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The contributions of Cav2.1 alternative splicing and calcium-dependent modulation to congenital migraine Adams, Paul Jacob 2010

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   THE CONTRIBUTIONS OF Cav2.1 ALTERNATIVE SPLICING AND CALCIUM-DEPENDENT MODULATION TO CONGENITAL MIGRAINE  by  Paul Jacob Adams  BSc, University of British Columbia, 2001    A THESIS SUMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY  in  The Faculty of Graduate Studies  (Neuroscience)    THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)  August 2010    © Paul Jacob Adams, 2010       ABSTRACT Cav2.1 calcium (Ca2+) channels are expressed throughout the mammalian central nervous system where they mediate P/Q-type Ca2+ currents essential for neurotransmitter release at most fast synapses. In humans, naturally occurring mutations in the CACNA1A gene encoding Cav2.1 are associated with several severe congenital disorders including familial hemiplegic migraine type 1 (FHM-1). Alternative splicing of the Cav2.1 transcript generates multiple functionally distinct channel variants with unique spatial and temporal expression patterns.  Yet, whether different Cav2.1 splice variants have distinct responses to FHM-1 missense mutations that relate to the localized, episodic nature of the FHM-1 phenotype has not been explored.  Using recombinant Cav2.1 channels, we systematically compared the biophysical effects of three FHM-1 mutations in two prevalent Cav2.1 splice variants.  All three FHM-1 mutations caused differential effects on voltage-dependent and kinetic properties when expressed in the short carboxyl terminus variant (Cav2.1 Δ47) compared to the long variant (Cav2.1 +47). Our findings provide important insight concerning the role of Cav2.1 alternative splicing and the pathophysiology of FHM-1. Ca2+-dependent facilitation (CDF) of Cav2.1 channels is a powerful means of channel control proposed to play a role in short-term facilitation of synaptic release during repetitive action potentials (APs).  However, empirical evidence to support CDF of Cav2.1 as a relevant mechanism of synaptic facilitation in the CNS is limited.  As such, short-term facilitation of synaptic release is generally attributed to enhanced vesicle release due to residual Ca2+ binding to sensor proteins that directly mediate vesicle fusion and transmitter release.  However, we found that two FHM-1 mutations occluded CDF of Cav2.1 in both recombinant and native systems and cause a corresponding attenuation in short-term synaptic facilitation at the cerebellar parallel fibre to Purkinje synapse.  This is the first evidence that presynaptic Ca2+ at this fast central synapse also enhances Ca2+ influx through Cav2.1 by means of CDF and acts as an additional required mechanism for short-term plasticity.  Thus, the data supports the notion that CDF of Cav2.1 underlies key aspects of short-term plasticity in the CNS and provides the first evidence that FHM-1 mutations directly affect Cav2.1 CDF.  ii TABLE OF CONTENTS  Abstract.................................................................................................................. ii Table of Contents ................................................................................................. iii List of Tables ........................................................................................................ vi List of Figures...................................................................................................... vii List of Abbreviations ........................................................................................... ix Acknowledgements ............................................................................................. xii Dedication ...........................................................................................................xiii Co-Authorship Statement ................................................................................. xiv 1 Introduction........................................................................................................ 1  1.1 Overview of voltage-gated calcium channels...................................................................1 1.1.1 Calcium channel discovery..................................................................................................... 1 1.1.2 There are multiple types of VGCCs in neurons...................................................................... 2 1.1.3 Current understanding of VGCC structure, classification and nomenclature......................... 4 1.1.4 Expression, properties and modulation of VGCCs................................................................. 9 1.2 The Cav2.1 VGCC............................................................................................................14 1.3 Alternative splicing ..........................................................................................................17 1.3.1 Alternative splicing overview for VGCCs............................................................................ 17 1.3.2 Mechanisms of alternative splicing ...................................................................................... 18 1.3.3 Alternative splicing of the Cav2.1 VGCC............................................................................. 20 1.4 Cav2.1 Ca2+ channelopathies ...........................................................................................26 1.4.1 General Cav2.1 channelopathies ........................................................................................... 26 1.4.2 Introduction to FHM............................................................................................................. 31 1.4.3 Mechanisms of migraine pathophysiology ........................................................................... 32 1.4.4 The role of Cav2.1 channels in molecular mechanisms of FHM-1....................................... 35 1.4.5 A key area in future FHM-1 research ................................................................................... 37 1.5 Ca2+-dependent modulation ............................................................................................39 1.5.1 Introduction to Ca2+-dependent modulation of VGCCs ....................................................... 39 1.5.2 CDF and CDI of Cav2.1 channels......................................................................................... 40 1.5.3 Ca2+-dependent modulation of Cav2.1 channels in synaptic plasticity ................................. 43 1.5.4 Key areas concerning future research on Cav2.1 modulation and synaptic plasticity .......... 45 1.6 Cav2.1 and synaptic plasticity at the PF-PC synapse ...................................................46 1.6.1 Brief cerebellum overview ................................................................................................... 46 1.6.2 Synaptic plasticity at the PF-PC synapse: a role for CDF of Cav2.1? .................................. 49  iii 1.7 Thesis hypotheses and objectives....................................................................................50 1.7.1 Hypothesis 1 ......................................................................................................................... 50 1.7.2 Hypothesis 2 ......................................................................................................................... 51 1.7.3 Thesis objectives................................................................................................................... 51 1.8 Bibliography .....................................................................................................................53 2 Cav2.1 Ca2+ Channel Alternative Splicing Affects the Functional Impact of Familial Hemiplegic Migraine Mutations: Implications for Ca2+ Channelopathies.................................................................................................. 73  2.1 Introduction......................................................................................................................73 2.2 Results ...............................................................................................................................76 2.2.1 Cav2.1 (+47) and Cav2.1 (∆47) variants are expressed in human cortex .............................. 76 2.2.2 FHM-1 mutations exhibit differential effects on the voltage-dependent properties of Cav2.1 splice variants..................................................................................................................... 79 2.2.3 FHM-1 mutations exhibit differential effects on recovery from inactivation of Cav2.1 splice variants ................................................................................................................................ 82 2.2.4 FHM-1 mutations exhibit differential effects on inactivation of Cav2.1 splice variants during tonic depolarization ............................................................................................................ 86 2.2.5 FHM-1 mutations exhibit differential effects on inactivation of Cav2.1 splice variants during bursts of depolarization ...................................................................................................... 90 2.3 Discussion..........................................................................................................................94 2.3.1 FHM-1 mutations differentially affect biophysical properties of Cav2.1 splice variants ..... 94 2.3.2 Differential effects of FHM-1 mutations on Cav2.1 splice variants may contribute to localized phenotype ....................................................................................................................... 95 2.3.3 Differential effects of FHM-1 mutations on Cav2.1 splice variants under different conditions may contribute to episodic nature of the phenotype .................................................... 96 2.3.4 The differential effects of mutations on Ca2+ channel function is likely multifaceted and important in all Ca2+ channelopathies ..................................................................................... 97 2.4 Experimental Procedures................................................................................................98 2.4.1 Site-directed mutagenesis ..................................................................................................... 98 2.4.2 Cell culture and transfection................................................................................................. 98 2.4.3 Electrophysiological recordings ........................................................................................... 99 2.4.4 Recording protocols and data analysis................................................................................ 100 2.4.5 RT-PCR of Cav2.1 carboxyl-terminal region from human cortex RNA............................. 101 2.5 Acknowledgements ........................................................................................................102 2.6 Bibliography ...................................................................................................................103 3 Contribution of Ca2+-Dependent Facilitation to Short-Term Synaptic Plasticity Revealed by Congenital Migraine Mutations in the Cav2.1 Ca2+ Channel .............................................................................................................. 107     3.1 Introduction....................................................................................................................107 3.2 Results .............................................................................................................................108 3.2.1 FHM-1 mutations occlude CDF and CDI of recombinant human Cav2.1 channels ........... 108 3.2.2 FHM-1 mutations occlude CDF of native P/Q-type currents in cerebellar PCs................. 114  iv 3.2.3 FHM-1 mutations occlude short-term synaptic facilitation at the PF-PC synapse ............. 117 3.2.4 FHM-1 mutant Cav2.1 channels appear to be in a basally facilitated state in PF boutons ................................................................................................................................... 122 3.3 Discussion........................................................................................................................126 3.3.1 A role for Cav2.1 CDF in short-term synaptic plasticity at the PF-PC synapse ................. 126 3.3.2 Facilitated Cav2.1 channels may increase susceptibility to CSD and aura ......................... 126 3.3.3 Impairment of synaptic efficacy may underlie cerebellar ataxia in FHM-1....................... 127 3.3.4 Beyond the PF-PC synapse................................................................................................. 128 3.4 Experimental procedures ..............................................................................................128 3.4.1 Cell culture and transfection............................................................................................... 128 3.4.2 Dissociated PCs .................................................................................................................. 129 3.4.3 Electrophysiological recording conditions in HEK and dissociated PCs ........................... 129 3.4.4 Protocols and data analysis for CDF and CDI.................................................................... 130 3.4.5 Cerebellar slices.................................................................................................................. 131 3.4.6 Extracellular field recordings ............................................................................................. 131 3.4.7 Two-photon microscopy: Ca2+ Imaging ............................................................................. 132 3.5 Acknowledgements ........................................................................................................133 3.6 Bibliography ...................................................................................................................134 4 Discussion........................................................................................................ 138  4.1 Overall significance and strengths ...............................................................................138 4.1.1 FHM-1 mutations differentially affect recombinant Cav2.1 splice variants ....................... 138 4.1.2 FHM-1 mutations occlude Ca2+-dependent modulation of Cav2.1 and synaptic Plasticity ...................................................................................................................................... 139 4.2 FHM-1 mutations differentially affect Cav2.1 splice variants: a role in the localized, episodic nature of the FHM-1 phenotype ...................................................141 4.2.1 Working hypothesis ............................................................................................................ 141 4.2.2 Possible limitations............................................................................................................. 145 4.2.3 Future directions ................................................................................................................. 146 4.3 A role for Cav2.1 CDF in synaptic plasticity ...............................................................147 4.3.1 Working hypothesis ............................................................................................................ 147 4.3.2 Possible limitations............................................................................................................. 153 4.3.3 Future directions ................................................................................................................. 156 4.4 Conclusions.....................................................................................................................157 4.4.1 General Conclusions........................................................................................................... 157 4.4.2 Relevance to treatment of human disease........................................................................... 158 4.5 Bibliography ...................................................................................................................160  Appendix 1: Ca2+ Channelopathies: Voltage-Gated Ca2+ Channels............ 166  Appendix 2: Other PhD Publications ............................................................. 203  Appendix 3: UBC Research Certificates of Approval................................... 204  v LIST OF TABLES  Table 2.1 - Mean values for voltage-dependent activation and inactivation parameters ..............................................................................................................................81  Table 2.2 - Time constant values and recovery from inactivation ....................................85                       vi LIST OF FIGURES  Figure 1.1 - HVA VGCCs........................................................................................................6 Figure 1.2 - Phylogeny of VGCCs ..........................................................................................8 Figure 1.3 - Alternative splicing in VGCCs.........................................................................19 Figure 1.4 - Human Cav2.1 splice-variants..........................................................................22 Figure 1.5 - Cav2.1 mutations implicated in EA2 and Epileptic and Ataxic Mouse Models .....................................................................................................................................28  Figure 1.6 - Pathophysiological mechanisms of migraine with aura.................................34 Figure 1.7 - Missense mutations in Cav2.1 associated with FHM-1 ..................................36 Figure 1.8 - CDF and CDI of Cav2.1 channels ....................................................................42 Figure 1.9 - Main microcircuitry in the cerebellar cortex .................................................48 Figure 2.1 - Human Cav2.1 Ca2+ channel topology and splice-variant expression in human cortex..........................................................................................................................78  Figure 2.2 - FHM-1 mutations differentially affect voltage-dependent properties of Cav2.1 (Δ47) and Cav2.1 (+47) variants ...........................................................................80  Figure 2.3 - Wild-type and FHM-1 mutant Cav2.1 (Δ47) and Cav2.1 (+47) variants exhibit different rates of recovery from inactivation..........................................................84  Figure 2.4 - Wild-type and FHM-1 mutant Cav2.1 (Δ47) and Cav2.1 (+47) variants exhibit different current decay during 25 Hz tonic depolarizations .................................88  Figure 2.5 - Wild-type and FHM-1 mutant Cav2.1 (Δ47) and Cav2.1 (+47) variants exhibit different current decay during bursts of depolarizations .....................................92  Figure 3.1 - The R192Q and S218L FHM-1 mutations occlude Ca2+-dependent modulation of human recombinant Cav2.1 channels........................................................110  Figure 3.2 - Measuring CDF using 100 Hz APW in Ca2+ and Ba2+.................................113 Figure 3.3 - CDF is similarly occluded in endogenous P/Q-type currents in acutely dissociated PCs from Cav2.1 R192Q and S218L knock-in mice......................................115  Figure 3.4 - Occlusion of Cav2.1 CDF results in a comparable attenuation in PPF at the PF-PC synapse......................................................................................................................118    vii Figure 3.5 - Kinetics of macroscopic currents of mutant channels is consistent with a preferentially facilitated gating mode.....................................................................120  Figure 3.6 - The kinetics of Ca2+ influx in S218L presynaptic terminals indicate facilitated Cav2.1 channels ..................................................................................................123  Figure 3.7 - Ca2+ transients in wild-type and S218L mice are identical in the presence of 500 nM ω-Aga-IVA .........................................................................................125  Figure 4.1 - Schematic model outlining the role of CDF of Cav2.1 channels as a key component of short-term synaptic facilitation ........................................................151  Figure 4.2 - R192Q and S218L FHM-1 mutant effects on Cav2.1 CDF are splice-variant dependent .....................................................................................................153                     viii LIST OF ABBREVIATIONS  Ca2+:  calcium ions Na+:  sodium ions APs:  action potentials Sr2+:  strontium ions Ba2+:  barium ions VGCC:  voltage-gated Ca2+ channel LVA:  low-voltage-activated HVA:  high-voltage-activated DHPs:  dihydropyridines N:  asparagine P:  proline K+:  potassium pS:  picosiemens EC:  excitation-contraction SA:  sinoatrial AV:  atrioventricular SNc:  substantia nigra pars compacta NMJ:  neuromuscular junction CNS:  central nervous system PNS:  peripheral nervous system SNARE:  soluble N-ethylmaleimide-sensitive factor attachment protein receptors VD:  voltage-dependent VI:  voltage-independent KO:  knock-out mRNA:  messenger RNA C-terminus:  carboxyl terminus  ix pre-mRNA:  precursor mRNA N-terminus:  amino terminus EA2:  Episodic Ataxia type 2 SCA6:  Spinocerebellar Ataxia type 6 FHM-1:  Familial Hemiplegic Migraine Type 1 LEMS:  Lambert-Eaton Myasthenic Syndrome tg:  Tottering tgla:  Leaner tgrol:  Rolling Nagoya HEK:  human embryonic kidney SCLC:  small-cell lung carcinoma CSD:  cortical spreading depression H+:  hydrogen ions NO:  nitric oxide TNC:  trigeminal nucleus caudalis VPM:  ventral posterior medial PAG:  periaqueductal gray region CaM:  calmodulin apoCaM:  Ca2+ free form of the CaM molecule IQ:  isolucine-glutamine CBD:  CaM binding domain CDF:  Ca2+-dependent facilitation CDI:  Ca2+-dependent inactivation NSCaTE:  N-terminal spatial Ca2+ transforming element Po:  open-channel probability APWs:  action potential waveforms CaS:  Ca2+ sensors  x SCG:  superior cervical ganglion EPSPs:  excitatory post-synaptic potentials PF:  parallel fibre PC:  Purkinje cell MFs:  mossy fibers GCs:  granule cells DCN:  deep cerebellar nuclei CFs:  climbing fibres LTP:  long-term potentiation LTD:  long-term depression EPSCs:  excitatory post-synaptic currents EGTA-AM:  ethylene glycol tetraacetic acid bound to acetoxymethyl ester τact:  kinetics of activation τinact:  kinetics of inactivation PPF:  paired-pulse facilitation PPR:  paired-pulse ratio fEPSPs:  field excitatory post-synaptic potentials nM:  nanomolar μM:  micromolar mM:  millimolar mV:  millivolt μs:  microseconds ms:  milliseconds s:  seconds mg:  milligrams μg:  micrograms   xi ACKNOWLEDGEMENTS I would like to thank my supervisor, Terry Snutch, for his patient guidance and encouragement throughout my PhD studies.  A special thanks to Esperanza Garcia for teaching me both the many techniques and theories of electrophysiology, but also for inspiring me to be excited about and enjoy science.  Many thanks to Sian Spacey for encouraging me to pursue research; I never would have made it this far without your support.  Thank you to my supervisory committee, Lynn Raymond, Steve Kehl, and Brian MacVicar, for their direction, advice, and feedback on this thesis. A special thanks to all of the past and present members of the Snutch lab.  The scientific collaborations and discussions were excellent, and the beach volleyball games, broomball games and drinks at the pub were absolutely essential.  Thank you to all members of the MacVicar lab, especially Ravi Rungta, for provided me with reagents, technical advice and performing the two-photon experiments found in Chapter 3.  A special thanks to Arn van den Maagdenberg for generating the R192Q and S218L knock-in mice, Simon Kaja for the initial set-up and care of the mice at UBC and Ray Gopaul for the continual care of the mice.  I am also very grateful to the Michael Smith Foundation for Health Research, the UBC graduate fellowship program and the William and Dorothy Gilbert Graduate Scholarship program for providing me with trainee funding during my graduate studies. I would like to thank all of my family and friends that helped and encouraged me to persevere through grad school.  A special thanks to my parents for always believing in me and teaching me the value of hard work, perseverance and integrity.  My highest appreciation is for my exceptional wife Ronee, for her unconditional support at every stage of this process.      xii DEDICATION                                  To Jesus,                   my beautiful wife Ronee, and my amazing children Katelynn and Jacob.                                              xiii CO-AUTHORSHIP STATEMENT I designed all experiments and analyzed all experimental results with input and feedback from other researchers.  