl.5 potassium channels. J. Physiol. 568:31-46. Kurata, H. T., G. S. Soon, J. R. Eldstrom, G. W. Lu, D. F. Steele, and D. Fedida. 2002. Amino- -terminal determinants of  U-type inactivation of  voltage-gated K+ channels. J. Biol. Chem. 277:29045-29053. Liu, Y., M. E. Jurman, and G. Yellen. 1996. Dynamic rearrangement of  the outer mouth of  a K+ channel during gating. Neuron 16:859-867. Loots, E. and E. Y. Isacoff.  2000. Molecular coupling of  S4 to a K+ channel's slow inactivation gate. J. Gen. Physiol. 116:623-635. Lopez-Barneo, J., T. Hoshi, S. H. Heinemann, and R. W. Aldrich. 1993. Effects  of  external cations and mutations in the pore region on C-type inactivation of  Shaker  potassium channels. Receptors. 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Mechanisms underlying modulation of neuronal KCNQ2/KCNQ3 potassium channels by extracellular protons. J. Gen. Physiol. , 122:775-793. Rehncrona, S. 1985. Brain acidosis. Ann. Emerg. Med. 14:770-776. Snyders, D. J. 1999. Structure and function  of  cardiac potassium channels. Cardiovasc. Res. 42:377-390. Starkus, J. G., Z. Varga, R. Schonherr, and S. H. Heinemann. 2003. Mechanisms of  the inhibition of  Shaker  potassium channels by protons. Pflugers  Arch. 447:44-54. Steidl, J. V. and A. J. Yool. 1999. Differential  sensitivity of  voltage-gated potassium channels Kvl.5 and Kvl.2 to acidic pH and molecular identification  of  pH sensor. Mol. Pharmacol. 55:812-820. Teisseyre, A. and J. W. Mozrzymas. 2006. Influence  of  extracellular pH on the modulatory effect of  zinc ions on Kvl.3 potassium channels. J. Physiol. Pharmacol. 57:131-147. Trapani, J. G. and S. J. Korn. 2003. Effect  of  external pH on activation of  the Kvl.5 potassium channel. Biophys. J. 84:195-204. Wang, H. S., Z. 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