I performed all experiments except for the two-photon microscopy experiments in chapter 3 which were performed by Ravi Rungta, and a few whole cell recordings of recombinant Cav2.1 channels in chapters 2 and 3 were performed by Esperanza Garcia and RT-PCR reactions in chapter 2 were performed by Laurence David.  The FHM-1 knock-in mice used in chapter 3 were kindly provided by Dr. Arn van den Maagdenberg.  I wrote the entire manuscripts for chapters 2 and 3, with subsequent editing from Terry Snutch and other authors.  I also wrote the entire review article shown in Appendix 1 excluding the introduction section, with subsequent editing by Terry Snutch.  Figures 1.6 and 4.1 in this thesis were created by Esperanza Garcia.   xiv 1 INTRODUCTION      1.1 Overview of voltage-gated calcium channels 1.1.1 Calcium channel discovery For some North Americans today, the year 1953 holds relevance in the inauguration of such things as Ian Fleming’s first James Bond novel Casino Royale, the release of Walt Disney’s Peter Pan, Hugh Hefner’s first issue of Playboy Magazine or perhaps the first successful climb to the summit of Mount Everest by Edmund Hillary and Tenzing Norgay.  Others may remember the significant political moments of 1953, such as the stroke and death of the Soviet Communist leader Joseph Stalin, the end of the Korean War, the coronation of Queen Elizabeth II, or the dawn of the Cold War following the announcement by both the United States and Russia that they had developed hydrogen bombs. Scientists, on the other hand, might be reminded of such epic discoveries as Jonas Salk’s polio vaccine (Salk, 1953), Eugene Aserinsky and Nathaniel Kleitman’s discovery of REM sleep (Aserinsky and Kleitman, 1953), or possibly the monumental publication of the double helix structure of DNA by Francis Crick and James Watson (Watson and Crick, 1953).  Yet for a small group of scientists, 1953 holds its significance in a classical discovery that would forever change research in the field of electrically excitable cells. In an article published in the Journal of Physiology in 1953, Paul Fatt and Bernard Katz made the unexpected finding that electrical responses persist in crustacean muscle fibres in sodium (Na+)-free media (Fatt and Katz, 1953).  This was an unexpected because up to that point the “Na+ theory” of electrical excitability of cells had prevailed.  The Na+ theory stated that action potentials (APs) arose as a result of regenerative processes depending on the permeability of cell surface membranes to Na+ (Hodgkin and Huxley, 1952).  Fatt and Katz were therefore cautious in concluding in their paper that “the mechanism of the action potential, and the species of ions involved in the movement of charge across the membrane, remain a puzzling problem…”, and in addition to other potential explanations, suggested “…influx of calcium or magnesium, or outflux of some internal anion may be responsible for  1 transport of charge”.  Their conclusions lead to further exploration in 1957 by Alan Hodgkin and Richard Keynes which showed an important role of calcium (Ca2+) in neurosecretion.  A year later, Paul Fatt and Bernard Ginsborg showed that full-blown APs in crustacean muscle can be generated in the absence of Na+ leading them to confidently conclude that the movement of divalents such as strontium (Sr2+), barium (Ba2+) or even Ca2+ could support APs across cell membranes (Fatt and Ginsborg, 1958).  Thus had begun a new pursuit for the identity of ions involved in the AP and neurosecretion (for full review of Ca2+ channel history see (Tsien and Barrett, 2005; Dolphin, 2006)).  1.1.2 There are multiple types of VGCCs in neurons By the end of the 1950’s, it was recognized that voltage-dependent Ca2+ conductances are an important component in excitable cells, and during the 1960’s Susumu Hagiwara verified the presence of Ca2+ conductances in numerous invertebrate tissues.  Reference to “a Ca2+ channel” in the singular form however, reveals the original perception that all measured Ca2+ currents were through a single type of voltage-gated Ca2+ channel (VGCC) and “…that Ca2+ channels everywhere are basically the same” (Brown et al., 1982).  The view of a single VGCC persisted into the early 1980s, despite the fact that as early as the mid 1970’s Hagiwara used voltage clamp experiments to show that in starfish eggs there are at least two distinct types of VGCCs which he termed type I and type II (Hagiwara et al., 1975). Subsequently, through efforts of a number of research groups including Llinas and Yarom, Carbone and Lux, Matteson and Armstrong and Bean, it was generally agreed that VGCCs are present in many systems and cell types and separable into two basic categories, the low voltage-activated (LVA) channels and high voltage-activated (HVA) channels (Carbone and Lux, 1984) depending upon the membrane potentials at which they first open.  LVA VGCCs open in response to small changes from the resting membrane potential, whereas HVA channels are activated by stronger depolarizations.  LVA channels have other distinguishing properties including a small single channel conductance (~ 5-12 picosiemens, pS), overlapping activation and inactivation ranges, rapid activation and inactivation kinetics, slow deactivation (closing) although a poorly defined pharmacology.  Contrastingly, HVA VGCCs generally  2 possess larger conductances (~ 15-25 pS), variable inactivation kinetics, faster deactivation, and a well- defined pharmacology (reviewed in (Snutch et al., 2005)). In 1985, classification of VGCCs was further refined.  Richard Tsien’s lab characterized properties of VGCCs in cell bodies of chick sensory neurons and found, based on single channel slope conductances and activation and inactivation properties, that VGCCs can be further classified into three categories.  Tsien’s lab termed the LVA channels T-type because they generate tiny unitary Ba2+ currents, give rise to a transient average current, have slow deactivation and resemble those previously described in heart.  They divided the HVA channels into either L-type or N-type.  L-type were named based on their large unitary conductance to Ba2+, long-lasting Ba2+ current and resemblance to those previously described in the heart.  The N-type were named based on the fact that they have similar characteristics to VGCCs previously described in neurons and because they have an intermediate conductance to Ba2+ and activate at higher voltages and thus are neither L- nor T-type. Identification and classification of VGCCs up to the mid 1980s had been primarily based on the biophysical properties of channels.  Starting in the mid 1980’s, however, Richard Tsien, Rodolfo Llinas, Michael Adams, Peter Hess and Bruce Bean began using VGCC pharmacological antagonists to further distinguish HVA VGCCs.  The first major class of antagonists utilized were the 1,4-dihydropyridines (DHPs) which specifically and potently block L-type VGCCs (Hess et al., 1984).  The peptide toxin ω- conotoxin-GVIA from marine snail venom was found to potently and specifically block N-type channels (McCleskey et al., 1987; Hirning et al., 1988; Plummer et al., 1989).  Use of these toxins by Llinas and collaborators led to the discovery of a non-inactivating, DHP and ω-conotoxin-GVIA-insensitive Ca2+ current in the cell bodies of cerebellar Purkinje neurons that is instead blocked by extract from venom of the American funnel-web spider.  They termed the newly identified channel P-type because of its dominance in Purkinje neurons (Llinas et al., 1989; Llinas et al., 1992).  In 1992 the lab of Michael Adams determined that the peptide ω-agatoxin IVA isolated from the venom of the American funnel web spider is the active component that blocks P-type Ca2+ currents (Mintz et al., 1992b).  3 In 1995, armed with channel specific blockers nimodipine (a DHP), ω-conotoxin GVIA and ω- agatoxin IVA, Andy Randell from the Tsien lab set out to identify the contribution of L-type, N-type and P-type currents, respectively, in rat cerebellar granule cells (Randall and Tsien, 1995).  In so doing, he identified two Ca2+ current types not previously identified in neurons but similar to VGCCs cloned from mammalian brain and characterized in exogenous mammalian expression systems by the labs of Shosaku Numa and Terry Snutch (cloning of VGCCs is discussed below).  The first of these, although resembling HVA P-type currents described in Purkinje neurons, has reduced sensitivity to ω-agatoxin IVA and faster inactivation kinetics and was thus thought to be distinct from the P-type channel and was termed the Q- type VGCC.  The second distinct current identified exhibited an activation threshold between HVA and LVA channels, a very rapid decay rate and insensitivity to high concentrations of nimodipine, ω- conotoxin GVIA and ω-agatoxin IVA.  Since this current was the residual current remaining after the application of the three known VGCC blockers it was designated the R-type VGCCs (Randall and Tsien, 1995). Thus, by the mid 1990’s it had been demonstrated by measuring Ca2+ currents in different tissues that VGCCs are separable into T-type, L-type, N-type, P-type, Q-type and R-type based on biophysical and pharmacological properties.  However, during the late 1980’s and early 1990s, identification of the protein composition and DNA sequences of distinct VGCCs greatly enhanced understanding of channel structure and lead to new ways of channel classification and naming.  1.1.3 Current understanding of VGCC structure, classification and nomenclature The protein composition of VGCCs was determined through work by Bill Catterall, Kevin Campbell, Franz Hoffman and others (Takahashi et al., 1987; Ellis et al., 1988; Ruth et al., 1989; Jay et al., 1990).  The DHP sensitive (L-type) VGCC complex was first purified from rabbit skeletal muscle and it was determined that this HVA VGCC is a large multimeric protein complex containing an equal stoichiometric ratio of a α1 pore-forming subunit and auxiliary subunits α2, δ, β and perhaps γ (Takahashi et al., 1987) (Fig. 1.1).  It is now clear that the α1 pore-forming subunit of all VGCCs is the largest  4 (ranging between 190-250 kDa) and contains most of the molecular components necessary to produce a functional channel, including the pore which allows ion permeation and selectivity, voltage sensors which allow the channel to respond to membrane depolarization, activation gate and the intrinsic inactivation machinery (Hille, 2001).  Functional differences in pharmacological sensitivities and basic biophysical properties such as ion conductance, threshold of activation and kinetics of inactivation between the VGCCs result primarily from variations in the primary structure of the channel.  The auxiliary subunits have important roles in channel surface expression and modulation.  The α2 and δ subunits are encoded by a single gene whose polypeptide product is post-translationally cleaved to yield an extracellular α2 product and a transmembrane δ product linked by disulfide bonds.  The α2δ subunit enhances channel expression (De Jongh et al., 1990; Jay et al., 1991).  β subunits bind the channel on the intracellular linker between domains II and III and enhance expression and modulate voltage and kinetic properties (Pragnell et al., 1994) (reviewed in (Dolphin, 2003a)).  Although the transmembrane γ subunit was originally shown to associate with the skeletal-muscle L-type VGCC, its necessity and functional role for this and other VGCCs remains uncertain (Freise et al., 2000).  More recently the γ subunit family has been shown rather to be a part of a group of transmembrane AMPA receptor regulatory proteins (Tomita et al., 2003) (for full review of Cav auxiliary subunits and their modulatory properties see (Dolphin, 2009)). Purification of LVA T-type channels has yet to be accomplished.  However, from molecular cloning it is known that mammalian T-type channels share only 20 to 30% amino acid sequence homology with mammalian HVA channels and that T-type channels lack entire structural motifs conserved within the HVA channels such as a β auxiliary subunit binding site in the I-II linker and an EF hand Ca2+ binding motif in the carboxyl terminus (C-terminus).  Thus, the association of typical HVA auxiliary subunits with native T-type channels is unlikely to have a significant role, although a minor role is possible (Dolphin et al., 1999; Dubel et al., 2004).    5  Figure 1.1: HVA VGCCs HVA VGCCs are large multimeric proteins containing a main α1 pore-forming subunit (blue) which includes four homologous domains (I-IV) each composed of six transmembrane segments and a pore forming loop between segments 5 and 6.  The 4th transmembrane segments have four regularly-spaced, positively charged amino acids (yellow +) which form the voltage sensor.  The N-terminus, C-terminus and inter-domain linkers are all on the cytoplasmic side of the membrane and are key sites for channel modulation.  Auxiliary subunits include the α2 and δ subunits (orange and light blue, respectively) (whose polypeptide product is post-translationally cleaved to yield an extracellular α2 product and a transmembrane δ product linked by disulfide bonds), β subunits (green), which bind the α1 pore-forming subunit on the intracellular linker between domains II and III and perhaps the transmembrane γ subunit (purple).  Knowledge of the individual subunits composing a VGCC opened the possibility for molecular cloning and reconstitution of recombinant VGCCs.  The first VGCC cloned was the DHP-sensitive L- type channel from skeletal and heart muscle by the lab of Shosaku Numa (Tanabe et al., 1987) followed by the demonstration that the mammalian brain expresses multiple distinct VGCCs by Terry Snutch (Snutch et al., 1990).  In 1991, a VGCC resembling the P-type channel was cloned from mammalian brain both by the labs of Snutch and Numa (Mori et al., 1991; Starr et al., 1991).  Within a year the  6 Snutch lab also cloned the neuronal N-type (Dubel et al., 1992) and that which was later called the R- type channel (Soong et al., 1993).  The last cloned VGCCs where the T-type channels by the lab of Ed Perez-Reyes (Cribbs et al., 1998; Perez-Reyes et al., 1998; Lee et al., 1999c) and the Snutch lab (McRory et al., 2001).   Cloning made it possible to express VGCCs in recombinant systems and to more thoroughly characterize their basic properties than was previously possible in native systems. In terms of VGCC discovery and classification, one of the most notable discoveries following the cloning of the channels was made by the Snutch lab in 1999.  They showed that P-type and Q-type currents are actually conducted through the same channel, rather than through distinct channels as originally believed.  The P- and Q-type currents result from alternative splicing of a single gene transcript in which the inclusion or exclusion of two residues (asparagine (N) and proline (P)) in the S3- S4 linker of domain IV determines the distinct pharmacological sensitivities, P-type (-NP; high ω- agatoxin IVA sensitivity) and Q-type (+NP; low ω-agatoxin IVA sensitivity) (Bourinet et al., 1999). From that point onward it has been generally agreed that P and Q-type currents are conducted through different splice variants of the same channel. Molecular cloning allowed researchers to identify the genes that encode for VGCCs and powerfully shaped the classification of channels.  Beyond the five categories of L-type, N-type, P/Q- type, R-type and T-type based on activation thresholds and pharmacological sensitivities of Ca2+ currents measured in native systems established by the end of the 1990s, it is now known that there are at least ten genes that encode distinct α1 pore-forming subunits in the VGCC family in mammals.  Sequence analysis of the ten genes has been used to create a systematic organization based on structural and evolutionary relationship.  Figure 1.2 shows the phylogeny of the ten mammalian VGCC α1 pore-forming subunits based on sequence similarities between conserved transmembrane and pore forming domains.  The channels are classified into three distinct branches: 1) L-type, 2) P/Q-type, N-type and R-type and 3) T- type.  The α1 subunits are more than 70% identical within a family, but less than 40% between families. All three families have a single representative in the C. elegans genome which suggests their divergence was phylogenetically ancient (Ertel et al., 2000).  7  Figure 1.2: Phylogeny of VGCCs Phylogenetic relationships between primary sequences of cloned mammalian VGCC α1 pore-forming subunits, including the membrane spanning segments and pore-forming loops.  Both the Cav and alphabetical (parentheses) nomenclatures are presented for each channel.  Gene names for each channel are shown in green.  Identification of VGCCs by different labs over the years led to numerous ways of naming the channels.  Initially an attempt was made to create a unifying naming system based on an alphabetical nomenclature, but this proved limited in its ability to reflect the structural homology within families and had the potential for overlap between names (Birnbaumer et al., 1994).  Nonetheless, the alphabetical nomenclature is sometimes still used today and is reflected in the gene names.  In 2000, a group of leaders in the field devised a systematic naming system of VGCCs based on the 10 α1 pore-forming subunits.  The naming system reflects the functional, structural and evolutionary relationships of the channels and is coordinated with nomenclature used for other voltage-gated channels such as potassium  8 (K+) and Na+ channels (Ertel et al., 2000).  The naming system adopted at that time was approved by the Nomenclature Committee of the International Union of Pharmacology and is the preferred use today. The system uses the symbol for the primary permeating ion (Ca) with the main physiological regulator (voltage) indicated as a subscript (Cav).  A numeric identifier between 1 and 3 is used to identify the α1 pore-forming subunit gene family (Cav1: L-type; Cav2: P/Q-type, N-type and R-type; Cav3: T-type), and a second numeric identifier after the decimal point to identify the order of discovery of the subunit within the family (figure 1.2).  It is now conventional to refer to each VGCC by the Cav nomenclature of the α1 pore-forming subunit and to refer to the currents elicited by the channel as L-type, P/Q-type N-type, R- type or T-type (Catterall et al., 2005).  This is the convention used in the remainder of this thesis. The auxiliary subunits often associated with VGCCs in mammals have a similar naming system that reflects the gene family and the specific splice-variant.  To date there have been four β subunit genes cloned (β1-4), four α2δ subunits (α2δ1-4), and seven γ subunits (γ2-8) (Ertel et al., 2000; Dolphin, 2009).  1.1.4 Expression, properties and modulation of VGCCs Over the past 57 years, the biophysical and modulatory properties of all ten VGCCs have been extensively explored using both recombinant channels in various expression systems and endogenous channels in many in vivo preparations.  The following section is a brief survey of the key properties and tissue expression of each of the ten VGCCs. The Cav1 family: Cav1 channels are the primary trigger for excitation-contraction (EC) coupling in cardiac, skeletal and smooth muscles (Bean, 1989).   They are also found in most central and peripheral neurons where they in part control Ca2+-dependent gene expression, as well as in endocrine cells and many types of non-excitable cells where they contribute to a variety of processes including exocytotic release.  The Cav1 channels conduct L-type Ca2+ currents and are potently blocked by DHPs (Glossmann and Striessnig, 1988; Xu and Lipscombe, 2001; Koschak et al., 2003; McRory et al., 2004), have large single  9 channel conductance in Ba2+ (25 pS), activate between -60 and -10 mV and have a slow rate of inactivation (τ > 500 ms) (Hille, 2001). Cav1.1 channels are important in striated muscle cells for coupling membrane depolarization to the release of Ca2+ from cytoplasmic stores and triggering EC coupling (Rios and Brum, 1987).  In this instance, depolarizing changes in membrane potential cause a conformational change in Cav1.1 and without a requirement for direct Ca2+ influx induce an allosteric interaction with the sarcoplasmic reticulum (SR) ryanodine receptor (RyR1) ultimately inducing Ca2+ release and muscle contraction (Tanabe et al., 1990; Rios et al., 1992; Flucher and Franzini-Armstrong, 1996; Kugler et al., 2004). Related to this role as a voltage sensor for the RyR1, several loss-of-function missense mutations in the CACNA1S gene encoding for Cav1.1 have been implicated in two muscle disorders, hypokalemic periodic paralysis and malignant hyperthermia susceptibility (reviewed in (Adams and Snutch, 2007), Appendix 1). The Cav1.2 channel is widely expressed in heart, brain (neurons, preferentially somatodendritic), smooth muscle, pituitary, gastrointestinal systems, lungs, immune system and testes (Ertel et al., 2000; Splawski et al., 2004).  These channels have prominent roles in EC-coupling in cardiac and smooth muscle and AP propagation in sinoatrial (SA) and atrioventricular (AV) node, hormone secretion in endocrine cells and synaptic plasticity in neurons (Hell et al., 1993; Bokvist et al., 1995; Striessnig, 1999; Catterall, 2000; Schulla et al., 2003; Sinnegger-Brauns et al., 2004).  Unlike EC coupling between Cav1.1 and RyR1 receptors in skeletal muscle, EC coupling in cardiac tissue requires Ca2+ influx through Cav1.2 in order to activate the RyR2 in the SR and release Ca2+ from internal stores to initiate muscle contraction (Meissner, 1994).  Two gain-of-function missense mutations in the sixth transmembrane segment of domain I of the Cav1.2 channel have recently been associated with a severe arrhythmic disorder, Timothy syndrome (reviewed in (Adams and Snutch, 2007), Appendix 1). Cav1.3 channels are the least sensitive to DHPs and are primarily found in photoreceptors, cochlear hair cells, endocrine cells (pancreatic cells, pituitary, adrenal chromaffin cells and pinealocytes), some in atrial muscle and SA and AV nodes in heart, vascular smooth muscle and neurons (preferentially  10 located on cell bodies and proximal dendrites) (Hell et al., 1993; Chik et al., 1997; Takimoto et al., 1997; Platzer et al., 2000; Garcia-Palomero et al., 2001; Mangoni et al., 2003; Michna et al., 2003).  Of the HVA channels, Cav1.3 channels activate at relatively negative membrane potentials and are often referred to as “low-voltage-activated” L-type channels.  The Cav1.3 channels play important physiological roles concerning neurotransmitter release in sensory cells and hormone secretion to affect mood behavior, the control of AV node conductance and cardiac rhythm, and also play a key role in the autonomous activity of adult substantia nigra pars compacta (SNc) dopaminergic neurons (Platzer et al., 2000; Mangoni et al., 2003; Sinnegger-Brauns et al., 2004; Chan et al., 2007). Cav1.4 channels have important roles in neurotransmitter release in retinal photoreceptors and bipolar cells as well as in spinal cord (Tachibana et al., 1993; Nachman-Clewner et al., 1999). Photoreceptor neurotransmission is atypical in that photoreceptor cells do not fire action potentials, but rather have continuous graded membrane potentials.  Also, photoreceptors are tonically depolarized in the absence of a light stimulus resulting in continuous glutamate release and subsequently hyperpolarize in response to light stimuli (Wu, 1994).  Considering their critical role in vision, it is perhaps not surprising that both loss-of-function and gain-of-function mutations in the CACNA1F gene encoding Cav1.4 in humans are implicated in the vision related diseases incomplete X-linked congenital stationary night blindness and X-linked cone-rod dystrophy (reviewed in (Adams and Snutch, 2007), Appendix 1). The Cav1.4 channels are also expressed in immune cells and involved in T-lymphocyte activation (Bech- Hansen et al., 1998; Strom et al., 1998; Naylor et al., 2000; Firth et al., 2001; Ball et al., 2002; McRory et al., 2004). The Cav2 family: The Cav2 family of channels is the primary conduit of Ca2+ entry at presynaptic terminals and required for neurotransmitter release in both the central nervous system (CNS) and peripheral nervous system (PNS).  Their single channel conductances in Ba2+ range between 10 and 20 pS, they can activate anywhere between -120 (mostly R-type) and -30 mV and they possess an intermediate inactivation rate (τ = 50-80 ms) (Catterall et al., 2005).  11 Cav2.1 channels conduct P/Q-type Ca2+ currents and are blocked by ω-agatoxin-IVA and ω- conotoxin MVIIC (Hillyard et al., 1992; Mintz et al., 1992a).  Cav2.1 channels are ubiquitously expressed in neurons throughout the CNS and PNS (presynaptic terminals, dendrites and some cell bodies) as well as in heart, pancreas (β-cells) and pituitary.  Cav2.1 channels play a primary role in neurotransmitter release in CNS and the neuromuscular junction (NMJ) and are known to contribute to excitation-secretion coupling in pancreas (Hillman et al., 1991; Mori et al., 1991; Starr et al., 1991; Mintz et al., 1992a; Ousley and Froehner, 1994; Randall and Tsien, 1995; Westenbroek et al., 1995; Day et al., 1997; Horvath et al., 1998; Ishikawa et al., 2005).  Considering the central role of Cav2.1 channels in neurotransmission, it is perhaps not surprising that mutations in the CACNA1A gene encoding Cav2.1 are associated with several severe human disorders (discussed in a subsequent section). Cav2.2 channels conduct N-type Ca2+ currents and are potently blocked by ω-conotoxin GVIA, ω-conotoxin MVIIA and MVIIC (McCleskey et al., 1987; Hillyard et al., 1992).  They are expressed in neurons (presynaptic terminals, dendrites and cell bodies) and contribute to neurotransmitter release in CNS and sympathetic neurons, the sympathetic regulation of the circulatory system and play an important role in sensation and transmission of pain (Westenbroek et al., 1992; Dunlap et al., 1995; Ino et al., 2001; Mori et al., 2002; Beuckmann et al., 2003). Cav2.3 channels conduct R-type Ca2+ currents, are resistant to block by ω-agatoxin-IVA, ω- conotoxin GVIA, ω-conotoxin MVIIA and MVIIC and under some conditions are blocked by SNX-482 (blocks recombinant channels, but is only partially effective at blocking native R-type currents) (Newcomb et al., 1998; Tottene et al., 2000).  Cav2.3 channels are primarily expressed in neurons (dendrites, cell bodies and some presynaptic terminals), heart, testes and pituitary where they have roles in neurotransmitter release, repetitive firing, long-term potentiation and post-tetanic potentiation (Randall and Tsien, 1995; Tottene et al., 1996; Dietrich et al., 2003; Jing et al., 2005; Pereverzev et al., 2005). The Cav3 family: The Cav3 family of channels conducts T-type Ca2+ currents.  These channels diverged early in VGCC evolution and differ significantly in primary sequence from HVA channels.  Although there is  12 some evidence for an interaction of Cav3 α1-subunits with VGCC auxiliary subunits (Dolphin et al., 1999; Dubel et al., 2004; Lin et al., 2008), recombinant Cav3 channels display properties similar to those of native T-type currents in the absence of auxiliary subunits and it is not yet clear what roles auxiliary subunits contribute towards Cav3 functional properties. Cav3 channels are blocked by the scorpion peptide kurtoxin from Parabuthus transvaalicus and Cav3.1 and 3.2 channels are blocked by the small organic antiepileptic ethosuximide (Chuang et al., 1998; Lee et al., 1999b; Gomora et al., 2001).  Cav3 channels have variable sensitivities to nickel (Ni2+), Cav3.2 is highly sensitive to Ni2+ with an IC50 of 12 μM compared to IC50’s of 250 μM and 216 μM for Cav3.1 and Cav3.3, respectively (Lee et al., 1999b).  Cav3 channels have relatively small single channel conductances (8-10pS), activate at membrane potentials between -90 and -60 mV and have fast inactivation time constants (τ=20-50 ms) (Hille, 2001).  The acute voltage sensitivity of Cav3 channels plays a pivotal role in regulating cellular excitability and oscillatory behaviours. Cav3.1 channels are predominantly expressed in neurons (soma and dendrites), heart (SA node), ovary and placenta, and have roles in burst firing associated with thalamic oscillations and cardiac pacemaking (Perez-Reyes et al., 1998; Craig et al., 1999; Talley et al., 1999; Monteil et al., 2000a; Perez-Reyes, 2003; Ono and Iijima, 2005).  In the cerebellum, mGluR1-mediated potentiation of Cav3.1 T-type currents may promote synapse-specific Ca2+ signalling in response to bursts of excitatory inputs (Hildebrand et al., 2009).   Cav3.2 channels are widely expressed in the juvenile and adult hippocampus, cerebellum, pons/medulla, striatum, thalamus/hypothalamus, olfactory bulb, heart (SA node), and cortex (Talley et al., 1999; McRory et al., 2001; Perez-Reyes, 2003; McKay et al., 2006).  They are also important for burst firing and oscillatory behavior and have key roles in smooth muscle contraction, proliferation, aldosterone secretion and cortisol secretion (Chuang et al., 1998; Cribbs et al., 1998; Lee et al., 1999b; Talley et al., 1999; Bohn et al., 2000; Gomora et al., 2001; Hansen et al., 2001; Schrier et al., 2001).  In the periphery, Cav3.2 channels are found in the primary nociceptor pathway and have been shown to contribute both to acute and chronic nociceptive behaviours (reviewed in (Snutch and David, 2006)).  In  13 the last few years, researchers have uncovered several point mutations in the CACNA1H gene encoding Cav3.2 in patients with idiopathic generalized epilepsy (IGE) and childhood absence epilepsy (CAE), some of which have gain-of-function effects on channel function (reviewed in (Adams and Snutch, 2007), Appendix 1). Cav3.3 channels are expressed primarily in neurons and have key roles in thalamic oscillations (Perez-Reyes et al., 1998; Talley et al., 1999; Monteil et al., 2000b; Perez-Reyes, 2003).  Whereas Cav3.1 and Cav3.2 have similar activation and inactivation kinetics, Cav3.3 activation and inactivation kinetics are much slower.  The Cav3.3 channels also have faster deactivation kinetics as well as more hyperpolarized voltage dependence of activation and inactivation compared with the other two Cav3 channels (McRory et al., 2001).  Gαq/11-coupled GPCRs exhibit a strong inhibitory effect on Cav3.3 T- type Ca2+ currents, but have either no effect on or increase Cav3.1 and Cav3.2 peak current amplitudes (Hildebrand et al., 2007).       1.2 The Cav2.1 VGCC The CACNA1A gene in mammals has 47 exons and encodes the 2,200-2,400 amino acid pore- forming α1 subunit (Cav2.1; α1A).  In exogenous systems, in order to recapitulate native P/Q-type currents Cav2.1 channels must be associated with both β and α2δ auxiliary subunits.  Endogenous Cav2.1 channels are likely associated with any one of four β subunits (β1-4) and any one of four α2δ subunits (α2δ1-4).  The different β subunits enhance trafficking of channels to the plasma membrane and can modulate activation threshold and kinetics of inactivation with varying degrees of effectiveness (Stea et al., 1994; Walker and De Waard, 1998) (reviewed in (Arikkath and Campbell, 2003; Richards et al., 2004; Dolphin, 2009)).  β subunits bind via their guanylate kinase-like domain to the α interaction domain within the I–II linker of Cav2.1 channels with different affinities (Pragnell et al., 1994; De Waard and Campbell, 1995).  The α2δ subunits promote trafficking of Cav2.1 channels by an unknown mechanism.   The α2δ subunit binds via a Von Willebrand factor-A domain (Canti et al., 2005; Davies et al., 2006) to one or more unidentified extracellular domains of the Cav2.1 channel.  Which of the various auxiliary subunits are complexed with  14 endogenous Cav2.1 channels in a particular cell type likely depends on both the expression of the subunits and their affinity for Cav2.1. Cav2.1 channels are generally concentrated to presynaptic terminals but are also detected on dendritic spines and shafts (Hillman et al., 1991; Westenbroek et al., 1995).  Cav2.1 (and Cav2.2 and Cav2.3 to a lesser degree) channels have been shown to be necessary for the release of neurotransmitters from presynaptic terminals at most fast synapses in spinal cord and brain stem, cerebellum, hippocampus, calyx of Held and cortex (Hillyard et al., 1992; Turner et al., 1992; Takahashi and Momiyama, 1993; Turner et al., 1993; Castillo et al., 1994; Regehr and Mintz, 1994; Wheeler et al., 1994; Regehr and Atluri, 1995).  Ca2+ influx through Cav2 channels initiates neurotransmitter release by triggering vesicle fusion with the plasma membrane by interacting with the SNARE (soluble N- ethylmaleimide-sensitive factor attachment protein receptors) protein complex composed of syntaxin, SNAP-25 and VAMP/synatobrevin (reviewed in (Catterall, 2000; Evans and Zamponi, 2006; Kisilevsky and Zamponi, 2008)).  The function of the SNARE complex in vesicle fusion is regulated by numerous proteins including the Ca2+-binding protein synaptotagmin which acts as a Ca2+ sensor.  It is believed that the physical interaction of the Cav2 channel with the SNARE complex and synaptotagmin enables a tight structural association and functional coupling of Ca2+ entry with vesicle fusion (Catterall, 2000; Mochida et al., 2003b).  Cav2.1 channels contain a specific synaptic protein-protein interaction region called the synprint site located within the intracellular loop linking domains II and III (Sheng et al., 1994).  The synprint site of Cav2.1 interacts with several proteins in the SNARE complex and can either function to modulate target proteins or receive modulation from other proteins (Sheng et al., 1994; Charvin et al., 1997; Sheng et al., 1997; Wiser et al., 1997). The regulation of Cav2.1 channels exerts a crucial influence on presynaptic Ca2+ influx and the precise tuning of neurotransmitter release.  In addition to modulation by SNARE proteins, Cav2.1 channels are also acutely regulated in a variety of ways by effector molecules acting via protein phosphorylation through second messenger-activated kinase pathways (reviewed in (Catterall, 2000; Snutch et al., 2005; Evans and Zamponi, 2006)).  For example, PKA increases Cav2.1 channel activity  15 indirectly by interfering with phosphatidylinositol-4,5-bisphosphate mediated regulation which normally produces voltage-dependent current inhibition (Fournier et al., 1993; Fukuda et al., 1996; Huang et al., 1998; Kaneko et al., 1998; Wu et al., 2002).  Also, activation of certain G-protein coupled receptors is a well known means of potent and fast inhibition of both Cav2.1 and Cav2.2 channels affecting both voltage-dependent (VD) and voltage-independent (VI) mechanisms (Ikeda and Schofield, 1989; Lipscombe et al., 1989; Mintz and Bean, 1993; Kisilevsky and Zamponi, 2008).  G-protein mediated VD inhibition is thought to involve a complex interplay between multiple cytoplasmic regions of Cav2.1 including the I-II linker and N-terminus (Dolphin, 2003b; Agler et al., 2005), and possibly the C- terminus, to enhance Gβγ binding affinity (Li et al., 2004).  Furthermore, modulation of Cav2.1 by Ca2+- calmodulin-dependent protein kinase II interacting with the C-terminus of the channel slows channel inactivation (Jiang et al., 2008).  Cav2.1 channel activity can also be indirectly regulated through phosphorylation of auxiliary subunits (reviewed in (Dolphin, 2009)). The ablation of P/Q-type currents by genetic deletion of Cav2.1 in mice (Cav2.1 knock-out; KO) results in numerous neurological dysfunctions including absence seizures characterized by 3-5 Hz cortical spike-wave discharges, ataxia and dystonia beginning around post-natal day 10 and progressing with age, and selective degeneration of the cerebellum (particularly the anterior vermis) in older mice.  If unaided, Cav2.1 homozygous KO mice typically do not survive past weaning (Jun et al., 1999; Fletcher et al., 2001; Song et al., 2004; Todorov et al., 2006).  Examination of pain-related behavioral responses of Cav2.1 KO mice also reveals that Cav2.1 plays a role in inhibiting pain signalling during non-injurious noxious thermal stimuli, although on the other hand is necessary for the conduction of pain signalling in inflammatory and neuropathic pain states (Luvisetto et al., 2006). The Cav2.1 properties and characteristics described to this point have largely been general. However, there are many variants of Cav2.1 that arise as a result of alternative splicing of the CACNA1A transcript and that produce unique channels with distinct spatial and temporal expression patterns, biophysical properties and pharmacological sensitivities (Bourinet et al., 1999; Soong et al., 2002; Timmermann et al., 2002; Chaudhuri et al., 2004; Chang et al., 2007).  Alternative splicing of Cav2.1 is a  16 critical means to achieve channel diversity relevant to a variety of functional roles.  The following sections explore alternative splicing and its relevance to Cav2.1 physiology and pathophysiology.       1.3 Alternative splicing 1.3.1 Alternative splicing overview for VGCCs In the early 1980s, researchers predicted that about 5% of genes in higher eukaryotes are alternatively spliced (Sharp, 1994).  Sequencing of the human genome, however, revealed only approximately 32,000 genes which is far less than the previously predicted 150,000 (Pennisi, 2000).  The disparity fueled a revolution in the minds of many scientists regarding biological complexity.  It is now believed that the number of human messenger RNA (mRNA) forms is much higher than the number of genes and that alternative splicing plays a central role in the production of biological complexity.  Recent estimates are that between 75 and 94% of all human genes produce primary transcripts which undergo alternative splicing to produce structurally and functionally distinct protein variants (Johnson et al., 2003; Wang et al., 2008). Alternative splicing of VGCCs is a critical component of their functional diversity.  As described, there have been ten genes identified encoding the Cavα1 subunits, yet this number is insufficient to account for the functional diversity amongst the native Ca2+ currents and their roles in Ca2+-dependent signalling in different brain regions and cellular compartments.  While there may only be ten Cav subunit genes, it is predicted that there are perhaps 1000 times as many splice variants, likely with their own unique functional properties and spatial and temporal expression patterns (Lipscombe et al., 2002; Vigues et al., 2002; Emerick et al., 2006; Gray et al., 2007). It has been shown for a variety of VGCCs that alternative splicing occurs within key functional domains of the channels.  For example, alternative splicing within the S3-S4 extracellular loops is known to occur in Cav2.1 (Bourinet et al., 1999), Cav2.2 (Lin et al., 1997; Stea et al., 1999) and Cav1 channels (Barry et al., 1995) and is thought to generally influence voltage sensitivity of the channel because of the  17 close proximity to the S4 voltage sensors which must move upon depolarization (Bezanilla, 2002).  The intracellular loop connecting homologous domains II and III (II-III linker) in VGCCs is a key site of channel modulation, important in membrane targeting and in some channels links to downstream effector proteins required for neurotransmitter release (Catterall, 2000).  Alternative splicing within the II-III linker has been reported for Cav2.2 (Pan and Lipscombe, 2000), Cav2.1 (Soong et al., 2002), Cav2.3 (Pereverzev et al., 2002) and Cav3 channels (Mittman et al., 1999a; Chemin et al., 2001).  Splicing in the II-III linker in Cav2.2 and Cav2.1 channels significantly affects channel targeting to the plasma membrane, causes differential interaction with β subunits and alters current-voltage relationships (Scott et al., 1996; Pan and Lipscombe, 2000; Lipscombe et al., 2002; Rajapaksha et al., 2008).  Another key region of alternative splicing is the C-terminus.  The C-terminus makes up about one-third of VGCCs and is prominent in determining functional diversity.  This region contains the sites for several regulatory elements such as the binding of Ca2+, calmodulin and G-proteins (Catterall, 2000) and is also important in channel targeting to the plasma membrane (Maximov and Bezprozvanny, 2002).  There are at least two C-terminus variants of different lengths for most VGCCs and the variants show distinct Ca2+- dependent modulation, G-protein modulation, changes in the voltage-dependence, kinetics of inactivation and current amplitude (Hell et al., 1994; Zhuchenko et al., 1997; Bourinet et al., 1999; Mittman et al., 1999a; Mittman et al., 1999b; Krovetz et al., 2000; Lu et al., 2001; Sandoz et al., 2001; Murbartian et al., 2002; Pereverzev et al., 2002; Soong et al., 2002).  1.3.2 Mechanisms of alternative splicing In the process of generating mature, stable mRNA that can be directly translated into protein, non-coding regions corresponding to introns must be removed from precursor mRNA (pre-mRNA) by the process of splicing.  A spliceosome protein complex recognizes introns by signature dinucleotide sequences within intron/exon boundaries (Sharp and Burge, 1997; Wu and Krainer, 1999; Modrek and Lee, 2002; Singh, 2002).  Splicing out sequences at any position in the pre-mRNA depends on the  18 combination of both the nucleotide sequences in key positions, as well as the spliceosome composition available within the cells (Grabowski and Black, 2001).  Figure 1.3: Alternative splicing in VGCCs There are six types of alternative splicing that occurs for VGCC pre-mRNA transcripts, A-F.  Exons are shown in dark purple, alternate exons in blue and segments of exons included or excluded as a result of alternative splice donor and acceptor sites are shown in light purple.  Small bent arrows indicate promoter start sites.  There are six types of alternative splicing thus far identified for VGCCs (Figure 1.3).  One described type is the inclusion or exclusion of an entire exon (Fig. 1.3A).  There are constitutively expressed exons that are essential for basic protein function and structure, and alternatively expressed exons that generally modify protein function and fine-tune the protein for specific cellular tasks.  The inclusion or exclusion of alternative exons in the final mRNA transcript is determined by a number of factors including tissue type, cell type, developmental stage, metabolic state of the cell and gender (Black, 1998; Grabowski, 1998; Xie and Black, 2001).  19 A second type of splicing observed for VGCCs is the mutual exclusion of a series of homologous exons whereby only one of the exons will be selected and included in the final mRNA transcript (Fig. 1.3B). A third and fourth form of alternative splicing occur at alternative 5’ donor and 3’ acceptor slice sites.  Splicing at an alternative intron sequence near the 5’ intron end can cause elongation or shortening of the preceding exon (alternative splice donor) (Fig. 1.3C), and splicing at an alternative intron sequence near the 3’ end can cause elongation or shortening of the following exon (alternative splice acceptor) (Fig. 1.3D). The remaining forms of splicing involve the use of alternative 5’ promoters (Fig. 1.3E) and 3’ polyadenylation/cleavage sites (Fig. 1.3F), respectively.  The use of alternative promoters may join additional exons to the 5’ end of the protein product, whereas the composition and length of 3’ untranslated regions can modify mRNA stability and alter targeting to specific regions of the cell (Modrek and Lee, 2002). An additional, albeit less common, means of channel variation arises through RNA editing which may include nucleoside modifications such as cytosine to uracil and adenosine to inosine deaminations, or non-templated nucleotide additions and insertions (Smith et al., 1998; Tsunemi et al., 2002) (for full review of VGCC alternative splicing see (Lipscombe and Castiglioni, 2004)).  1.3.3 Alternative splicing of the Cav2.1 VGCC Fitting with the diverse roles of Cav2.1 channels described in detail above, their functional properties can be fine-tuned on a short time scale by gating modulation through channel phosphorylation (Zamponi et al., 1997), G-protein interaction (Colecraft et al., 2001), and assembly with various auxiliary subunits (Stea et al., 1994; Dunlap et al., 1995; Patil et al., 1998).  However, even as early as the first studies to isolate full-length Cav2.1 cDNA from mammals, it was already evident that there are multiple variants predicted to result from alternative splicing of the CACNA1A gene (Mori et al., 1991; Starr et  20 al., 1991).  In these original studies, both groups independently identified multiple versions of the Cav2.1 channel with sequence diversity primarily in the intracellular loop between domains II and III and in the C-terminus.  Since these original reports, at least seven loci of alternative splicing in the Cav2.1 channel have been identified in neuronal tissues from mammals, many of which have functionally distinct biophysical and modulatory properties (Fig. 1.4).         21  Figure 1.4: Human Cav2.1 splice variants Seven known locations of alternative splicing in human Cav2.1 channels.  Top, the location of each of the seven splice sites on the Cav2.1 channel are indicated by green squares.  Bottom, all possible amino acid changes resulting from alternative splicing at the seven loci are shown.  Bold lettering indicates amino acids involved in splicing.  Dotted lines indicate where amino acids have been spliced out.  Slanted lines indicate a break in the amino acid sequence.  Adapted from (Soong et al., 2002)  Starting from the amino terminus (N-terminus), the first known region of alternative splicing occurs at the beginning of exon 10 within the intracellular loop connecting domains I-II (10/Δ10A/ Δ10B) (Fig. 1.4, #1) and has been identified in human (Soong et al., 2002) and rodent (Bourinet et al., 1999; Tsunemi et al., 2002; Kanumilli et al., 2006; Richards et al., 2007).  There can be an insertion of a valine and glycine (+VG; 10), insertion of glycine alone (+G; Δ10A), or no insertion of either amino acid (- ; Δ10B).  These alternative splice variants are a result of alternative splice acceptors within the 5’ region of exon 10 (Fig. 1.3D), wherein the two insertions use a non-canonical GT/TG donor-acceptor site and the Δ10B variant results from conventional splicing using a GT/AG donor-acceptor site.  In human  22 cerebellar RNA, the percentage of (10) was identified to be 17% with the Δ10A and Δ10B containing variants together making up the remaining 83% of transcripts (Soong et al., 2002).  In rats and mice, both variants are present in cerebellum, although in single Purkinje neurons all Cav2.1 transcripts lacked the valine insertion (Tsunemi et al., 2002; Kanumilli et al., 2006; Richards et al., 2007).  In the Cav2.1 rat homologue, the valine insertion (10) was shown to both enhance G-protein inhibition and up-regulate channel activity via PKC, as well as to decrease inactivation kinetics (Bourinet et al., 1999). A second region of splicing occurs within the IIS6 region of the channel where exons 16 and 17 can either be included (+16/17) or excluded (-16/17) in the final mRNA product (Fig. 1.4, #2), identified in human brain tissue (Soong et al., 2002).  Both exons are either included or excluded together and follow a customary exon inclusion/exclusion splicing event using a conical GT/AG donor-acceptor site pair (Fig. 1.3A).  Notably, the exclusion of the two exons has only been observed during transcript scanning of Cav2.1 channel fragments and both exons were found included in 100% of full-length cDNA clones analyzed from human cerebellum samples (Soong et al., 2002). A third alternative splice region is near the end of exon 17 in the II-III intracellular loop and results in either the insertion (+VEA; 17) or exclusion (-VEA; Δ17A) of a tripeptide repeat (Fig. 1.4, #3), identified in human brain (Hans et al., 1999a; Soong et al., 2002).  The tripeptide insertion/exclusion is due to the use of alternative splice donor sites at the 5’ end of intron 17 (Fig. 1.3C).  In cDNA clones obtained from human cerebellar samples only 0.4% possessed the VEA insertion (Soong et al., 2002). The functional consequences of the tripeptide insertion on Cav2.1 biophysical properties or channel modulation have not been reported. A fourth region of alternative splicing is the only site of alternative splicing that occurs in a segment constituting an extracellular portion of the channel (±31*) (Fig. 1.4, #4) and has been identified in human brain (Hans et al., 1999a; Soong et al., 2002), human spinal cord (Krovetz et al., 2000) and rodent brain (Bourinet et al., 1999; Toru et al., 2000).  The insertion (+NP; +31*) or exclusion (-NP; - 31*) of a dipeptide segment occurs within the IVS3-IVS4 extracellular loop.  Interestingly, the six nucleotide sequence encoding for NP is a small exon (exon 31*) lying within intron 31 and flanked by  23 canonical GT/AG acceptor-donor sites.  The ±31* variants are thus predicted to result from exon inclusion/exclusion (Fig. 1.3A).  In 1999 the Snutch lab determined that inclusion/exclusion of the dipeptide NP segment in rats determines Cav2.1 sensitivity to ω-Aga IVA (Bourinet et al., 1999).  It is now generally agreed that –NP corresponds to native P-type currents with a high sensitivity to ω-Aga IVA (IC50 in the low nanomolar (nM) range), whereas +NP corresponds to Q-type currents with a lower sensitivity to ω-Aga IVA (IC50 ~ 90 nM).  Ninety-five percent of cDNA clones isolated from human whole cerebellum possess the NP insertion (Soong et al., 2002).  In mice, the –NP variant was found almost exclusively in single Purkinje neurons and the +NP variant in granule cells (Toru et al., 2000; Tsunemi et al., 2002; Kanumilli et al., 2006; Richards et al., 2007). Within the proximal end of the C-terminus is a fifth location of alternative splicing involving the use of mutually exclusive exons 37a and 37b (37a/37b) (Fig. 1.4, #5) identified in human brain (Zhuchenko et al., 1997; Soong et al., 2002), human spinal cord (Krovetz et al., 2000) and rat brain (Bourinet et al., 1999).  The exons 37a and 37b are mutually exclusive exons with canonical GT/AG acceptor-donor sites (Fig. 1.3B).  The exons encode two versions of an EF-hand motif that contains conventional Ca2+ binding sites important in the Ca2+-dependent regulation of Cav2.1 channels (Kretsinger, 1976; Bourinet et al., 1999; Chaudhuri et al., 2004).  The two EF-hand variants alone appear to determine Ca2+-dependent modulatory properties of Cav2.1 with incredible selectivity.  The EFa variant confers the ability of channels to undergo forms of Ca2+-dependent modulation selective to local Ca2+ concentrations, whereas the EFb variant is locked in a normal gating mode (Chaudhuri et al., 2004; Chaudhuri et al., 2007).  There is approximately equal expression of the 37a (40.5%) and 37b (59.5%) variants in mammalian cerebellum, hippocampus and cerebral cortex.  Contrastingly, there is a large preferential expression of EFb in the amygdala and predominant expression of EFa in the substantia nigra and thalamus (Soong et al., 2002; Chaudhuri et al., 2004).  In rodent cerebellum, Purkinje neurons express almost exclusively EFa-containing channels (Kanumilli et al., 2006; Richards et al., 2007). Furthermore, there are critical changes in the expression pattern of the two variants during development and a clear gender bias in rodent and human brain (Chaudhuri et al., 2005; Chang et al., 2007).  24 A sixth location of alternative splicing identified in human brain and spinal cord involves exons 43 and 44 which lie within the C-terminus near the Ca2+ binding domain (±43/±44) (Fig. 1.4, #6) (Zhuchenko et al., 1997; Krovetz et al., 2000; Soong et al., 2002).  Exons 43 and 44 can be either included or excluded in all four combinations and result from two exon inclusion/exclusion splicing events that use canonical GT/AG acceptor-donor sites (Fig. 1.3A).  The presence of both exons was identified in 90% of cDNA clones isolated from human cerebellum, whereas 6% had +43/-44, 2% had - 43/+44 and 2% lacked both exons (Soong et al., 2002).  Similar findings were obtained from mouse cerebellum and single cerebellar Purkinje neurons (Kanumilli et al., 2006).  On the other hand, the proportion of the various combinations varied substantially in other areas of the human brain including the amygdala, cerebral cortex, hippocampus, hypothalamus, thalamus and substantia nigra.  In general, the proportion of transcripts containing the +43/-44 combination equaled those containing +43/+44, and the proportion containing neither exon was approximately 20-25% (Soong et al., 2002).  The inclusion/exclusion of these exons has been reported to either decrease voltage-dependent inactivation of Cav2.1 (Krovetz et al., 2000) or to have no effect (Soong et al., 2002), discrepancies likely resulting from coexpression with different β subunits (Patil et al., 1998). The seventh locus of alternative splicing for Cav2.1 channels is in the distal portion of the C- terminus (47/Δ47)(Fig. 1.4, #7) and has been identified in human brain (Zhuchenko et al., 1997; Hans et al., 1999a; Soong et al., 2002), human spinal cord (Krovetz et al., 2000) as well as in mouse Purkinje neurons (Toru et al., 2000; Tsunemi et al., 2002; Kanumilli et al., 2006; Richards et al., 2007).  The 47 and Δ47 splice variants result from the use of alternate canonical acceptor sites (Fig. 1.3D).  The use of an acceptor site immediately 5’ of exon 47 results in a pentanucleotide insertion (GGCAG) and frame shift that allows the full translation of exon 47.  Alternatively, the use of an acceptor site precisely at the exon 47 boundary yields an in-frame stop codon at the very beginning of exon 47.  In whole human cerebellum, 65% of transcripts have the pentanucleotide insertion and full translation of exon 47 (47) (Soong et al., 2002), whereas isolated human Purkinje and granule cells have approximately equal amounts of the 47 and Δ47 Cav2.1 variants (Tsunemi et al., 2008).  Studies in mice showed similar findings for whole cerebellum, although for isolated Purkinje neurons all Cav2.1 variants contained the  25 pentanucleotide insertion (Kanumilli et al., 2006).  Functional differences in the basic channel properties of the 47 and Δ47 Cav2.1 variant channels have not been reported. Two additional Cav2.1 variants recently identified in rat neuroendocrine cells from the supraoptic nucleus of the hypothalamus have large, 200-300 base pair deletions in the II-III cytoplasmic loop region containing the synprint site.  The deletion variants are predicted to have multiple consequences including altered coupling to synaptic release machinery, altered subcellular targeting and altered channel function (Rajapaksha et al., 2008). Taken together, alternative splicing is an important mechanism necessary in achieving the functional diversity of the Cav2.1 channel.  However, the relevance of Cav2.1 alternative splicing in the context of human diseases associated with the Cav2.1 channel has not been investigated.  The next section introduces the diseases associated with Cav2.1 channels and raises critical questions regarding a role for Cav2.1 alternative splicing in disease pathophysiology.       1.4 Cav2.1 Ca2+ channelopathies 1.4.1 General Cav2.1 channelopathies Owing to their diverse physiological roles, it is perhaps not surprising that disruption of VGCC function has been implicated in numerous severe human pathologies.  Diseases associated with VGCCs are referred to as Ca2+ channelopathies and to date there have been ten human channelopathies associated with five of the ten VGCC subunit genes (Appendix 1 for complete review of VGCC Ca2+ channelopathies). Of the three Cav2 α1 subunits, the only one known to be associated with mammalian genetic disorders is the Cav2.1 channel.  In humans, mutations in the CACNA1A gene encoding Cav2.1 are associated with three congenital, autosomal dominantly inherited neurological disorders: episodic ataxia type 2 (EA2), spinocerebellar ataxia type 6 (SCA6) and familial hemiplegic migraine type 1 (FHM-1). In mice, Cav2.1 genetic disorders include tottering (tg), leaner (tgla), rolling nagoya (tgrol) and rocker.  In  26 addition to the defined mutations in the Cav2.1 channel causing disease, an apparent autoimmune attack on Cav2.1 channels is associated with Lambert-Eaton Myasthenic Syndrome (LEMS).  I will first provide a brief overview of each of the human and mouse diseases associated with Cav2.1 channels and then a more in-depth discussion of FHM-1 as it pertains to this thesis. EA2 patients experience spontaneous episodes of ataxia (poor muscle coordination) that last for hours to days.  In between attacks, patients often experience gaze-evoked or down-beat nystagmus (rapid, involuntary eye oscillations).  Approximately 50% of patients also experience migraine-like symptoms and cerebellar atrophy is common (Lorenzon and Beam, 2000).  Attacks are often initiated by emotional stress, exercise or alcohol.  Most patients respond well to treatment with acetazolamide (reviewed in (Jen et al., 2004)).  EA2 is genetically variable and has been associated with Cav2.1 missense, truncation and alternative splice site mutations.     27  Figure 1.5: Cav2.1 mutations implicated in EA2 and Epileptic and Ataxic Mouse Models Location of missense mutations (black circles) and truncation mutations (white diamonds) in the human Cav2.1 channel associated with Episodic Ataxia Type-2 (EA2).  Also shown are mutations in the Cav2.1 mouse homolog associated with tottering (tg; purple circle), leaner (tgla; green circle), rolling nagoya (tgrol; yellow triangle) and rocker (orange triangle) phenotypes.  To date, more than 40 individual mutations in the CACNA1A gene have been associated with EA2.  The EA2 genetic alterations are distributed throughout the channel, with a large number of missense and premature truncations identified within the pore forming P-loops (Fig. 1.5).  Overall, the structure-function findings to date are relatively consistent across experimental conditions.  A number of both truncation and missense mutations consistently show a reduction in current density, proposed to be a result of fewer channels being properly folded and reaching the plasma membrane (Imbrici et al., 2004; Wan et al., 2005; Jeng et al., 2006; Pietrobon, 2010).  The three EA2 missense mutations G293R, C287Y, H1736L and a deletion mutation, nt 4778-4780, all demonstrate a net reduction of available channels due to a depolarizing shift in the voltage of half-maximum activation (V50act), an increased rate  28 of inactivation and a reduced rate of recovery from inactivation (Wappl et al., 2002; Spacey et al., 2004; Wan et al., 2005). The EA2 phenotype in humans is paralleled by the epilepsy and ataxia mouse models tg, tgla, tgrol and rocker (Fig. 1.5) which all contain Cav2.1 genetic mutations and show varying degrees of epilepsy and ataxia.  The tg, tgla and tgrol mutations have similar biophysical effects as those for EA2, namely decreased current densities and depolarizing shifts in V50act (Dove et al., 1998; Lorenzon et al., 1998; Wakamori et al., 1998; Mori et al., 2000).  In vivo studies in the animal models show that the reduced channel function seen in the heterologous systems translates to decreased neurotransmitter release in neocortex and at the parallel fibre to Purkinje synapses (Ayata et al., 2000; Matsushita et al., 2002) (for review on Cav2.1 models see (Pietrobon, 2005, 2010)). SCA6 is characterized by progressive cerebellar atrophy resulting in progressive gait ataxia, incoordination, nystagmus, proprioceptive sensory loss and dysarthria (Zhuchenko et al., 1997). Zhuchenko and coworkers found that patients with SCA6 possess a polyglutamine (CAG) expansion in exon 47, making SCA6 a member of the group of neurodegenerative disorders containing CAG repeats that includes Huntington’s disease amongst others.  Whereas unaffected people tend to have CAG repeats numbering between 4 and 16 in CACNA1A, patients with SCA6 have expansions of greater than 21 CAG repeats.  The length of the expansion appears directly correlated with age of onset, e.g., greater CAG expansion is associated with early age of disease onset (Ishikawa et al., 1997).  The CAG repeat expansion is associated with severe cerebellar Purkinje cell loss, moderate granule cell and dentate nucleus neuronal loss, and mild neuronal loss in the inferior olive (Zhuchenko et al., 1997).  The mechanisms involved concerning SCA6 mutations in the Cav2.1 channel and neuronal death have not been completely resolved.  However, biophysical analyses of polyglutamine expansions in the Cav2.1 channel expressed in heterologous systems show a range of effects on voltage and time-dependent properties with a strong dependence on both auxiliary subunit and α1 subunit splice-variant composition (Restituito et al., 2000; Toru et al., 2000).  It has been generally accepted that the polyglutamine stretches exert toxic effects by forming aggregates.  Analysis of cerebellar tissue from SCA6 patients revealed  29 perinuclear aggregates in Purkinje cells, and transfection of polyglutamine expanded Cav2.1 channel cDNA into human embryonic kidney (HEK) 293 cells demonstrated that cell death is likely due to the perinuclear aggregates (Ishikawa et al., 1999).  An interesting recent report showed that the C-terminus of the Cav2.1 channel is cleaved and translocated to the nucleus under wild-type conditions, but when the polyglutamine expansion is extended to greater than 33 repeats the nuclear translocated channel segment somehow induces cell death (Kordasiewicz et al., 2006).  In contrast, it has also been speculated that the loss-of-function effects on Cav2.1 channel function may result in reduced intracellular Ca2+ concentration and that the reduced Ca2+ alone can induce Purkinje neuron apoptosis and cerebellar atrophy (Matsuyama et al., 1999).  Overall, it appears that neuronal loss observed in SCA6 brains is likely due to a combination of altered channel properties resulting in abnormal intracellular Ca2+ concentrations and perinuclear and/or nuclear channel protein aggregates ultimately causing cell death. LEMS is a neuromuscular transmission disorder characterized by reduced acetylcholine quantal release and is associated with small-cell lung carcinoma (SCLC) in approximately 60% of patients (Lang et al., 1983).  Cav2.1 channels are implicated in LEMS, although unlike FHM-1, EA2 and SCA6, LEMS is not a true channelopathy as it does not result from defined mutations in the channel.  Rather the sera from LEMS patients contain auto-antibodies against VGCCs, with an apparent preferential targeting of Cav2.1 at the neuromuscular junction (Lennon et al., 1995; Pinto et al., 2002).  Clinical features of LEMS includes skeletal muscle weakness in proximal and trunk muscles, with the most severe effects observed in lower limbs (for review see (Flink and Atchison, 2003)).  Auto-antibodies are thought to be initiated in response to the SCLC tumour (O'Neill et al., 1988) and, via targeting Cav2.1 channels at the neuromuscular junctions, reduce channel availability for neurotransmission (Lennon et al., 1995).  It has been shown that the LEMS auto-antibodies do not alter channel voltage or kinetic properties, but instead act in an all-or-none fashion, likely eliminating available channels from the population (Grassi et al., 1994; Magnelli et al., 1996).  Drugs that prolong the duration of APs and enhance intracellular Ca2+ levels, such as 4-aminopyridine and 3,4-diaminopyridine, offer symptomatic relief in some LEMS patients, however, the side effects from treatment can often be severe (Flink and Atchison, 2003).  30  1.4.2 Introduction to FHM Migraine is a severe neurological condition that affects approximately 15% of the North American and Western European populations (Lipton et al., 1994; Lipton et al., 2001).  Migraine headaches are characterized by recurring unilateral headache that are often accompanied with nausea, phonophobia and/or photophobia.  The headaches can occur in isolation, or in approximately 20% of sufferers, the migraine headache can be preceded by or concurrent with an aura.  Auras are a subjective sensation most often associated with vision, although other sensory auras can occur  (reviewed in (Goadsby et al., 2002)).  The complicated genetics and physiology of migraine has slowed the development of adequate treatments and our understanding of underlying disease mechanisms (Montagna, 2004). FHM is a rare autosomal dominant subtype of migraine with aura, and other than a characteristic hemiplegia, has similar clinical features to typical migraine with aura and may share some pathogenetic mechanisms (Thomsen et al., 2002; Thomsen et al., 2003; Thomsen and Olesen, 2004).  Due to its relatively simple genetics, FHM has become a popular research model and has lead to relevant hypotheses for the pathophysiology underlying some aspects of typical migraine (Montagna, 2004).   About 50% of FHM patients have a mutation in the CACNA1A gene (Ophoff et al., 1996) (FHM type 1; FHM-1), whereas the other approximately 50% have mutations in either the ATP1A2 Na+/K+-ATPase gene (FHM-2) (De Fusco et al., 2003) or the SCN1A sodium channel gene (FHM-3) (Dichgans et al., 2005).  In FHM-1, the aura typically precedes the headache pain and manifests as an obligatory motor aura in combination with one or more other symptoms including visual disturbance, dysphasia (difficulty with speech) and/or sensory loss (usually numbness or paraesthesias of an extremity or the face).  The characteristic motor aura most frequently manifests as hemiplegia in both the upper and lower extremities.  In the majority of FHM-1 cases, the headache pain directly follows the aura phase of the migraine attack.  The headache pain can last from less than 30 minutes to greater than 72 hours, with the mean duration being approximately 24 hours.  In addition to these common symptoms, some FHM-1  31 patients also show permanent cerebellar symptoms that may include progressive cerebellar ataxia, and/or nystagmus, with cerebellar atrophy in some cases (for an extensive review of FHM-1 features and statistics see (Thomsen et al., 2002)).  1.4.3 Mechanisms of migraine pathophysiology It is generally accepted that both typical migraine and FHM-1 migraine attacks start in the brain (Lauritzen, 1994; Charles, 2009; Goadsby et al., 2009; Levy et al., 2009; Olesen et al., 2009).  Based on evidence from neuroimaging studies in human migraine patients (Bowyer et al., 2001; Hadjikhani et al., 2001) and other animal studies, it is apparent that the aura phase of migraine results from cortical spreading depression (CSD).  CSD is a transient wave of neuronal hyperexcitability that begins at a focal point and slowly progresses over the cortex (2-6 mm/min), followed by a long neuronal depression lasting minutes (reviewed in (Lauritzen, 1994)).  The headache pain component of migraine is thought to involve the meningeal nociceptors and brainstem.  Pain sensitivity within the skull is primarily restricted to meningeal blood vessels, and in conscious patients, electrical stimulation of the dura can cause headache pain through activation of branches of the ophthalmic division of the trigeminal nerve afferents that innervate the meningeal blood vessels (Penfield and McNaughton, 1940; Ray and Wolff, 1940) (reviewed in (Pietrobon and Striessnig, 2003; Pietrobon, 2005)).  An important link between CSD and headache pain was made by two separate animal studies that showed CSD can cause activation of meningeal trigeminovascular afferents and evoke a series of alterations in the meninges and brainstem that are consistent with the activation of trigeminal nociceptive pathways (Bolay et al., 2002; Moskowitz et al., 2004).  Thus at the macroscopic level, the current model of migraine is that CSD underlies the aura phase of migraine and activates the trigeminal pain pathway resulting in headache pain (Fig. 1.6), although a mechanism has not been proposed for the cerebellar dysfunction observed in some FHM-1 patients. At the tissue and cellular levels, during CSD in rat cortex (Fig. 1.6; 1) there is a substantial change in the composition of glutamate, K+, hydrogen ions (H+), nitric oxide (NO), arachidonic acid and  32 prostaglandins in the extracellular fluid (Somjen, 2002).  These molecules activate and/or sensitize the meningeal trigeminovascular afferents either directly or via perivascular inflammation (Wei et al., 1992; Strassman et al., 1996) (Fig. 1.6; 2).  Activation of the trigeminovascular afferents causes the release of vasoactive neuropeptides in their peripheral nerve endings which cause vasodilation of meningeal vessels and secretion of proinflammatory substances into the dura (Goadsby and Duckworth, 1987; Markowitz et al., 1988; Goadsby and Edvinsson, 1993; Williamson et al., 1997).  In addition, activation of trigeminovascular afferents leads to a subsequent activation of second-order neurons comprising the trigeminal nucleus caudalis (TNC) and the upper divisions of the cervical spinal cord (Fig. 1.6; 3,4). From there, two distinct pathways are activated.  First, activation of the superior salivatory nucleus sends impulses that cause further vasodilation of the meningeal blood vessels (Fig. 1.6; 5) and further activation of the trigeminal nerve creating a positive feedback cycle.  Second, TNC activation leads to stimulation of trigeminal efferents that project to the ventral posterior medial (VPM) nucleus of the thalamus and periaqueductal gray region (PAG).  The VPM sends projections that activate pain centers in the cortex (Fig. 1.6; 6).  A self-sustained central sensitization or disinhibitory sensitization of second- order trigeminovascular neurons involving sensory innervation from periorbital skin and neck muscles, descending inhibitory pathway from the PAG (Knight et al., 2002) and facilitatory pathways from the medulla (Urban et al., 2005) likely play key roles in maintaining the severe prolonged pain of migraine headache (for comprehensive review of migraine mechanisms see (Pietrobon and Striessnig, 2003; Moskowitz et al., 2004; Burstein and Jakubowski, 2005; Edvinsson and Uddman, 2005; Pietrobon, 2005; Sanchez-Del-Rio et al., 2006; Goadsby et al., 2009)).    33    Figure 1.6: Pathophysiological mechanisms of migraine with aura. 1) Cortical spreading depression (CSD) begins at a focal point and propagates as a wave across the cortex.  2) As the wave of neuronal hyperexcitability moves, excess glutamate, K+ ions, H+ ions, NO, arachidonic acid and prostaglandins build up in the extracellular space surrounding cortical neurons.  3) As these ions diffuse, they depolarize meningeal trigeminovascular afferents (trigeminal nerve).  4) Activation of the trigeminal nerve leads to activation of the superior salivatory nucleus which in turn sends projections that cause vasodilation of the meningeal blood vessels (5).  Dilation of the blood vessels leads to further activation of the trigeminal nerve, creating a positive feedback cycle.  6) Activation of the trigeminal nucleus caudalis also leads to stimulation of trigeminal efferents that project to the ventral posterior medial (VPM) nucleus of the thalamus.  The VPM sends projections that activate pain centers in the cortex.     34 1.4.4 The role of Cav2.1 channels in molecular mechanisms of FHM-1 Cav2.1 channels are highly expressed within areas associated with FHM-1 pathophysiology. They predominate in most neurons in the cerebral cortex (Westenbroek et al., 1995; Timmermann et al., 2002) and are the dominant mediators of glutamate release in cortical neurons (Turner et al., 1992; Qian and Noebels, 2001).  Cav2.1 channels account for about 40% of the whole cell Ca2+ current in trigeminal ganglion neurons (Borgland et al., 2001; Xiao et al., 2008; Davies and North, 2009) and contribute to the release of vasoactive neuropeptides from perivascular terminals of meningeal nociceptors (Akerman et al., 2003).  Cav2.1 channels are also involved in the descending inhibitory pathway from the PAG (Knight et al., 2002) and facilitatory pathways from the medulla (Urban et al., 2005), which regulate trigeminal pain transmission.  Cav2.1 channels are also involved in inhibitory control of the TNC by inputs from the dura (Ebersberger et al., 2004).  Furthermore, Cav2.1 channels are highly expressed in cerebellum in both Purkinje and granule cells (Randall and Tsien, 1995) and play predominant roles in both excitatory and inhibitory neurotransmission in cerebellum (Mintz et al., 1995; Iwasaki et al., 2000; Stephens et al., 2001). To date, there have been 21 mutations in the CACNA1A gene identified in patients with FHM-1 (Fig. 1.7).  The mutations all produce amino acid substitutions in conserved functional domains including the S4 voltage sensors and flanking regions, and the pore forming P-loops and S6 transmembrane segments.  Analyses of 13 FHM-1 mutations in heterologous expression systems and two FHM-1 knock-in mouse models show a trend in FHM-1 mutation effects that begin to explain the molecular mechanisms behind at least the aura phase of migraine.   35  Figure 1.7: Missense mutations in Cav2.1 associated with FHM-1 There are 21 reported missense mutations in Cav2.1 channels identified in patients with FHM-1.  Top, shows the location of mutations throughout conserved functional domains of the Cav2.1 channel. Bottom, lists the locations and changes in amino acids.  The colours indicate the phenotype associated with the mutation in patients: green = pure FHM-1, red = FHM-1 with cerebellar signs, and blue = severe FHM-1, cerebellar signs, diffuse encephalopathy and sometimes coma.  In recombinant human Cav2.1 channels, the introduction of mutations associated with both pure FHM-1 and FHM-1 with cerebellar signs show a hyperpolarizing shift in V50act (Kraus et al., 1998; Hans et al., 1999b; Melliti et al., 2003; Mullner et al., 2004), indicating the majority of mutant channels are available for opening at lower membrane potentials.  Similarly, several mutations cause an increase in open-channel probability and single channel conductance (Hans et al., 1999b; Tottene et al., 2002; Tottene et al., 2005).  All of these effects are predicted gain-of-function phenotypes with the potential to increase Ca2+ influx at lower membrane potentials (for full review of results from recombinant studies see (Adams and Snutch, 2007), Appendix 1) The impact of FHM-1 mutations on endogenous P/Q-type currents and neurotransmission in the cortex has been determined directly using two FHM-1 knock-in mouse strains.  One of the FHM-1  36 strains contains the R192Q mutation which in human patients is associated with a mild form of pure FHM-1 (migraine without cerebellar signs) (Ophoff et al., 1996; Ducros et al., 2001), whereas the other strain contains the S218L mutation which in human patients causes severe hemiplegic migraine and may include progressive cerebellar ataxia and atrophy, epileptic seizures, coma or stupor, and severe (sometimes fatal) cerebral oedema triggered by minor head trauma (Fitzsimons and Wolfenden, 1985; Kors et al., 2001; Chan et al., 2008).  Neither homozygous nor heterozygous R192Q mice show any overt phenotype, however, homozygous S218L mice exhibit the main clinical features identified in humans, including mild permanent cerebellar ataxia, spontaneous attacks of hemiparesis and/or generalized seizures and can include brain oedema after mild head impact (van den Maagdenberg et al., 2004; van den Maagdenberg et al., 2010).  Studies using the mouse strains have provided compelling evidence that the gain-of-function effects of the FHM-1 mutations measured in recombinant systems are recapitulated in cortical neurons from the knock-in mice.  The mutations have been shown to enhance excitatory neurotransmission in the FHM-1 mice due to increased Ca2+ influx during APs and increased glutamate release at pyramidal cell synapses, predicted to both initiate and perpetuate components of a positive feedback cycle that leads to CSD in FHM-1 (Tottene et al., 2009).  In support, in vivo studies in FHM-1 mice show a lower threshold for stimulation of CSD and an increased velocity of propagation across the cortex (van den Maagdenberg et al., 2004; Gherardini et al., 2006; van den Maagdenberg et al., 2010).  1.4.5 A key area in future FHM-1 research In some regards, there is compelling evidence from both the heterologous systems and knock-in mice to support the hypothesis that FHM-1 mutations increase channel availability, enhance Ca2+ influx and increase neurotransmitter release and thereby cause greater susceptibility to CSD causing aura.  To date however, no data have been presented to explain the headache pain phase or cerebellar dysfunction associated with FHM-1.  Furthermore, the FHM-1 phenotype is both highly localized to specific brain regions and episodic in nature, yet the Cav2.1 channels are expressed at nearly all fast synapses in the  37 CNS and PNS, and during most stages of development.  Why aren’t all regions that express Cav2.1 channels affected by the FHM-1 mutations?  Also, there is often considerable discrepancy in how the FHM-1 mutations affect recombinant Cav2.1 channels when tested by different research groups.  Over the past ten years several labs have investigated the effects of FHM-1 mutations on the biophysical properties of recombinant Cav2.1 channels expressed in Xenopus oocytes and mammalian expression systems.  Although many findings agree that FHM-1 mutations result in gain-of-function effects, some results vary considerably between research groups, some even contradictory.  For example, some researchers find that the R192Q mutation increases Cav2.1 whole cell current density (Hans et al., 1999b; van den Maagdenberg et al., 2004) while others find a decrease in whole cell current density (Tottene et al., 2002; Cao and Tsien, 2005).  In addition, while one lab found that the most prevalent FHM-1 mutation (T666M) shifts the voltage dependence of activation to more hyperpolarized potentials (Kraus et al., 1998), another group reported no change in voltage-dependent properties (Cao and Tsien, 2005). Also of note, mutations found in patients with similar clinical phenotypes have reported opposing effects on the biophysical properties of Cav2.1 channels.  For example, K1336E and V714A have both been associated with pure FHM-1 (without cerebellar signs) but cause either increased (Mullner et al., 2004) or decreased (Kraus et al., 1998) current decay in response to 1 Hz square pulse repetitive stimulations. These types of disparities create significant challenges in interpreting results obtained from different researchers as well as in correlating findings between recombinant and native Cav2.1 channels. These discrepancies and how FHM-1 mutations might only affect some Cav2.1 channels in specific brain regions under certain conditions remain largely unexplored.  Different expression systems (Tottene et al., 2002) or association with different auxiliary subunits (Mullner et al., 2004) may account for a portion of the disparity and might contribute to the localized, episodic nature of the FHM-1 phenotype.  There are other possibilities, however, that have not been explored.  Critically, whether the various known Cav2.1 splice variants respond differently to the presence of FHM-1 missense mutations, and whether this contributes to spatial and temporal aspects of the disease phenotype has not been explored.  38 Additional questions regarding FHM-1 that have not been previously explored include whether mutations associated with FHM-1 alter Cav2.1 Ca2+-dependent modulation and whether any effects on channel modulation might alter synaptic signalling mechanisms.       1.5 Ca2+-dependent modulation 1.5.1 Introduction to Ca2+-dependent modulation of VGCCs In addition to various forms of protein phosphorylation and G-protein modulation, HVA VGCCs are powerfully modulated by Ca2+ itself.  Ca2+ ions entering through channels bind various Ca2+ sensors and modulate channel activity under certain conditions (Meyers et al., 1998; Lee et al., 2002; Haeseleer et al., 2004; Zhou et al., 2004; Few et al., 2005; Lautermilch et al., 2005).  The most robust and well understood forms of Ca2+-dependent modulation of Cav1 and Cav2 channels are mediated by the ubiquitous Ca2+ sensing protein calmodulin (CaM) (reviewed in (Halling et al., 2005)). CaM consists of an N-terminal and C-terminal lobe connected by a central α helix.  Each lobe contains two Ca2+-binding E-F hand motifs, which when occupied by Ca2+, expose hydrophobic pockets that modulate the activity of target proteins (Halling et al., 2005).  A single Ca2+-free form of the CaM molecule (apoCaM) is constitutively complexed with the C-terminus of HVA VGCCs at an isolucine- glutamine (IQ) domain, and interacts with a second downstream site called the CaM binding domain (CBD).  Ca2+ binding to the C-terminal or N-terminal lobes of CaM, can each cause distinct forms of channel regulation (DeMaria et al., 2001; Pitt et al., 2001; Erickson et al., 2003; Lee et al., 2003; Liang et al., 2003; Mori et al., 2004).  Ca2+-dependent facilitation (CDF) is an augmentation of channel opening in response to a series of depolarizations.  Ca2+-dependent inactivation (CDI) is an enhancement of channel closing that occurs during prolonged depolarizations (a process distinct from VDI which progresses at a slower rate and persists when Ba2+ is the permeant ion).  Although only Cav2.1 channels undergo CDF, all Cav1 and Cav2 channels undergo CDI.  39 CDI regulation of Cav1 and Cav2 channels by CaM exhibits exquisite modes of spatial Ca2+ sensitivity.  The N-terminal lobe of CaM responds to a global rise in intracellular Ca2+ levels and causes CDI of the Cav2 channel family.  Alternatively, the C-terminal lobe of CaM responds to a local rise in Ca2+ levels within microdomains surrounding the channel pore and causes CDI of the Cav1 channel family (Liang et al., 2003; Chaudhuri et al., 2007).  The difference in spatial selectivity of CaM when associated with Cav1 and Cav2 channels is due, at least in part, to an N-terminal spatial Ca2+ transforming element (NSCaTE).  NSCaTE is a 13 amino acid sequence located between the first 80 and 100 amino acids in the proximal N-terminus of Cav1 channels that interacts with CaM molecules bound to the C- terminus of the channel.  Cav2 channels on the other hand lack the NSCaTE motif (Dick et al., 2008). CDI is not a an all-or-nothing process as some Cav1 channels contain a C-terminal CDI-inhibiting module (Singh et al., 2006; Singh et al., 2008) that can act as an enzymatic competitive inhibitor that retunes channel affinity for apoCaM and alters the magnitude of CDI (Liu et al., 2010). Critical roles of CDI of Cav1 and Cav2 channels in biological processes are becoming more apparent.  For example, Ca2+/CaM association with Cav1 channels is necessary for activation of the Ras/mitogen-activated protein kinase pathway, which conveys local Ca2+ signals from Cav1 channels to the nucleus to regulate gene transcription (Dolmetsch et al., 2001).  Also, Ca2+/CaM regulation of Cav1 channels is essential for moment-to-moment control of heart rate by controlling the duration of APs. Disruption of CaM binding to Cav1 channels causes a 4- to 5-fold prolongation of the cardiac AP (Alseikhan et al., 2002).  In addition, roles for Cav1.3 channel CDI in the pathophysiology of human diseases such as Parkinson’s disease, Alzheimers and/or schizophrenia have been proposed (Liu et al., 2010).  1.5.2 CDF and CDI of Cav2.1 channels To date, Cav2.1 channels are the only VGCCs that exhibit dual regulation mediated by CaM. When associated with Cav2.1 channels, the N-terminal lobe of CaM responds to a global rise in the intracellular Ca2+ level and induces CDI of Cav2.1.  The C-terminal lobe of CaM on the other hand,  40 responds to a local rise in Ca2+ levels within microdomains surrounding the pore of the channel and causes CDF.  Both recombinant and endogenous Cav2.1 channels have been shown to possess robust forms of CDF and CDI (Lee et al., 1999a; Lee et al., 2000; DeMaria et al., 2001; Lee et al., 2003; Chaudhuri et al., 2005; Kreiner et al., 2010). Human recombinant Cav2.1 channels transiently expressed in HEK cells (along with auxiliary subunits β2a and α2δ) is a well characterized system and allows for the clear isolation and measurement of CDF and CDI (illustrated in Figure 1.8A).  With a single depolarization to +5 mV from a holding potential of -90 mV, Cav2.1 channels initially open into a normal mode of gating characterized by a relative low steady-state open probability (Po) and elicit a relatively small Ca2+ current  (Fig. 1.8A; 1, left panel).  As Ca2+ enters through the channel, two Ca2+ ions bind the C-terminal lobe of preassociated apoCaM and subsequent Ca2+ currents through the channel are enhanced (i.e. CDF) (Fig. 1.8A; 2, left panel) (Lee et al., 1999a; DeMaria et al., 2001; Lee et al., 2003).  The Ca2+/CaM complex is thought to selectively induce a conformational change in the channel structure that favors channel opening. Facilitated Cav2.1 channels have a “facilitated mode of gating” featuring a genuine enhancement of Cav2.1 steady-state Po (Fig. 1.8C; left two panels) (Chaudhuri et al., 2007).  During paired depolarizations, channels become facilitated during the first depolarization as Ca2+ enters, and during the second depolarization, Cav2.1 channels are already facilitated and open immediately with a facilitated mode of gating that elicits maximum Ca2+ currents (Fig. 1.8A; 2, middle panel).  When Ba2+ is the permeant ion, channels remain in the  low Po state and exhibit the “unfacilitated” or “normal” gating mode and do not transition to the facilitated state regardless of prepulse potential (Fig. 1.8A; right panel), confirming CDF is a true Ca2+-dependent process (Chaudhuri et al., 2007).  Strong CDI of Cav2.1 is evident during prolonged depolarizations, in which whole cell Ca2+ currents decay much faster relative to Ba2+ currents (Fig. 1.8B).  As global Ca2+ levels rise, two Ca2+ ions bind the N-terminal lobe of resident CaM molecules and induce additional conformational change that inactivates channels (Fig. 1.8 C; right panel) (Chaudhuri et al., 2007).  41  Figure 1.8: CDF and CDI of Cav2.1 channels A.  Left; when Ca2+ is used as the charge carrier, wild-type Cav2.1 channels respond to a +5 mV square test pulse with an initial rapid activation (1) followed by a slow phase of current increase as Ca2+ enters through the channel, due to CDF (2).  Middle; on the other hand, when preceded by a +20 mV prepulse, channels are already facilitated at the beginning of the test pulse and activate rapidly with a maximum current response (2).  Right; when Ba2+ is used as the charge carrier, channels open to the low current level and remain there.  B. During a 1 second +10 mV test pulse, Ca2+ currents inactivate faster than Ba2+ currents due to CDI (DeMaria et al., 2001).  C. Mechanisms that underlie the two phenomena in A and B.  Left panel; without a prepulse wild-type channels open with a normal gating mode (1) then transition to a facilitated mode of gating as Ca2+ enters and binds the C-lobe of CaM constitutively bound to the C- terminus of the channel (2) (CDF) (middle panel).  Right panel; a rise in global Ca2+ levels through many channels, causes Ca2+ to bind the N-lobe of CaM which induces further conformational change and channel inactivation (Chaudhuri et al., 2007).  D. 100 Hz APW derived from APs recorded in the calyx of Held (Borst and Sakmann, 1998; Patil et al., 1998) (left panel).  Right two panels show representative traces when using either Ca2+ or Ba2+ as the charge carrier.  All currents were obtained from transiently transfected human recombinant Cav2.1 channels in HEK cells.    42 Although square voltage pulses allow maximum resolution of both CDF and CDI, the physiological relevance of CDF and CDI is more evident under simple action potential waveforms (APWs) (Fig. 1.8D).  As evident, the dual feedback regulation of Cav2.1 channels by CaM causes activity-dependent changes of Ca2+ currents during repetitive APs, first by enhancing the Ca2+ transients during initial APs (Fig. 1.8D; 1 and 2) and then reducing Ca2+ transients during later APs (Fig. 1.8D; 3). These powerful forms of rapid regulation have the potential to strongly influence Ca2+ dynamics in presynaptic terminals expressing Cav2.1 channels in the CNS.  1.5.3 Ca2+-dependent modulation of Cav2.1 channels in synaptic plasticity Short and long forms of synaptic plasticity including facilitation and/or depression of synaptic response during repetitive APs affect the dynamics and strength of neural circuits and are essential for the encoding, processing and storage of information in the CNS.  It has been known for many years that activity-dependent dynamics of Ca2+ signalling within presynaptic terminals is a central determinant of short-term synaptic plasticity (Katz and Miledi, 1968).  The earliest empirical evidence showed that presynaptic Ca2+ currents themselves do not change during repetitive stimulation at the squid giant axon synapse (Charlton et al., 1982).  This established the notion that mechanisms of synaptic plasticity at most fast synapses in the CNS are likely downstream of Ca2+ entry and rather associated with molecular complexes directly involved in vesicle fusion (termed the “residual Ca2+ hypothesis”).  The residual Ca2+ hypothesis has been the most widely accepted mechanism of short-term synaptic plasticity for the past thirty years.  It states that facilitation of synaptic responses during repetitive APs is due to the accumulation of intracellular Ca2+ in presynaptic terminals, and that build-up of residual Ca2+ enhances binding to Ca2+ sensor proteins (CaS) (such as synaptotagmin) which directly mediate vesicle fusion and transmitter release.  Conversely, decay in synaptic responses during repetitive APs is believed to primarily be due to the depletion of the readily available pool of neurotransmitter containing vesicles (reviewed in (Zucker and Regehr, 2002)).  However, this traditional explanation of short-term synaptic plasticity is being challenged by recent findings that residual Ca2+ can act on other CaSs independent of  43 the release machinery (Blatow et al., 2003; Felmy et al., 2003; Muller et al., 2007), and further, that modulation of presynaptic Ca2+ currents themselves can in fact be a means to achieve short-term synaptic plasticity at a CNS synapse. The calyx of Held is a large presynaptic terminal in the mammalian CNS which synapses onto cell bodies of principal neurons in the medial nucleus of the trapezoid body.  Whole-cell voltage clamp recordings on the calyx of Held in mice show that an activity- and Ca2+-dependent enhancement of presynaptic Ca2+ currents accounts for about 40% of the total facilitation of synaptic response during repetitive APs (Forsythe, 1994; Borst and Sakmann, 1998; Cuttle et al., 1998; Forsythe et al., 1998; Inchauspe et al., 2004; Xu and Wu, 2005; Muller et al., 2008).  There is also some evidence that inactivation of presynaptic currents mediates short-term synaptic depression under some conditions (Xu and Wu, 2005).  Although both Cav2.1 and Cav2.2 channels are expressed presynaptically at this synapse, facilitation of presynaptic Ca2+ currents and the corresponding changes in synaptic facilitation are dependent solely on Cav2.1 channels (Inchauspe et al., 2004; Ishikawa et al., 2005; Xu and Wu, 2005; Inchauspe et al., 2007).  At the mechanistic level, it has been proposed that these processes in the calyx of Held are mediated by the neuronal Ca2+ sensor NCS-1 (Tsujimoto et al., 2002), although some descrepencies remain (Rousset et al., 2003).  Clarity regarding the mechanism of Cav2.1 channel modulation and whether it represents a general mechanism of synaptic plasticity at other CNS synapses remains to be established. In 2008, work from Catterall and Mochida provided the first evidence in a model system that CDF and CDI of Cav2.1 is mediated through CaS such as CaM that bind the IQ-like domain and contributes to both the induction of short-term synaptic facilitation and rapid synaptic depression (Mochida et al., 2008).  The model system consisted of recombinant Cav2.1 channels transfected into cultured superior cervical ganglion (SCG) neurons.  SCG neurons do not express endogenous Cav2.1 channels and when cultured in vitro form fast cholinergic synapses (Mochida et al., 2003a; Mochida et al., 2003b).  Mochida et al. showed that after transfection of cultured SCG neurons with wild-type Cav2.1 channels, both robust CDF and CDI of Ca2+ currents could be measured in the presynaptic neurons.  44 Further, a corresponding synaptic facilitation and depression, respectively, could be detected in excitatory post-synaptic potentials (EPSPs) evoked from pairs of synaptically connected SCG neurons in which only the presynaptic neuron was transfected with Cav2.1 channels.  In contrast, when Cav2.1 channels harboring mutations in the IQ-like and CBD binding domains that prevent CaS binding were transfected into the SCG neurons, CDF and CDI of presynaptic currents was not detected.  Furthermore, paired-pulse facilitation (PPF) and facilitation and depression of EPSPs in response to trains of APs were reduced significantly (Mochida et al., 2008).  The results provide compelling evidence for a model in which CaM or other CaS proteins respond to residual Ca2+ and mediate Ca2+-dependent modulation of Cav2.1 channels by binding to the IQ-like motif in the C-terminus of the channel and induce short-term synaptic plasticity under repetitive APs.  However, it remains to be described whether this mechanism described in the model system applies to intact fast synapses in the CNS.  1.5.4 Key areas concerning future research on Cav2.1 modulation and synaptic plasticity The centrality of Cav2.1 in mediating presynaptic Ca2+ influx at nearly all fast presynaptic terminals in the CNS suggests such forms of CDF and CDI of Cav2.1 may be widespread mechanisms of short-term synaptic plasticity.  Interestingly, some FHM-1 missense mutations result in an overall gain- of-function Cav2.1 channel phenotype as a result of enhanced Po (Hans et al., 1999b; Tottene et al., 2002) and as described, CaM mediated CDF of Cav2.1 renders channels toward a “facilitated mode of gating” due to an enhancement of Cav2.1 Po.  Although the effects of FHM-1 mutations on CDF and CDI of Cav2.1 have not been reported, the fact that CDF/CDI and FHM-1 mutations both alter Cav2.1 channel gating suggests FHM-1 mutations may also interfere with the normal Ca2+-dependent regulation of Cav2.1 channels.  Further, if these forms of CDF and CDI of Cav2.1 are required components of synaptic plasticity, FHM-1 mutations might directly affect synaptic efficacy and have significant implications concerning disease pathophysiology. A prototypical CNS synapse where Cav2.1 CDF may play an important role in synaptic plasticity and also be important in FHM-1 pathophysiology (at least in severe forms of FHM-1 associated with  45 cerebellar signs), is the parallel fibre (PF) to Purkinje cell (PC) synapse in the cerebellar cortex.  The PF- PC synapse is a well-characterized central synapse that displays robust presynaptic forms of short-term facilitation via both paired-pulse and AP trains and is known to rely predominantly on Cav2.1 channels for neurotransmitter release (Mintz et al., 1995; Randall and Tsien, 1995).  Also, facilitation at this synapse is mediated by CaSs similar to those reported to mediate Cav2.1 CDF (Atluri and Regehr, 1996; Lee et al., 1999a; Lee et al., 2000; DeMaria et al., 2001; Lee et al., 2002; Tsujimoto et al., 2002; Lee et al., 2003; Chaudhuri et al., 2004; Chaudhuri et al., 2005; Few et al., 2005; Chaudhuri et al., 2007).       1.6 Cav2.1 and synaptic plasticity at the PF-PC synapse 1.6.1 Brief cerebellum overview The cerebellum plays a central role in motor control, reflex adaptation and motor learning.  The structure of the cerebellum in mammals is highly organized and composed of repeated modules consisting of three layers of well-defined cell types (molecular, Purkinje and granular layers) (Fig. 1.9 A) (for full review of cerebellar anatomy and circuitry see (Voogd and Glickstein, 1998; Ito, 2000, 2002; Boyden et al., 2004)).  In brief, inputs to the cerebellum derive from various regions of the CNS and PNS through precerebellar nuclei axons called mossy fibers (MFs).  MFs branch extensively and create large presynaptic terminals (glomeruli) that form excitatory glutamatergic synaptic connections with dendrites of hundreds of granule cells (GCs) within the granule layer of the cerebellar cortex (Fig. 1.9 B).  GCs axons ascend to the molecular layer where they bifurcate to form the PFs, which extend 2-3 mm in either direction along the transverse plane making many en passant synapses on the dendritic spines of tens of thousands of PCs (the PF– PC synapse) (Fig. 1.9B).  Each PC receives as many as 60,000 to 175,000 PF inputs (Napper and Harvey, 1988).  PCs have large cell bodies that reside in the Purkinje cell layer and have extensive apical dendritic arbourization in the parasaggital plane that protrude into the molecular layer.  This elaborate dendritic tree is the largest and most highly branched of all neurons in the CNS, yet is confined within the saggital plane and at a right angle to the PFs.  PC cells also receive excitatory input from climbing fibres (CFs) that originate in the inferior olive and medulla oblongata.  Only a single  46 CF contacts each PC, however the CF wraps around the PC and can form upwards of 26,000 synapses along dendritic spines (Nieto-Bona et al., 1997).  Powerful inhibitory inputs from basket cells are made onto cell bodies of PCs and inhibitory input to PC dendrites is mediated by stellate cells that receive input from PFs.  The apical dendrites of Golgi cells lie within the molecular layer and receive input from the PFs and then provide inhibitory feedback to the GCs (Ito, 2000, 2002).  The PCs are responsible for the sole output of the cerebellar cortex.  PC axons project through the granule cell layer and white matter to make inhibitory GABAergic connections with neurons of the deep cerebellar nuclei (DCN).                  47            A B Figure 1.9: Main microcircuitry in the cerebellar cortex A) Three-dimensional illustration of the cerebellar cortex.  PC somas (brown) form the Purkinje cell layer, while extensive dendritic arbourizations of the PCs lie within the parasagittal plane and form the molecular layer.  The soma of GCs (light green) form the granule cell layer.   Axons of GCs ascend through the Purkinje cell layer to the molecular layer, bifurcate and run within the transverse plane of the cerebellum as PFs.  There are several GABAergic interneurons, including stellate cells (blue), basket cells (red), and golgi cells (purple) found in the cerebellar cortex that form synaptic connections with various cellular elements.  B) MFs (dark green) from various brain regions form excitatory glutamatergic synaptic connections with dendrites of hundreds of GCs.  Each PF (light green) forms many en passant glutamatergic excitatory inputs onto the distal dendritic spines of many PCs.  CFs (deep green) project directly from the inferior olive of the brainstem and form glutamatergic excitatory inputs onto PCs.  Each CF forms thousands of synaptic connections with the proximal dendrites of an individual PC.  The axons of PCs projects to DCN (purple) via inhibitory GABAergic synaptic connections and form the sole output of the cerebellum.  DCN neurons relay signals to the motor cortex via the thalamus. Adapted with permission from Purves et al., 2001, Neuroscience 2/e, 416-417.     48 1.6.2 Synaptic plasticity at the PF-PC synapse: a role for CDF of Cav2.1? The geometry and spatial equilibration of Ca2+ in the PF presynaptic terminals (or boutons) are ideal for measuring the role of Ca2+ dynamics on a tens-of-milliseconds time scale using Ca2+-sensitive fluorescent dyes while simultaneously measuring excitatory post-synaptic currents (EPSCs) in PCs (Mintz et al., 1995; Atluri and Regehr, 1996).  Such measurements have provided substantial information as to mechanisms of synaptic plasticity at this synapse.  Short-term synaptic facilitation is robust under conditions of both paired-pulse and short AP trains at the PF-PC synapse and is derived from presynaptic mechanisms (Schulz et al., 1994; Goto et al., 2006).  Although modulation of presynaptic Ca2+ currents via adenosine A1, GABAB and cannabinoids receptors can reduce synaptic strength at this synapse, synaptic facilitation is driven by Ca -dependent processes in PF boutons (Dittman and Regehr, 1996; Kreitzer and Regehr, 2000). B 2+ Though the precise Ca2+-dependent mechanisms that drive facilitation have not been fully elucidated, several properties of this terminal suggest CDF of Cav2.1 channels may be an important player.  For one, although a significant portion of facilitation can be explained by traditional mechanisms described by the residual Ca2+ hypothesis, some facilitation persists in the absence of residual Ca2+.  This has been demonstrated through experiments in which presynaptic terminals were treated with high (100 μM) concentrations of the Ca2+ chelator ethylene glycol tetraacetic acid bound to acetoxymethyl ester (EGTA-AM) to make it hydrophobic and amenable for uptake across cell membranes in live cells.  In these experiments, EGTA-AM sped the decay of free intracellular Ca  to only a brief impulse lasting a few milliseconds.  During paired pulses, facilitation of the second pulse was still present with interpulse intervals between 40 and 50 milliseconds (Atluri and Regehr, 1996).  The authors concluded that there are at least two mechanisms of facilitation at the PF-PC synapse, one driven by residual 2+ Ca2+ in the traditional sense, and another driven by CaSs with high Ca  affinity that can detect modest, transient levels of Ca  likely near the pore of presynaptic VGCCs.  Of note, it is known that CaM and other CaS proteins are both present in PFs and sensitive to moderate levels of Ca  (Ullrich et al., 1994; Li et al., 1995). 2+ 2+ 2+  49  While GCs contain Ca 2.1, Ca 2.2 and Ca 2.3 channels (Randall and Tsien, 1995), Ca 2.1 is responsible for about 60% of Ca  influx in PF terminals during APs and is most effective at triggering transmitter release.  As such, Ca 2.1 channels are responsible for nearly 93% of the synaptic response at the PF-PC synapse (Mintz et al., 1995) and even small changes in the Ca  influx through Ca 2.1 channels at these terminals have profound effects on the post-synaptic response.  In fact, there is a steep correlation between Ca  influx in PF boutons and transmitter release such that the power law of synaptic transmission at this synapse can range between 2 and 4 (Mintz et al., 1995; Sabatini and Regehr, 1997). Thus, the enhancement and broadening of APs predicted by CaM mediated CDF of Ca 2.1 (Chaudhuri et al., 2007) could have important implications for the robust short-term facilitation observed at this synapse. v v v v 2+ v 2+ v 2+ v While the PF-PC synapse is a prototypical synapse in the CNS, and the available evidence indicates Cav2.1 CDF may play an important role in synaptic plasticity, whether this is in fact an important regulatory mechanism at this synapse has not been experimentally shown.       1.7 Thesis hypotheses and objectives 1.7.1 Hypothesis 1 The Cav2.1 channel plays a central role in neurotransmitter release at nearly all fast synapses in the CNS, yet not all Ca 2.1 channels are the same.  As a result of alternative splicing, there are potentially thousands of functionally distinct Ca 2.1 splice variants that are differentially expressed within brain regions, cell types and subcellular compartments and at different developmental stages. Critically, the Ca 2.1 channel is associated with several autosomal dominant human diseases including FHM-1, yet whether the FHM-1 mutations differentially affect Ca 2.1 splice variants has not been tested. In this thesis I specifically hypothesize: v v v v That Ca 2.1 splice variants respond differentially to FHM-1 mutations and contribute to both the episodic nature of the disease phenotype and the localization of the disease to specific brain regions and cell types. v  50  1.7.2 Hypothesis 2 Ca -dependent modulation of Ca 2.1 channels is a robust form of channel modulation, however, whether mutations in the Ca 2.1 channel associated with human disease alter CDF or CDI has not been reported.  Additionally, CDF and CDI of Ca 2.1 are predicted to play critical roles in short-term synaptic plasticity in the CNS.  A direct demonstration of CDF or CDI of Ca 2.1 channels as important determinants of short-term synaptic plasticity at an intact central synapse has not been demonstrated.  In this thesis I specifically hypothesize: 2+ v v v v That FHM-1 mutations alter Ca -dependent modulation of Ca 2.1 channels, and further, that the changes in channel modulation affect synaptic plasticity. 2+ v  1.7.3 Thesis objectives The following scientific objectives were formulated to address the above hypotheses: 1. To use molecular genetic tools in combination with whole-cell electrophysiological analyses to compare the effects of several FHM-1 mutations on voltage-dependent and kinetic properties of two common human Ca 2.1 splice variants expressed in HEK cells.    v 2. To analyse the effects of FHM-1 mutations on CDF and CDI of human recombinant Ca 2.1 channels expressed in HEK cells. v 3. To verify effects on CDF or CDI utilizing two FHM-1 knock-in mouse strains expressing endogenous levels of Cav2.1 channels containing FHM-1 mutations. 4. To examine whether effects of FHM-1 mutations on CDF or CDI in recombinant and endogenous Cav2.1 channels affect short-term synaptic plasticity by measuring excitatory post- synaptic potentials in PCs during evoked APs in PFs in both wild-type and knock-in mice  51 cerebellar slices.  Also, to compare Ca  dynamics through the 2+ Cav2.1 channels in PF boutons from wild-type and FHM-1 knock-in mice using two-photon microscopy.                     52       1.8 Bibliography Adams PJ, Snutch TP (2007) Calcium channelopathies: voltage-gated calcium channels. Sub- cellular biochemistry 45:215-251. 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Since the first mutation in a Ca2+ channel was identified (Fontaine et al., 1994; Ptacek et al., 1994), over 150 individual mutations have now been reported in five of the ten genes encoding Ca2+ channel pore forming α1-subunits (Cav) and are associated with nine distinguishable disorders (“Ca2+ channelopathies”).  Over the past decade, studies using recombinant channels in various expression systems have shown many of these mutations have significant positive or negative effects on channel gating and/or expression levels, while others result in non-functional channels or have dominant negative effects (reviewed in (Adams and Snutch, 2007)).  It is noteworthy that the effects of mutations on channel function have thus far only been tested in a small subset of known Ca2+ channel variants and a direct comparison of how mutations affect channel alternative splice variants has been largely unexplored.     _______________________________ * A version of this chapter has been published. Adams, P.J., Garcia, E., David, L.S., Mulatz, K.J., Spacey, S.D. and Snutch, T.P. (2009) Cav2.1 P/Q-type Calcium Channel Alternative Splicing Affects the Functional Impact of Familial Hemiplegic Migraine Mutations: Implications for Calcium Channelopathies. Channels, Mar-Apr; 3(2):110-21.  73 It has been predicted that the ten genes encoding Cav subunits have the potential to generate thousands of functionally distinct splice variants (Lipscombe and Castiglioni, 2004; Emerick et al., 2006). Indeed, isolation and characterization of some variants has shown that alternative splicing can be a means to obtain specialized Ca2+ channel function and to optimize Ca2+ signalling regionally, temporally and under altered environmental conditions (reviewed in (Lipscombe et al., 2002; Lipscombe and Castiglioni, 2004; Gray et al., 2007)).  It is evident that mutations directly at a splice-site or in an alternate exon can have effects on pre-mRNA splicing and/or affect a subset of splice variants expressing alternate exons (Zhuchenko et al., 1997; Splawski et al., 2004; Splawski et al., 2005; Graves et al., 2008).  However, the majority of identified mutations associated with Ca2+ channelopathies are missense mutations in coding sequences other than splice-sites and alternate exons (reviewed in (Adams and Snutch, 2007)).  Whether channel splice variants have different functional responses to disease-causing missense mutations has not been explored.  We hypothesized that point mutations associated with Ca2+ channelopathies might have splice-variant specific effects with important implications for both understanding disease pathophysiology and also towards interpreting results obtained from heterologous studies using recombinant channels. Familial Hemiplegic Migraine (FHM) is an autosomal dominant subtype of migraine characterized by an aura of hemiplegia that is associated with at least one other aura symptom such as hemianopsia, hemisensory deficit, or aphasia (Ducros et al., 2001; Thomsen et al., 2002).  Approximately 20 missense mutations associated with FHM have been identified in the CACNA1A gene (Ophoff et al., 1996) (called FHM-1) which encodes the α1 subunit (Cav2.1; α1A) that conducts P/Q-type Ca2+ currents.  Cav2.1 channels are abundantly expressed throughout mammalian brain and spinal cord where they mediate Ca2+ influx essential for neurotransmitter release, Ca2+-mediated second messenger signalling and Ca2+-dependent gene transcription (Starr et al., 1991; Takahashi and Momiyama, 1993; Westenbroek et al., 1995; Bourinet et al., 1999; Sutton et al., 1999).  The functional consequences of FHM-1 mutations on Cav2.1 channel properties have been investigated in heterologous Xenopus oocyte and mammalian expression systems, and more recently in neurons and whole brains of FHM-1 mutant R192Q and S218L knock-in mice (Hans et al., 1999b; Kraus et al., 2000; Tottene et al., 2002; Mullner et al., 2004; van den Maagdenberg et al., 2004; Cao and Tsien, 2005; Tottene et al., 2005; Tottene A., 2005).  Biophysical analysis of FHM-1 effects on  74 Cav2.1 channels is controversial as both loss-of-function and gain-of-function effects have been reported, as well as no effect (Kraus et al., 1998; Hans et al., 1999b; Kraus et al., 2000; Tottene et al., 2002; Barrett et al., 2005; Cao and Tsien, 2005).  Yet despite the noted discrepancies, in both heterologous and knock-in mice systems there is a general demonstrated trend for FHM-1 mutations to exhibit gain-of-function properties: increased channel availability and increased Ca2+ influx at lower membrane potentials resulting in a greater susceptibility to the CSD thought to be the underlying mechanism of aura (Kraus et al., 1998; Hans et al., 1999b; Melliti et al., 2003; Mullner et al., 2004; van den Maagdenberg et al., 2004; Gherardini L., 2006). There are seven identified alternatively spliced sites within the Cav2.1 subunit gene and the various splice variants exhibit distinct biophysical characteristics, Ca2+-dependent properties, pharmacological sensitivities and subtype-specific temporal and regional localizations in human brain (Bourinet et al., 1999; Soong et al., 2002; Timmermann et al., 2002; Chaudhuri et al., 2004; Chang et al., 2007).  However, it is not known whether the functional impact of FHM-1 mutations is similar amongst the different Cav2.1 splice variants or whether alternative splicing contributes to the spatial and temporal nature of the FHM-1 phenotype.  The carboxyl terminus of Cav2.1 channels is known to affect several physiological processes and alternative splicing in this region confers functional changes in channel properties (Bourinet et al., 1999; Maximov et al., 1999; Krovetz et al., 2000; Restituito et al., 2000; Soong et al., 2002; Chaudhuri et al., 2004).  The most substantial changes induced by alternative splicing in the C-terminus of Cav2.1 channels results from the use of an alternative three prime acceptor site in the intron upstream of the last exon, exon 47 (Mori et al., 1991; Hans et al., 1999a; Krovetz et al., 2000; Soong et al., 2002).  Alternative splicing at exon 47 introduces a frame-shift resulting in a stop codon at the beginning of exon 47.  As a result, Cav2.1 channels can be of either the short form (isoform 1; Cav2.1 (Δ47)) or the long (isoform 2; Cav2.1 (+47)).  The voltage-dependent and kinetic properties of the Cav2.1 (+47) and Cav2.1 (Δ47) splice variants and their relative contributions concerning FHM-1 mutations has not been explored. In the present study we compared the biophysical properties of wild-type Cav2.1 (+47) and Cav2.1 (Δ47) channel splice variants and also explored the effects of three FHM-1 mutations introduced into the  75 two variants.  We investigated two mutations, K1336E and R192Q, that are associated with an FHM-1 phenotype of pure hemiplegia and migraine without any other neurological symptoms (Ophoff et al., 1996; Ducros et al., 2001).  We further investigated the S218L FHM-1 mutation which is associated with a severe clinical phenotype wherein typical FHM-1 attacks induced by minor head trauma are often followed by a delayed cerebral oedema, fever, stupor and sometimes coma (fatal in one reported instance) (Fitzsimons and Wolfenden, 1985; Kors et al., 2001; Chan et al., 2008).  We find that the two Cav2.1 channel carboxyl tail splice variants exhibit functionally distinct properties and also that the three FHM-1 mutations have differential splice-dependent effects on voltage-dependent and kinetic properties.  We discuss the potential importance of the splice-variant differential effects in the context of FHM-1 pathophysiology as well as the implications for other Ca2+ channelopathies.       2.2 Results 2.2.1 Cav2.1 (+47) and Cav2.1 (∆47) variants are expressed in human cortex  Mutations in the Cav2.1 channel underlie FHM-1 and the current consensus is that initiation of migraine attacks is in the cortex; however the expression of splice variants has not yet been described in the human cortex.  In order to determine whether the Cav2.1 (+47) and Cav2.1 (∆47) variants are expressed in human cortex we utilized RT-PCR to amplify a ~1.1Kb carboxyl terminal fragment of Cav2.1 from adult human cortex RNA using oligonucleotide primers that recognize both carboxyl alternatively spliced variants in a non-biased manner.  The PCR products were subsequently re-amplified using splice-variant specific primers.  Figure 2.1 shows that the Cav2.1 (+47) and Cav2.1 (∆47) splice variants are both expressed in human cortex.  To determine their relative proportions, the human cortical Cav2.1 carboxyl terminal PCR products were sub-cloned and individual cDNAs analyzed using splice-variant specific primers and direct DNA sequencing.  From the 53 cDNA clones analyzed we determined that the Cav2.1 (+47) and Cav2.1 (∆47) splice variants were present in whole cortex in relative proportions of 79% and 21%, respectively.  76  All subsequent biophysical analyses were performed using human long Cav2.1 (+47) and short Cav2.1 (∆47) splice-variant cDNA clones with either wild-type or FHM-1 mutant K1336E, R192Q or S218L changes introduced (see Fig. 2.1A for the location of the FHM-1 mutations).      77  Figure 2.1: Human Cav2.1 Ca2+ channel topology and splice-variant expression in human cortex. A, schematic showing the location of the three FHM-1 mutations and the carboxyl terminal splice site in the human Cav2.1 channel.  In the box below the channel diagram are partial sequences of the Cav2.1 (+47) and Cav2.1 (Δ47) variants at the exon 46/ exon 47 boundary.  The pentanucleotide insertion is shown in bold for the Cav2.1 (+47) variant.  B, the last ~ 1 Kb of the Cav2.1 carboxyl terminus was amplified from human cortical RNA and purified.  Subsequently, splice-variant specific forward primers designed to exclusively bind either Cav2.1 (Δ47) or Cav2.1 (+47) transcripts were used in PCR reactions to generate an ~500bp fragment from the purified carboxyl fragments; Cav2.1 (Δ47)-SP and Cav2.1 (+47)-SP, respectively (dotted line is above the sequence that Cav2.1 (Δ47)-SP binds and the solid line is above the sequence that Cav2.1 (+47)-SP binds in A.  Both splice-variant specific primers generated the expected product from the carboxyl PCR fragment of the Cav2.1 cDNA obtained from human cortex, verifying both Cav2.1 (Δ47) or Cav2.1 (+47) are present in human cortex.  Products were verified by direct DNA sequencing and determined to be in relative proportions of 79% Cav2.1 (+47) and 21% Cav2.1 (Δ47) (bar graph); for protocol details see Experimental Procedures.   78 2.2.2 FHM-1 mutations exhibit differential effects on the voltage-dependent properties of Cav2.1 splice variants Whole cell current analysis of transiently transfected cells showed that the wild-type Cav2.1 (Δ47) and wild-type Cav2.1 (+47) variants possess similar membrane potentials at which half the channels are activated (V50act = -14.02 ± 1.49, and -15.08 ± 1.20, respectively) and similar membrane potentials at which half of the channels are inactivated (V50inact = -58.20 ± 2.04, and -62.07 ± 1.87, respectively; see Table 2.1 and Fig. 2.2).  The K1336E, R192Q and S218L mutations have been previously reported to cause a hyperpolarizing shift in the current-voltage relationship relative to wild-type Cav2.1 channels (Melliti et al., 2003; Mullner et al., 2004; Cao and Tsien, 2005; Tottene et al., 2005; Weiss et al., 2008).  Examining the FHM-1 mutations in the Cav2.1 +47 and ∆47 carboxyl tail splice variants we found differential effects. The K1336E Cav2.1 (+47) and R192Q Cav2.1 (+47) channels both exhibited a small but significant shift in V50act relative to wild-type Cav2.1 (+47) channels (-21.53 ± 1.35 and -19.11 ± 1.11 vs. -15.08 ± 1.20, respectively; p <0.05; ANOVA), while the S218L mutation had no significant affect on V50act of Cav2.1 (+47) channels relative to wild-type (Table 1 and Fig. 2.2).  In contrast, all three FHM-1 mutations caused large significant hyperpolarizing shifts in V50act when expressed in Cav2.1 (Δ47) variant channels (p < 0.001; ANOVA; Table 1 and Fig. 2.2).  Similar differential splice-dependent effects of the FHM-1 mutations were apparent in examining V50inact.  Figure 2.2 shows, that the R192Q and S218L mutations resulted in large (~15 – 17 mV) hyperpolarizing shifts in V50inact in the Cav2.1 (∆47) variant relative to wild-type Cav2.1 (∆47) channels (p < 0.001; ANOVA), while these same two FHM-1 mutations had a smaller effects on Cav2.1 (+47) variant channels  (p < 0.05; ANOVA; Table 1).  Interestingly, the K1336E mutation did not cause a significant change in V50inact in either splice-variant.  Also, all three mutations had significantly different effects on kinetics of activation and inactivation as well as steepness of the curves for activation and inactivation (Table 2.1).  Overall, these data indicate that the impact of individual FHM- 1 mutations on Cav2.1 channel gating properties is differentially affected by the nature of the splice-variant background in which the mutation is expressed.   79   Figure 2.2: FHM-1 mutations differentially affect voltage-dependent properties of Cav2.1 (Δ47) and Cav2.1 (+47) variants.  A, shows the comparison between current-voltage relationships (IV-curves) for wild-type (WT; black squares), FHM-1 mutant K1336E (KE; grey circles), R192Q (RQ; grey triangles), and S218L (SL; grey diamond) in both the short Cav2.1 (Δ47) (filled symbols) and long Cav2.1 (+47) (open symbols) C-terminus splice variants.  IV-curves for all constructs were determined from currents evoked during 90 ms square pulse depolarizations shown between -50 mV and +20 mV from a holding potential of -90 mV.  B, conductance values were calculated from IV curves to obtain activation curves.  C, steady-state inactivation curves were generated using a standard protocol in which 5s prepulse holdings of -100 to +10 mV were elicited prior to the 80 ms, 0 mV test pulse from a holding of -120 mV.  Normalized current evoked during the test pulse is plotted vs. prepulse membrane potential. For complete statistics see Table 1 and for details of protocols see Experimental Procedures.   80    V50 Activation (mV) k Activation τact (ms) V50 Inactivation (mV) k Inactivation Cav2.1 (Δ47) Wild-type -14.02 ± 1.49 (n = 16) 4.82 ± 0.27 1.42 ± 0.09 -58.20 ± 2.04 (n = 21) 7.03 ± 0.29 Cav2.1 (Δ47) K1336E -24.12 ± 1.33 # (n = 10) 2.80 ± 0.20 # 1.16 ± 0.21 -62.27 ± 1.81 (n = 7) 7.23 ± 0.45 Cav2.1 (Δ47) R192Q -20.84 ± 0.94 # (n = 16) 4.30 ± 0.19 1.31 ± 0.07 -73.41 ± 3.15 # (n = 11) 7.58 ± 0.57 Cav2.1 (Δ47) S218L -24.10 ± 1.15 # (n = 9) 5.53 ± 0.39 0.97 ± 0.11 * -75.07 ± 3.96 # (n = 10) 6.34 ± 0.62  Cav2.1 (+47) Wild-type -15.08 ± 1.20 (n = 16) 4.33 ± 0.24 1.28 ± 0.07 -62.07 ± 1.87 (n = 16) 6.85 ± 0.38 Cav2.1 (+47) K1336E -21.53 ± 1.35 * (n = 9) 3.10 ± 0.18* 1.33 ± 0.10 -65.02 ± 1.87 (n = 8) 8.39 ± 0.78 Cav2.1 (+47) R192Q -19.11 ± 1.11 * (n = 16) 4.79 ± 0.26 1.43 ± 0.10 -70.27 ± 2.36 * (n = 16) 7.19 ± 0.47 Cav2.1 (+47) S218L -18.08 ± 1.28 (n = 11) 6.26 ± 0.31 # 1.06 ± 0.08 * -72.54 ± 1.70 * (n = 9) 5.92 ± 0.37  * p<0.05, #p<0.001     Table 2.1:  Mean values for voltage-dependent activation and inactivation parameters. The voltage at which half of the channels are in the activated state (V50act) and inactivated state (V50inact), and the steepness of the curves for activation (kact) and inactivation (kinact) were obtained by fitting the data with the Boltzmann equation for the indicated number of cells in parentheses.  The kinetics of activation (τact) were obtained by fitting the maximum current trace from the IV curves with a single exponential. Asterisks (*) and number signs (#) indicate significant difference relative to wild-type with p-values less than either 0.05 or 0.001 (one-way ANOVA), respectively.  N.A. = not applicable.        81 2.2.3 FHM-1 mutations exhibit differential effects on recovery from inactivation of Cav2.1 splice variants  Analysis of wild-type Cav2.1 (∆47) and Cav2.1 (+47) variants showed different rates of recovery from inactivation for these Cav2.1 channel splice variants.  Wild-type Cav2.1 (∆47) channels exhibit faster rates of recovery (τ 1 = 0.669 ± 0.108 ms and τ 2 = 3.08 ± 0.614 ms) than the Cav2.1 (+47) variant channels (τ 1 = 0.760 ± 0.212 ms and τ 2 = 3.39 ± 0.835 ms).  As a result, Cav2.1 (∆47) channels show a significantly higher percentage of channels recovered at 7.5 s after inactivation relative to Cav2.1 (+47) channels (89.0 ± 1.8 vs. 77.3 ± 3.8 %, respectively; p<0.05; ANOVA) (Fig. 2.3 and Table 2.2). Examining the effects of FHM-1 mutations in the Cav2.1 (∆47) background, Figure 2.3 shows that the K1336E Cav2.1 (∆47) and R192Q Cav2.1 (∆47) variants exhibit a significant decrease in current recovered at 7.5 s relative to wild-type Cav2.1 (∆47) channels (70.4 ± 3.7% and 83.3 ± 2.3% vs. 89.0 ± 1.8 %, respectively; p<0.05; ANOVA).  Contrastingly, K1336E Cav2.1 (+47) and R192Q Cav2.1 (+47) channels showed increases in recovery relative to wild-type Cav2.1 (+47) channels (86.8 ± 1.7 and 87.0 ± 2.0 vs. 77.3 ± 3.8 %, respectively; p<0.05; ANOVA) (Fig. 2.3 and Table 2.2).  The S218L mutation was found to increase the rate of recovery in both splice variants, however, only the S218L Cav2.1 (+47) channels showed a significant increase in recovery relative to wild-type Cav2.1 (+47) channels at 7.5 s (94.7 ± 1.5 vs. 77.3 ± 3.8 %, respectively; p<0.001; ANOVA) (Fig. 2.3 and Table 2.2). Overall, in agreement with previous reports (Mullner et al., 2004; Tottene et al., 2005), we observed that both the K1336E and S218L mutations can cause significant changes to recovery from inactivation and show for the first time that the R192Q mutation also changes recovery from inactivation. Importantly, we also show that the quantitative effects of the FHM-1 mutations on channel function are dependant upon the nature of the Cav2.1 splice-variant.  We also note that the differential effects of the mutations resulted in significant changes to the functional distinction observed between the two wild-type channel variants; that is, while the wild-type Cav2.1 (∆47) channel variant recovered significantly faster than the wild-type Cav2.1 (+47) channel variant, the K1336E Cav2.1 (∆47) channels recovered significantly slower than K1336E Cav2.1 (+47) channels.  In addition, the R192Q and S218L mutations altered recovery  82 such that the R192Q Cav2.1 (∆47) and R192Q Cav2.1 (+47) channels and S218L Cav2.1 (∆47) and S218L Cav2.1 (+47) channels were not functionally distinct in this parameter (Fig. 2.3 and Table 2.2).    83    84      Figure 2.3: Wild-type and FHM-1 mutant Cav2.1 (Δ47) and Cav2.1 (+47) variants exhibit different rates of recovery from inactivation. A, graphs show percentage of the current recovered vs. time given to recover for all wild-type and FHM-1 mutated constructs.  Recovery from inactivation was examined for wild-type (WT; black squares), FHM-1 mutant K1336E (KE; grey circles), R192Q (RQ; grey triangles), and S218L (SL; grey diamond) in both the short Cav2.1 (Δ47) (filled symbols) and long Cav2.1 (+47) (open symbols) C-terminus splice variants.  B, shows a representative trace (capacitive currents removed for clarity) and the two pulse protocol used.  The protocol consists of a 2 second, 0 mV prepulse followed by a 50 ms, 0 mV test pulse elicited after inter- pulse intervals between 10 ms and 7.5 s.  Time constants were determined by fitting the average values for percent recovery with a single exponential or double exponential (values shown in Table 2).  C, bar graph shows percent recovery at 7.5 s for all wild-type and FHM-1 mutant clones studied.  Single asterisks and number signs indicate significant difference between mutant and wild-type of the same variant with p- values less than either 0.05 or 0.001 (one-way ANOVA), respectively.  Double asterisks indicate significant difference between the Δ47 and +47 variants containing the same sequence (i.e. wild-type or mutant) with p-value less than 0.05 (one-way ANOVA).      τ 1 (fast) (ms) τ 2 (slow) (ms) % recovery at 7.5 s Cav2.1 (Δ47)  Wild-type 0.669 ± 0.108 3.08 ± 0.614 89.0 ± 1.8       (n = 5) Cav2.1 (Δ47)  K1336E 2.89 ± 0.12 N.A. 70.4 ± 3.7 #      (n = 8) Cav2.1 (Δ47)  R192Q 0.918 ± 0.188 4.97 ± 2.65 83.3 ± 2.3 *    (n = 7) Cav2.1 (Δ47)  S218L 0.617 ± 0.095 2.71 ± 1.18 91.6 ± 0.8       (n = 5)  Cav2.1 (+47)  Wild-type 0.760 ±  0.212 3.39 ± 0.835 77.3 ± 3.8 **   (n = 7) Cav2.1 (+47)  K1336E 0.776 ± 0.149 4.34 ± 2.31 86.8 ± 1.7 *     (n = 5) Cav2.1 (+47)  R192Q 1.08 ± 0.200 5.78 ± 4.96 87.0 ± 2.0 *     (n = 6) Cav2.1 (+47)  S218L 0.500 ± 0.119 1.71 ± 0.593 94.7 ± 1.5  #      (n = 5) * p<0.05, #p<0.001  Table 2.2:  Time constant values and recovery from inactivation. Time constants were determined by fitting the average percent recovery with a double exponential for all constructs except the Cav2.1 (Δ47) K1336E which was best fit with a single exponential.  Percent recovery is a measure of the percentage of current evoked during the test pulse, given at 7.5 s after the prepulse, relative to the maximum current evoked during the prepulse.  Asterisks and number signs indicate significant difference between mutant and wild-type of the same variant with p-values less than either 0.05 or 0.001 (one-way ANOVA), respectively.  Double asterisks indicate significant difference between the Cav2.1 Δ47 and +47 variants containing the same sequence (i.e. wild-type or mutant) with p-value less than 0.05 (one-way ANOVA).  Number of cells recorded for each clone is indicated in parenthesis.  N.A. = not applicable.    85 2.2.4 FHM-1 mutations exhibit differential effects on inactivation of Cav2.1 splice variants during tonic depolarization  Wild-type Cav2.1 (+47) and Cav2.1 (∆47) variants exhibit functional differences with regard to accumulation of inactivation during short (3.5 ms) pulses to +5 mV applied at a frequency of 25 Hz (Fig. 2.4).  While Cav2.1 (∆47) variant channels showed 95 ± 1.4 % of current remaining at the end of 25 pulses, Cav2.1 (+47) variant channels had 90 ± 0.9 % (p < 0.05; ANOVA) (Fig. 2.4C). All three FHM-1 mutations examined significantly altered accumulation of inactivation (Fig. 2.4A); however, again the effects were contingent on the nature of the Cav2.1 variant in which mutations were expressed.  The K1336E Cav2.1 (∆47) and K1336E Cav2.1 (+47) channels had a significant increase in accumulation of inactivation and thus a lower percent of current remaining at the end of 25 pulses relative to wild-type Cav2.1 (∆47) and wild-type Cav2.1 (+47) (88 ± 2.2 % and 84 ± 1.7 % vs. 95 ± 1.4 and 90 ± 0.9 %, respectively; p < 0.05; ANOVA) (Fig. 2.4C).  The changes were such that the K1336E Cav2.1 (+47) and K1336E Cav2.1 (∆47) variants had similar current remaining at the end of the repetitive stimulation and thus lacked the clear functional distinction observed between the wild-type channel variants. In the context of the Cav2.1 (∆47) variant background the R192Q mutation caused a significant increase in accumulation of inactivation relative to wild-type Cav2.1 (∆47) (current remaining at the end of the 25 pulses = 90 ± 1.6% vs. 95 ± 1.4 %: p < 0.05; ANOVA).  In contrast, R192Q Cav2.1 (+47) channels were similar to wild-type Cav2.1 (+47) channels (92 ± 1.3% vs. 90 ± 0.9 %) (Fig. 2.4C).  As a result, similar to K1336E channels, the R192Q Cav2.1 (∆47) and R192Q Cav2.1 (+47) channel variants lacked the clear functional distinction observed between the wild-type variants for this property. The S218L mutation showed a large and significant increase in accumulation of inactivation and thus a lower percent of current remaining at the end of 25 pulses relative to both the wild-type Cav2.1 (∆47) and Cav2.1 (+47) channel variants (81 ± 2.5% and 77 ± 1.7 % vs. 95 ± 1.4 and 90 ± 0.9 %, respectively; p < 0.05; ANOVA).  Similar to that for R192Q and K1336E, a further overall effect of the  86 S218L mutation is to decrease the relative difference in current remaining observed between wild-type Cav2.1 (∆47) and Cav2.1 (+47) variant channels.    87   88      Figure 2.4: Wild-type and FHM-1 mutant Cav2.1 (Δ47) and Cav2.1 (+47) variants exhibit different current decay during 25 Hz tonic depolarizations. A, graphs show normalized current remaining vs. time after initial onset of depolarizations.  Current decay was measured for wild-type (WT; black squares), FHM-1 mutant K1336E (KE; grey circles), R192Q (RQ; grey triangles), and S218L (SL; grey diamond) in both the short Cav2.1 (Δ47) (filled symbols) and long Cav2.1 (+47) (open symbols) C-terminus splice variants.  B, to investigate current decay during repetitive stimulations we used 25 square pulses from a holding of -100 mV to a depolarizing potential of -5 mV for a duration of 3.4 ms. The test pulses were given at a rate of 25 Hz.  Representative current trace and pulse protocol indicated at bottom (capacitive currents were compensated using a P/4 protocol), with single current response enlarged.  C, bar graph shows the percent of current remaining at the end of 25 pulses for each clone.  Single asterisks and number signs indicate significant difference between mutant and wild- type of the same variant with p-values less than either 0.05 or 0.001 (one-way ANOVA), respectively. Double asterisks indicate significant difference between the Δ47 and +47 variants containing the same sequence (i.e. wild-type or mutant) with p-value less than 0.05 (one-way ANOVA). Number of cells recorded for WT Cav2.1 (Δ47) (n = 14), WT Cav2.1 (+47) (n = 13), KE Cav2.1 (Δ47) (n = 15), KE Cav2.1 (+47) (n = 18), RQ Cav2.1 (Δ47) (n = 17), RQ Cav2.1 (+47) (n = 15), SL Cav2.1 (Δ47) (n =15), SL Cav2.1 (+47) (n = 14).                 89 2.2.5 FHM-1 mutations exhibit differential effects on inactivation of Cav2.1 splice variants during bursts of depolarization  In addition to tonic depolarizations, neurons experience various frequencies of burst firing in which brief periods of tonic firing are interspersed with silent periods as the membrane potential drops below threshold (McCormick et al., 1985; Brumberg, 2002; Womack and Khodakhah, 2002; Fernandez et al., 2007; Shin et al., 2007).  During the tonic firing periods Cav2.1 channels will inactivate and during silent periods they will have the opportunity to recover from inactivation.  Based upon the above noted splice-variant changes in accumulation of inactivation and recovery from inactivation, we predicted that bursts of depolarization would also differentially affect wild-type and FHM-1 mutated Cav2.1 (+47) and Cav2.1 (∆47) variant channels. Figure 2.5 shows that wild-type Cav2.1 (+47) and Cav2.1 (∆47) variants exhibit significant differences in the amount of current remaining at the end of five 25 Hz bursts given at 3.5 Hz.  Current through the wild-type Cav2.1 (+47) variant decayed to 73 ± 3.4% by the end of the fifth burst while current through the wild-type Cav2.1 (∆47) variant decayed to 88 ± 3.6% (p < 0.05; ANOVA) (Fig. 2.5C).  The increased inactivation during the depolarizations and the slower recovery from inactivation of wild-type Cav2.1 (+47) variants discussed above likely contributed to the overall 15% decrease in current relative to the wild-type Cav2.1 (∆47) variant. Figure 2.5 shows that during burst firing the K1336E mutation in the Cav2.1 (∆47) variant background results in an overall lower percentage of current remaining at the end of five bursts (70 ± 4.9 % vs. 88 ± 3.6%, respectively; p < 0.05; ANOVA) (Fig. 2.5C), likely resulting from the increased accumulation of inactivation and slowed recovery from inactivation of K1336E Cav2.1 (∆47) channels relative to wild-type Cav2.1 (∆47) (see Figs. 2.3 and 2.4).  Contrastingly, the K1336E mutation in the Cav2.1 (+47) variant background did not show significant current decay relative to wild-type, likely due to the fact that although the K1336E Cav2.1 (+47) variant channels exhibit a small increase in accumulation of inactivation during tonic stimulation (Fig. 2.4), they also possess an increased rate of recovery from  90 inactivation (Fig. 2.3A).  We note that unlike wild-type channel variants, the K1336E Cav2.1 (+47) and K1336E Cav2.1 (∆47) variants did not differ significantly relative to one another   91   92      Figure 2.5: Wild-type and FHM-1 mutant Cav2.1 (Δ47) and Cav2.1 (+47) variants exhibit different current decay during bursts of depolarizations. A, graphs show normalized current remaining vs. time after initial onset of depolarizations.  Current decay was measured for wild-type (WT; black squares), FHM-1 mutant K1336E (KE; grey circles), R192Q (RQ; grey triangles), and S218L (SL; grey diamond) in both the short Cav2.1 (Δ47) (filled symbols) and long Cav2.1 (+47) (open symbols) C-terminus splice variants.  B, to investigate current decay during bursts of repetitive stimulations we used five bursts of 25 square pulses to -5 mV for 3.4 ms from a holding of -100 mV; bursts given at 290 ms intervals (3.5 burst firing).  Each burst contained 25 pulses at a rate of 25 Hz. Representative current trace indicated at bottom (capacitive currents were compensated using a P/4 protocol).  C, bar graph shows the percent of current remaining at the end of 6 s for each clone.  Single asterisks and number signs indicate significant difference between mutant and wild-type of the same variant with p-values less than either 0.05 or 0.001 (one-way ANOVA), respectively.  Double asterisks indicate significant difference between the Cav2.1 Δ47 and +47 variants containing the same sequence (i.e. wild-type or mutant) with p-value less than 0.05 (one-way ANOVA).  Number of cells recorded for WT Cav2.1 (Δ47) (n = 8), WT Cav2.1 (+47) (n = 7), KE Cav2.1 (Δ47) (n = 8), KE Cav2.1 (+47) (n = 10), RQ Cav2.1 (Δ47) (n = 9), RQ Cav2.1 (+47) (n = 7), SL Cav2.1 (Δ47) (n =8), SL Cav2.1 (+47) (n = 8).                 93 Similar to that for the K1336E mutation in the ∆47 background, examination of burst firing effects on the R192Q mutation showed an increase in current decay during burst firing relative to wild-type Cav2.1 (∆47) channels (70 ± 3.6% vs. 88 ± 3.6% current remaining, respectively; p < 0.05; ANOVA) (Fig. 2.5C). In contrast, the R192Q mutation in the +47 background resulted in a higher degree of current remaining compared to wild-type Cav2.1 (+47) channels (83 ± 2.7% vs. 73 ± 3.4% current remaining, respectively; p < 0.05; ANOVA).  This may reflect the fact that R192Q Cav2.1 (+47) channels exhibit reduced inactivation during tonic firing and increased recovery from inactivation (see Figs. 2.3 and 2.4).  We note that an overall effect concerning current decay during burst firing is for R192Q Cav2.1 (+47) channels to behave more similar to wild-type ∆47 channels and for R192Q Cav2.1 (∆47) channels to behave more similar to the wild-type +47 variant. Similar to that for the K1336E mutation, the S218L mutation only caused a significant current decay during the burst firing in the Cav2.1 (∆47) variant background (73 ± 4.0% vs. 88 ± 3.6% current remaining at the end of five bursts, respectively; p < 0.05; ANOVA).  Although the S218L mutation increased accumulation of inactivation substantially in both splice variants (see Fig. 2.4), S218L Cav2.1 (+47) channels had a larger increase in the rate of recovery from inactivation (Fig. 2.3) which likely slowed overall accumulation of inactivation during the burst firing.  The S218L Cav2.1 (+47) and S218L Cav2.1 (∆47) variants did not differ significantly relative to one another.       2.3 Discussion 2.3.1 FHM-1 mutations differentially affect biophysical properties of Cav2.1 splice variants We report here that FHM-1 missense mutations confer differential effects on the biophysical properties of the Cav2.1 (+47) and Cav2.1 (Δ47) channel splice variants.  Although the current-voltage relationships and steady-state properties of the two wild-type Cav2.1 splice variants are similar, all three FHM-1 mutations exhibited a greater hyperpolarizing shift when expressed in the Cav2.1 (Δ47) variant compared to the Cav2.1 (+47) variant (Fig. 2.2).  In addition, we show for the first time that wild-type  94 Cav2.1 (Δ47) and wild-type Cav2.1 (+47) variants have both different kinetics of recovery from inactivation and accumulation of inactivation during tonic depolarization that are likely relevant to the differential response of channel variants during burst of depolarization (Figs. 2.3-2.5).  It is known that Cav2.1 channels in different states possess alternative modes of gating that are reflected in biophysical properties at the whole cell current level (Fellin et al., 2004; Luvisetto et al., 2004).  Furthermore, it has been shown that alternative splicing in the EF-hand region of the Cav2.1 carboxyl terminus can shift gating modes (Chaudhuri et al., 2007).  It is therefore possible that wild-type Cav2.1 (Δ47) and wild-type Cav2.1 (+47) variants also have distinct gating modes that respond differently to FHM-1 mutations which are localized to voltage sensor regions (e.g. R192Q, S218L, K1336E).  Detailed single channel analyses would be required to fully explore this hypothesis.  2.3.2 Differential effects of FHM-1 mutations on Cav2.1 splice variants may contribute to localized phenotype Our findings provide the first suggestion for a potential role of Cav2.1 channel alternative splicing in FHM-1 pathophysiology and raise the notion that even though Cav2.1 channels are widely expressed in the central and peripheral nervous systems, point mutations can have greater or lesser functional affects on specific splice variants.  Although the mechanism of FHM-1 pathophysiology is not completely resolved, the current opinion is that the migraine usually initiates with aura due to cortical spreading depression (CSD) which leads to headache pain through activation of the trigeminovascular pain pathway (Bolay et al., 2002; Pietrobon, 2005).  In this regard, specific Cav2.1 variants within the cortex may have important roles in the onset of migraine attacks. We show that the Cav2.1 (Δ47) and Cav2.1 (+47) variants are both expressed in whole human cortex (Fig. 2.1B), and that the three FHM-1 mutations all cause a greater hyperpolarizing shift in the voltage-dependence of activation in Cav2.1 (Δ47) channels relative to that for Cav2.1 (+47) variant channels.  A hyperpolarizing shift in Cav2.1 channel activation has been suggested as an underlying mechanism of increased susceptibility to CSD and the initiation of migraine (van den Maagdenberg et al.,  95 2004; Pietrobon, 2007).  CSD begins within small domains of the cortex and propagates outward from a focal point.  Our data supports the notion that Cav2.1 splice variants with greater sensitivity to hyperpolarizing shifts in the voltage-dependence of activation (e.g., Cav2.1 (Δ47)) could result in cortical regions with greater susceptibility to CSD and migraine initiation.  Conversely, the effects of FHM-1 mutations on other Cav2.1 splice variants (e.g., Cav2.1 (+47)), expressed elsewhere in the cortex or other brain regions may be below the threshold to initiate CSD and/or other pathological effects.  Future exploration of the exact regional and cellular distributions of these and other Cav2.1 splice variants within the cortex and throughout the human brain using in situ hybridization and/or RT-PCR analyses will be necessary to fully understand the role of Cav2.1 splice variants in FHM-1 pathology.  2.3.3 Differential effects of FHM-1 mutations on Cav2.1 splice variants under different conditions may contribute to episodic nature of the phenotype Our results examining tonic and burst firing patterns also suggests the possibility of differential effects of FHM-1 mutations on Cav2.1 channel splice variants under different firing conditions.  This is most clearly seen with the S218L mutation in the Cav2.1 (+47) variant; during tonic depolarization current decay is significantly faster relative to wild-type, yet during burst firing the S218L Cav2.1 (+47) variant has similar current decay to wild-type channels after five bursts, likely due to rapid recovery from inactivation (see Figs. 2.4 and 2.5).  On the other hand, in Cav2.1 (Δ47) variant channels the S218L mutation has significant effects on current decay under both tonic and burst firing conditions. Interestingly, certain initiating factors of FHM-1 attacks such as emotional stress (Ducros et al., 2001) are known to alter neuronal firing patterns in the brain (Weiss and Simson, 1988; McEwen, 2007).  Although the exact firing conditions directly associated with precipitating factors of migraine are unknown, the episodic nature of the FHM-1 phenotype may in part be associated with changes in neuronal firing pattern and/or frequency that could be relevant to specific Cav2.1 splice variants expressed in localized brain regions.   96 2.3.4 The differential effects of mutations on Ca2+ channel function is likely multifaceted and important in all Ca2+ channelopathies It is likely that there exists a complex relationship between channel missense mutations and disease mechanism.  While we show alternative splicing at a single Cav2.1 splice-site can determine the functional impact of FHM-1 mutations, we recognize that across the entire brain many additional factors are likely to be involved in ultimately defining disease pathophysiology.  These likely include the expression of multiple splice Cav2.1 variants with distinct combinations of alternative splicing as well as the interaction with different auxiliary subunits (Mullner et al., 2004) and other structural and regulatory proteins.  Nonetheless, our results demonstrate the relevance of alternative splicing as an important factor in considering underlying disease molecular mechanisms and also the need for a comprehensive understanding of the splice-variant profile of Cav2.1 channels across brain regions and developmental stages as they might relate to FHM-1 pathology. While in the present chapter we show that individual FHM-1 mutations can have differential effects on the biophysical properties of the short and long Cav2.1 channel splice variants, we predict this phenomenon is likely relevant to both other FHM-1 mutations and Cav2.1 variants and also to other types of Cav channels and Ca2+ channelopathies.  Understanding the differential effects of channelopathy mutations on ion channel splice variants is likely to be important for interpreting results obtained in both heterologous and native systems, as well as for making inferences concerning disease mechanisms and phenotypes.  Mutations in the Cav1.1 L-type channel are associated with hypokalemic periodic paralysis, the Cav1.2 L-type with Timothy syndrome, the Cav1.4 L-type with incomplete X-linked congenital stationary night blindness and X-linked cone-rod dystrophy, and the Cav3.2 T-type with idiopathic generalized epilepsy and autism spectrum disorder (reviewed in (Adams and Snutch, 2007)).  Similar to FHM-1, many of these disorders exhibit phenotypes with episodic and/or developmentally specific attributes localized to a subset of regions or tissues that express the respective channels, and like Cav2.1, these channels also undergo alternative splicing that generates functionally distinct channel variants (reviewed in (Lipscombe et al., 2002; Lipscombe and Castiglioni, 2004)).  The identification of specific  97 Cav splice variants involved in disease pathophysiology may also provide the opportunity for targeted therapeutic approaches.  For example, while the Cav2.2 N-type channels have a central role in nociceptive signalling, distinct Cav2.2 splice variants are involved in the transmission of specific types of pain and has led to new strategies for splice-variant-specific targeting in pain therapy (Altier et al., 2007).       2.4 Experimental Procedures 2.4.1 Site-directed mutagenesis Standard PCR-based in vitro mutagenesis was performed using the Pfu Turbo DNA Polymerase (Stratagene, La Jolla, CA), 10 mM dNTPs (Invitrogen) and paired forward and reverse mutagenesis primers (Zoller and Smith, 1984).  The human Cav2.1 long (+47) (isoform 2) (NCBI accession number NM_023035.1) cloned in pcDNA 3.1 Zeo (+) was used as the source for the generation of the wild-type short human Cav2.1 (Δ47) (isoform 1) cDNA (the other known six splice sites are: ∆10A, 16+/17+, -VEA, -NP, EFa, 43+/44+).  Paired forward and reverse primers were designed to adhere to the C-terminus of the Cav2.1 isoform 2 at the exon 46/47 boundary nucleotide number 6784 and removed the GGCAG pentanucleotide sequence creating the premature stop in exon 47 (Cav2.1 (Δ47)).  Both Cav2.1 splice- variant cDNAs were used in site-directed mutagenesis reactions to generate human Cav2.1 K1336E, R192Q and S218L mutants in the short and long variants; paired forward and reverse primers were designed to convert codon 1336 from AAA to GAA, codon 192 from CGG to CAG, and codon 218 from TCG to TTA.  The integrity of all constructs generated through site-directed mutagenesis were verified by direct DNA sequencing.  2.4.2 Cell culture and transfection  Human embryonic kidney (HEK 293) cells were grown in standard Dulbecco’s modified Eagle’s medium supplemented with 10% fetal bovine serum (heat inactivated) and 50 U/ml penicillin-50 µg/ml streptomycin. Cells were incubated at 37oC in a humidified incubator with 95% atmosphere and 5% CO2  98 and grown to 8-15% confluency for transfection.  HEK 293 cells were transiently transfected with either wild-type human Cav2.1 (Δ47) or Cav2.1 (+47), or mutant K1336E Cav2.1 (Δ47), R192Q Cav2.1 (Δ47), S218L Cav2.1 (Δ47) or K1336E Cav2.1 (+47), R192Q Cav2.1 (+47), S218L Cav2.1 (+47) in combination with Ca2+ channel auxiliary subunits β4, α2δ1, and the CD8 marker plasmid in a 1:1:1:0.25 molar ratio using Lipofectamine (Invitrogen, La Jolla, CA).  To ensure accurate comparisons, transfections were performed at the same time and electrophysiological recordings alternated within the same day for all channel types.  2.4.3 Electrophysiological recordings  On the second day after transfection, macroscopic Ba2+ currents where recorded at room temperature using the whole-cell patch-clamp technique (Hamill et al., 1981).  The internal pipette solution used contained 105 mM CsCl, 25 mM TEACl, 1 mM CaCl2, 11 mM EGTA, 10 mM HEPES and 5 mM ATP (pH 7.2 with CsOH); external: 5 mM BaCl2, 1 mM MgCl2, 10 mM HEPES, 40 mM TEACl, 10 mM glucose and 87.5 mM CsCl (pH 7.4 with TEAOH).  Patch pipettes (borosilicate glass BF150-86-10; Sutter Instrument Company, Novato, CA) were made using a horizontal puller (P-87; Sutter Instruments Company) and fire polished using a microforge (Narishige, Tokyo, Japan), with resistances typically of 3 to 5 MΩ when containing internal solution.  External solution bath was connected to ground with a 3M KCl agar bridge.  Whole cell currents were recorded and filtered at 2-5 kHz bandwidth using an Axopatch 200A amplifier and monitored and stored on a personal computer running pClamp software package version 9.  Sampling frequencies were between 2 and 10 kHz.  Recordings were analyzed using Clampfit 9 and figures, fittings and statistics (ANOVA) were made using the software program Origin version 7.5 (OriginLab Corp., Northampton, MA).     99 2.4.4 Recording protocols and data analysis  Current-voltage relationships were determined by measured currents obtained using a series of 90 millisecond depolarization pulses applied from a holding potential of -90 mV to membrane potentials from -50 mV to +45 mV, increasing by 5 mV increments.  Current-voltage relationships were fitted, and IV curves generated, using a modified Boltzmann equation: I= (Gmax*(Vm-Er))/ (1+exp ((Vm-V50)/k)), where Gmax is the maximum slope conductance, Vm is the test potential, Er is the extrapolated reversal potential, V50 is the half-activation potential, and k reflects the slope of the activation curve.  Activation curves were constructed by calculating conductance from the IV curves and plotting the normalized conductance as a function of the membrane potential.  The data were fit with the Boltzmann equation: G/Gmax = A2 + (A1- A2)/ (1 + exp ((Vm-V50)/k)), where A1 is minimum normalized conductance, A2 is maximum normalized conductance, Vm is the test potential, V50 is the half-activation potential, and k reflects the slope of the activation curve (goodness of fit had R2 values ≥ 0.998). Voltage-dependence of inactivation was analyzed using depolarizations to 0 mV for 80 ms following 5 s prepulses ranging from -100 to +10 mV at 10 mV increments (holding potential of -120 mV). Steady state inactivation curves were constructed by plotting the maximum normalized current during the test pulse as a function of the prepulse potential.  The data were fit with the Boltzmann equation: I/Imax = A2 + (A1-A2)/ (1 + exp ((Vm-V50)/k)), where A1 is minimum normalized current, A2 is the maximum normalized current, Vm is the test potential, V50 is the half-inactivation potential, and k reflects the slope of the inactivation curve (goodness of fit had R2 values ≥ 0.998). The kinetics of activation (τact) were determined from currents obtained from the IV protocol. Current traces were fit with a standard single exponential equation: I = A*exp (-t/τ), where A is the amplitude of the current, and τ is the time constant.  Recovery from inactivation was determined using a double-pulse protocol.  The first depolarization was to 0 mV for 2 s (the prepulse), followed by a return to the holding potential of -100 mV for variable lengths between 10 ms and 7.5 s.  At the end of the variable repolarization period, a second 0 mV (the test pulse) was given for 50 ms.  The time interval between sweeps was a total of 1 minute to  100 ensure maximum recovery between sweeps.  All traces were normalized to the maximum current during the prepulse for each sweep.  The peak current from the test pulse was plotted as a percentage of maximum prepulse current vs. repolarization time.  Average traces were fit with either a single or double exponential equation (goodness of fit had R2 values ≥ 0.998). Current decay during a tonic depolarization was examined using a 25 Hz train of 25 square pulses from a holding of -100 mV to a depolarizing potential of -5 mV for 3.4 ms.  Current decay curves were generated by plotting normalized maximum current during the test pulses as a function of the time of pulse onset.  Current decay during bursts of depolarization was examined using square pulses to -5 mV for 3.4 ms from a holding of -100 mV.  Five bursts were given with 290 ms intervals (3.5 Hz burst firing).  Each burst contained 25 pulses at a rate of 25 Hz.  Current decay curves were generated by plotting normalized maximum current during the test pulses as a function of the time of pulse onset.  2.4.5 RT-PCR of Cav2.1 carboxyl-terminal region from human cortex RNA  Prior to reverse transcription, 1 µg total RNA from human cortex (Clontech; 636561) was treated with 1X DNase I reaction buffer and 1 unit DNase I (Invitrogen) in a final volume adjusted to 10 µL using sterile DEPC-treated H2O.  Following a 15 minute incubation period at room temperature, the reaction was inactivated by adding 1 µL of 25 mM EDTA and heating at 65ºC for 10 minutes.  cDNA synthesis was performed using Superscript II Reverse Transcriptase (Invitrogen) following manufacturer’s instructions with slight modification.  A ~1.1Kb nucleotide fragment of the carboxyl end of the Cav2.1 channel was amplified from the human cortex cDNA using standard PCR.  The reaction mixture consisted of 3% DMSO, 1X Phusion enzyme buffer, 0.4 pmol/µL of forward and reverse primers, 0.2 mM dNTPs, 1 µL of cortex cDNA, and 1 unit of Phusion enzyme in a final volume of 25 µL.  The forward primer (5’GGCACATGGAGTCCGGAACA 3’) corresponds to nucleotide position 6130 and the reverse primer (5’GGTAGTAGCCATGGTGCC3’) to nucleotide position 7211 of the human Cav2.1 α1 subunit (NCBI accession number NM_023035.1).  The cycling profile included an initial activation step of 98oC for 30s followed by 30 cycles of 98oC for 30s, 65oC for 30s, 72oC for 2.5 minutes and a final extension period of  101 72oC for 10 minutes.  As positive controls, the same PCR reaction was performed on human Cav2.1 (Δ47) and Cav2.1 (+47) cDNA plasmids to demonstrate unbiased amplification. PCR reactions using primers for tubulin were used to verify the success of RT reactions.  As negative controls, identical PCR reactions without template and containing no RT were performed.  To analyze individual Cav2.1 amplified carboxyl terminal transcripts, PCR products were ligated into pGEMT-Easy (Promega) and then transformed into competent XL-1 E.coli bacterial cells.  Bacteria containing PCR inserts were identified by blue-white screening and collected and subject to Cav2.1 (Δ47) and Cav2.1 (+47) specific PCR reactions.  The reverse primer (5’GGTAGTAGCCATGGTGCC3’) was used for both Cav2.1 (Δ47) and Cav2.1 (+47) specific PCR reactions.  Forward primer (5’ATGGCGCACCGGCAGTA3’) and (5’CATGGCGCACCGGCAGGG3’) were designed to specifically amplify Cav2.1 (Δ47) and Cav2.1 (+47), respectively.  All PCR products were run on a 1% agarose gel.  In determining the percentage of each variants, only colonies positive for Cav2.1 (Δ47) and negative for Cav2.1 (+47), and visa versa, were included and ambiguous results discarded. Direct DNA sequence determination of several representative clones confirmed both the veracity of the splice-variant specific PCR reactions and the identity of the cloned PCR products.       2.5 Acknowledgments  This work was funded by operating grant #10677 from the Canadian Institutes of Health Research (CIHR) to T.P. Snutch, a Tier 1 Canada Research Chair in Biotechnology and Genomics-Neurobiology to T.P. Snutch, an operating grant from the National Ataxia Foundation and a salary award from the Vancouver Coastal Health Authority to S.D. Spacey, graduate fellowships from the Michael Smith Foundation for Health Research to P.J. Adams and K.J. Mulatz, and a doctoral fellowship from the Heart and Stroke Foundation of Canada to L.S. David.  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DNA 3:479-488.    106  3 CONTRIBUTION OF Ca2+-DEPENDENT FACILITATION TO SHORT- TERM SYNAPTIC PLASTICITY REVEALED BY CONGENITAL MIGRAINE MUTATIONS IN THE Cav2.1 Ca2+ CHANNEL*       3.1 Introduction Short-term facilitation of synaptic release has historically been attributed to enhanced vesicle release resulting from the accumulation of intracellular Ca2+ in presynaptic terminals during repetitive APs, in which the build-up of residual Ca2+ enhances binding to CaS proteins that directly mediate vesicle fusion and transmitter release (Zucker and Regehr, 2002).  However, there is some evidence that residual Ca2+ can interact with other CaSs independent of the release machinery to enhance presynaptic Ca2+ currents and contribute towards synaptic facilitation, although the exact underlying mechanism remains uncertain (Forsythe, 1994; Borst and Sakmann, 1998; Cuttle et al., 1998; Forsythe et al., 1998; Tsujimoto et al., 2002; Rousset et al., 2003; Inchauspe et al., 2004; Muller et al., 2008).  In a recent study using a model system, Mochida and collaborators demonstrated that CDF and CDI of Cav2.1 channels via CaS protein interaction with the IQ-like and CBD domains in the Cav2.1 C-terminus can be a mechanism to induce short-term synaptic facilitation and rapid synaptic depression, respectively (Mochida et al., 2008).  Whether similar forms of CDF and/or CDI of Cav2.1 channels represent crucial mechanisms of synaptic plasticity at intact central synapses has not been reported.   ________________________________ * A version of this chapter has been accepted for publication. Adams, P.J., Rungta, R.L., Garcia, E., van den Maagdenberg, A.M.J.M, MacVicar, B.A., Snutch, T.P. (2010) Contribution of calcium-dependent facilitation to short-term synaptic plasticity revealed by congenital migraine mutations in the P/Q-type calcium channel. PNAS, in press.  107 In the current study, we explored the effects of FHM-1 missense mutations on CDF and CDI of the Cav2.1 channel.  FHM-1 is a congenital, autosomal dominant form of migraine with aura, typified by hemiparesis and linked to missense mutations in the CACNA1A gene encoding the Cav2.1 α1-subunit that conducts P/Q-type Ca2+ currents (Ophoff et al., 1996).  Biophysically, FHM-1 missense mutations result in an overall gain-of-function Cav2.1 channel phenotype as a result of an underlying shift in channel gating allowing increased Ca2+ influx at lower membrane potentials (Hans et al., 1999; Tottene et al., 2002).  CDF and CDI are robust forms of Cav2.1 channel modulation mediated by calmodulin and other Ca2+ sensor proteins which interact with the Cav2.1 carboxyl terminus in bipartite regulatory processes; CDF is mediated by a local rise in Ca2+ and CDI through a global rise in Ca2+ (Lee et al., 1999; Lee et al., 2000; DeMaria et al., 2001; Lee et al., 2002; Tsujimoto et al., 2002; Lee et al., 2003; Chaudhuri et al., 2004; Chaudhuri et al., 2005; Few et al., 2005; Chaudhuri et al., 2007).  The underlying mechanisms of CaM mediated CDF is also attributed to changes in channel gating (Chaudhuri et al., 2007) and it was of interest to examine whether FHM-1 mutations affect these important modulatory properties of Cav2.1 channels and to explore physiological implications using transgenic models. We found that FHM-1 gain-of-function missense mutations significantly occlude CDF in both recombinant and native systems and correlate with a reduction in short-term synaptic facilitation. Collectively, the data provide the first evidence that FHM-1 mutations directly affect the Ca2+-dependent regulation of Cav2.1 channels and support the notion that selective Ca2+-dependent regulation of presynaptic Cav2.1 channels may underlie several key aspects of short-term plasticity at the PF-PC synapse in cerebellum.       3.2 Results 3.2.1 FHM-1 mutations occlude CDF and CDI of recombinant human Cav2.1 channels CaM mediated CDF and CDI of Cav2.1 channels have been well characterized using human recombinant Cav2.1 channels expressed in HEK cells (along with auxiliary subunits β2a and α2δ1) and  108 allows for clear isolation and measurement of CDF and CDI (Lee et al., 1999; DeMaria et al., 2001). Consistent with previous findings, human recombinant wild-type Cav2.1 channels transiently expressed in HEK cells showed typical CDF with prepulse dependent relative facilitation when Ca2+ was used as the charge carrier (Fig. 3.1A, black squares) (all clones are the Cav2.1 (Δ47) splice-variant).  Currents elicited by a test pulse showed a rapid initial activation with a normal gating mode and then slowly transitioned to a facilitated mode as Ca2+ entered (Fig. 3.1E, grey traces).  In contrast, an applied prepulse evoked Ca2+ entry that rendered channels in the facilitated mode of gating such that currents evoked by an ensuing test pulse lacked the slower component and displayed instead a fast activation rate (Fig. 3.1E, black traces) (facilitated mode being an enhancement of Po over the basal level in the normal gating mode (Chaudhuri et al., 2007)).  When Ba2+ was used as the charge carrier, channels opened directly into the normal gating mode and did not transition to the facilitated state, regardless of the prepulse potential (Fig. 3.1A; grey symbols).  A measure of “pure” CDF was calculated by subtracting relative facilitation determined using Ca2+ as the charge carrier from relative facilitation when using Ba2+ (Fig. 3.1A; denoted g) (Chaudhuri et al., 2004).  The FHM-1 mutations, R192Q and S218L, represent distinct ends of the clinical spectrum with patients possessing the R192Q alteration exhibiting a mild, pure FHM-1 without other neurological symptoms (Ophoff et al., 1996; Ducros et al., 2001), whereas patients with the S218L mutation have an associated severe clinical migraine phenotype most often associated with ataxia or cerebellar symptoms (Fitzsimons and Wolfenden, 1985; Kors et al., 2001; Chan et al., 2008).  Figure 3.1 shows that channels containing either R192Q (Fig. 3.1A; triangles) or S218L (Fig. 3.1A; diamonds) mutations precluded CDF across multiple prepulse potentials, and overall significantly reduced pure CDF relative to wild-type channels (Fig. 3.1B).  109   110     Figure 3.1:  The R192Q and S218L FHM-1 mutations occlude Ca2+-dependent modulation of human recombinant Cav2.1 channels. A, Using the paired-pulse protocol previously described (Chaudhuri et al., 2004), we show both the R192Q and S218L FHM-1 mutations reduced CDF across several prepulse potentials shown as relative facilitation vs. prepulse potential; results are means ± s.e.m.  B, Pure CDF (the difference between relative facilitation obtained using 5 mM Ca2+ as the charge carrier minus relative facilitation obtained using 5 mM Ba2+; g) (Chaudhuri et al., 2004) following a +20 mV prepulse compared to wild-type (g = 0.290±0.046) was significantly reduced by the R192Q (g = 0.097±0.042) and S218L (g = 0.023±0.031) mutations.  C, Using 1 s square test-pulses between -10 and +30 mV, we show current remaining at 800 ms (r800) across several prepulse potentials was reduced by both FHM-1 mutations; results are means ± s.e.m.  D, Comparison of pure CDI (the difference between r800 values obtained using 5 mM Ca2+ as the charge carrier minus r800 obtained using 5 mM Ba2+; f) shows that relative to wild-type (f = 0.418±0.067), the reduction by R192Q was modest (f = 0.214±0.082) although S218L caused a significant reduction (f = 0.083±0.038).  E, Paired-pulse protocol used, and representative traces for peak CDF obtained following a +20 mV prepulse for wild-type, R192Q and S218L Cav2.1 channels using Ca2+ (top) and Ba2+ (bottom) as charge carrier; traces normalized to the end of the test pulse.  F, Representative traces in which traces are normalized to the peak of a +10 mV test pulse.  G, APW used and representative traces for wild-type, R192Q and S218L Cav2.1 channels.  First hashed line represents the level of Ca2+ response during the first action potential, and the second line represents the peak (maximum facilitation).  100 Hz APW used was previously derived from action potentials recorded in the calyx of Held (Borst and Sakmann, 1998; Patil et al., 1998).  n refers to the number of cells recorded. All statistics were performed using a one-way ANOVA.  Asterix denotes p<0.05.           111 The effect of FHM-1 mutations on CDI of exogenous Cav2.1 channels was tested using a 1 s test pulse to varying potentials in Ca2+ and Ba2+.  Wild-type Cav2.1 channels showed a typical CDI characterized by faster inactivation when Ca2+ was used as the charge carrier (Fig. 3.1C; black squares) relative to Ba2+ (Fig. 3.1C; grey, open squares) (Fig. 3.1F; left panel) (DeMaria et al., 2001; Chaudhuri et al., 2005).  A measure of pure CDI was determined by subtracting currents obtained using Ca2+ as the charge carrier from currents obtained using Ba2+ (Fig. 3.1C; denoted f, and Fig. 3.1D) (Chaudhuri et al., 2004).  Although the R192Q mutation modestly reduced CDI relative to that of wild-type (Fig. 3.1C; triangles), the more severe S218L mutation (Fig. 3.1C; diamonds) completely removed the ability of channels to undergo CDI (Fig. 3.1D and F, right panel).  Importantly, because of its reliance on global Ca2+ levels (unlike that for CDF), CDI depends on Cav2.1 current density (Soong et al., 2002). While the R192Q mutation has little affect on current density, the S218L mutation causes a large reduction in current density (Tottene et al., 2005) (and see Fig. 3.2 for similar findings). As such, at the mechanistic level the reduced ability of mutant channels to undergo CDI is likely, at least in part, a result of reduced current density. While rectangular depolarizations allow for optimal biophysical resolution of CDF and CDI (Fig. 3.1), it was also important to test the effects of FHM-1 mutations under conditions more resembling neuronal firing, such as APWs (DeMaria et al., 2001).  In response to application of a 100 Hz APW, wild-type Cav2.1 channels displayed typical CDF and CDI that shaped the Ca2+ currents; CDF caused an initial facilitation of Ca2+ currents during the first few APs, and CDI caused a cumulative reduction as the APWs continued to be applied (Fig. 3.1G) (Chaudhuri et al., 2004; Chaudhuri et al., 2007).  The FHM-1 R192Q and S218L mutations both strongly suppressed the dynamics of the response to APWs, consistent with their effects on CDF and CDI described above (Fig. 3.1G) (see Fig. 3.2 for average responses and responses obtained using Ba2+ as the charge carrier).  112  Figure 3.2: The effects of the R192Q and S218L mutations on current density and on CDF and CDI during 100 Hz APWs in Ca2+ and Ba2+ A, Current density (CD) for wild-type, R192Q and S218L human recombinant Cav2.1 channels. B, 100 Hz APW derived from APs recorded in the calyx of Held (Borst and Sakmann, 1998; Patil et al., 1998). C, Representative traces of transiently transfected human recombinant Cav2.1 channel responses when Ca2+ or Ba2+ were used as the charge carrier. D, Average responses obtained for wild-type, R192Q and S218L Cav2.1 channels. n refers to the number of cells recorded.        113 As the biophysical effects of FHM-1 mutations can be affected by a number of factors including Cav2.1 splice-variation (Adams et al., 2009), β subunit coexpression (Mullner et al., 2004) and the nature of the expression system (Kraus et al., 1998; Hans et al., 1999), it was important to test the FHM-1 mutations in the context of endogenous Cav2.1 channel modulation.  In this regard, we measured P/Q- type currents through endogenous Cav2.1 channels in acutely dissociated cerebellar PCs from wild-type and R192Q and S218L knock-in mice (van den Maagdenberg et al., 2004; Eikermann-Haerter et al., 2009; Tottene et al., 2009).  3.2.2 FHM-1 mutations occlude CDF of native P/Q-type currents in cerebellar PCs P/Q-type currents are robustly expressed in dissociated cerebellar PCs (see refs (Mintz et al., 1992a; Mintz et al., 1992b) and Fig. 3.3C) and recapitulate the key features of CDF currents observed for human recombinant Cav2.1 channels, while CDI is highly variable (Chaudhuri et al., 2005) (and our own unpublished observations).  We examined CDF of endogenous P/Q-type currents in acutely dissociated cerebellar PCs from wild-type and homozygous R192Q and S218L knock-in mouse strains (van den Maagdenberg et al., 2004; Eikermann-Haerter et al., 2009; Tottene et al., 2009).  P/Q-type currents in PCs from wild-type mice showed similar CDF to currents through wild-type human recombinant Cav2.1 channels (Fig. 3.3).  In contrast, P/Q-type currents from dissociated PCs from R192Q and S218L knock- in mice exhibited reduced CDF both across multiple prepulse potentials (Fig. 3.3A) and during APWs (Fig. 3.3D); with the more severe S218L mutation resulting in a statistically significant reduction in pure CDF (Fig. 3.3B).  Taken together, the findings from both recombinant and endogenous Cav2.1 channels support the notion that the FHM-1 R192Q and S218L mutations reduce or even preclude CDF of Cav2.1 channels.  The effects on Cav2.1 CDF predict that FHM-1 mutations alter Cav2.1 channel-dependent functions as they relate to synaptic signalling.   114    115   Figure 3.3:  CDF is similarly occluded in endogenous P/Q-type currents in acutely dissociated PCs from Cav2.1 R192Q and S218L knock-in mice. A, Both the R192Q and S218L mutations reduced CDF across several prepulse potentials; results are means ± s.e.m (representative traces below).  B, Pure CDF (g) relative to wild-type (g = 0.210 ± 0.028) was not significantly reduced by R192Q (g = 0.136 ± 0.03) while the S218L mutation resulted in a significant reduction (g = 0.0686 ± 0.035).  C, P/Q-type currents were isolated from freshly dissociated cerebellar PCs from 15-25 day old mice and identified by their characteristic large size and tear-shaped morphology.  Exemplar trace below shows that the pharmacologically isolated P/Q-type currents were completely blocked with 0.2 μM ω-Aga-IVA.  D, Action potential waveform used and representative traces for wild-type, R192Q and S218L Cav2.1 channels.  First hashed line represents the level of Ca2+ response during the first action potential, and the second line represents the peak (maximum facilitation). n refers to the number of cells recorded.  All statistics were performed using a one-way ANOVA. Asterix denotes p<0.05.               116 Neither a contribution of Cav2.1 CDF towards synaptic plasticity in cerebellum nor demonstration as to whether FHM-1 mutations might affect fast synaptic cerebellar signalling has been reported.  The PF-PC synapse is a well-characterized central synapse that displays robust presynaptic forms of short-term facilitation via both paired-pulse and AP trains, relies predominantly on Cav2.1 channels for neurotransmitter release (Mintz et al., 1995; Randall and Tsien, 1995) and facilitation at this synapse is mediated by CaSs similar to those reported to mediate Cav2.1 CDF in recombinant and native systems (Atluri and Regehr, 1996; Lee et al., 1999; Lee et al., 2000; DeMaria et al., 2001; Lee et al., 2002; Tsujimoto et al., 2002; Lee et al., 2003; Chaudhuri et al., 2004; Chaudhuri et al., 2005; Few et al., 2005; Chaudhuri et al., 2007).  While the specific modulators of facilitation at the presynaptic terminals of PFs have not been defined, we hypothesized that CDF of Cav2.1 channels is an important means of short-term synaptic facilitation and further, that FHM-1 mutations affecting CDF would have a corresponding effect on synaptic plasticity.  3.2.3 FHM-1 mutations occlude short-term synaptic facilitation at the PF-PC synapse Synaptic transmission at the PF-PC synapse was measured using extracellular field recordings in transverse slices from wild-type and homozygous R192Q and S218L knock-in mice.  A typical facilitation response was evoked by paired, 180 μs stimulations of PFs (50 ms interpulse interval), in which the second EPSP was facilitated relative to the first (termed paired-pulse facilitation: PPF) (Mintz et al., 1995) (Fig. 3.4A; left, representative trace).  Of note, the paired pulse ratio (size of the second pulse relative to the first; PPR) was significantly reduced by both the R192Q and S218L mutations (Fig. 3.4A; bar graph), and in a manner quantitatively consistent with the reduction in Cav2.1 CDF in exogenous and native systems (compare with Fig. 3.1B and Fig. 3.3B).  Following five, 180 μs stimulations at 20 Hz, similar results were observed in that both the R192Q and S218L mutations significantly decreased the successive EPSPs relative to the second pulse (Fig. 3.4B).  These results support the notion that CDF of Cav2.1 channels is a contributing and necessary component of synaptic plasticity in presynaptic terminals of PFs.  An unresolved question is whether FHM-1 mutant presynaptic  117 channels are reluctant to facilitate and have a reduced Ca2+ influx and decreased transmitter release, or whether they predominate within a facilitated state with enhanced Po that increases presynaptic Ca2+ influx and transmitter release.   Figure 3.4:  Occlusion of Cav2.1 CDF results in a comparable attenuation in PPF at the PF-PC synapse. A, Left, exemplar field recording of synaptic responses from the PC layer evoked by two extracellular stimuli of ~15V, 180 μs delivered at a 50 ms interval to PFs in the molecular layer (wild-type mouse). The paired-pulse ratio (PPR) is a quantification of facilitation obtained by dividing response two by response one. Right, PPR was significantly reduced relative to wild-type (2.36±0.096) by both the R192Q (1.86±0.13) and S218L mutations (1.60±0.075); results are means ± s.e.m.  B, Left, exemplar field recording of synaptic responses from the PC layer evoked by five extracellular stimuli of ~15V, 180 μs delivered at 20 Hz (wild-type mouse).  Right, relative facilitation was measured by dividing the peak response from each stimulus by the response obtained from the first stimulus, plotted vs. pulse number; results are means ± s.e.m.  Both mutations reduced relative facilitation during 5 pulses at 20 Hz.  n refers to the number of slices recorded (from 8 wild-type, 4 R192Q and 5 S218L mice).  All statistics were performed using a one-way ANOVA.  Asterix denotes p<0.05.   118 Several lines of evidence suggest that the FHM-1 mutant channels favour a facilitated state that precludes further CDF.  For one, the facilitated gating mode of Cav2.1 channels induced by CaM mediated CDF is an increase in Po predicted to both enhance and elongate AP induced Ca2+ currents through the channels (Chaudhuri et al., 2007), and critically, both the R192Q and S218L mutations can render channels in a state of increased Po in some systems (Hans et al., 1999; Tottene et al., 2005) similar to that described for facilitated channels (Chaudhuri et al., 2007). Furthermore, the kinetics of mutant channels suggest a predominant gating mode consistent with a facilitated state (Fig. 3.5).  Without a prepulse, wild-type channels open with a normal gating mode and slowly transition to a facilitated mode of gating as Ca2+ enters (Fig. 3.5A; -1- and -2-).  When paired with a prepulse, channels open rapidly with the facilitated mode of gating (Fig 3.5A; -3-).  Conversely, in the presence of an FHM-1 mutation, channels appear to activate rapidly to the maximum current level regardless of prepulse (Fig 3.5B; -3-).  These support a kinetic model whereby FHM-1 mutations favour a functional state with enhanced Po that shifts the equilibrium of channels toward a gating mode that is the same as (or analogous to) the one achieved during CDF (Fig 3.5A and B; lower panels). Lastly, the results in Figure 3.4B show that compared to transmission measured in wild-type animals, synapses from both the R192Q and S218L mice exhibit decreased PPF and accelerated decay of EPSPs with successive stimulations.  Both effects can be explained by mutant channels being in a facilitated state whereby greater Ca2+ influx through facilitated presynaptic Cav2.1 channels during initial APs results in vesicle depletion and thus a decrease in successive synaptic responses. Taken together, we predicted that the observed changes in synaptic plasticity at the PF-PC synapse result from a larger initial Ca2+ influx through basally facilitated mutant channels relative to unfacilitated wild-type channels in PF boutons.     119   120   Figure 3.5:  Kinetics of macroscopic currents of mutant channels are consistent with a preferentially facilitated gating mode. A, upper panel; when Ca2+ is used as the charge carrier, wild-type Cav2.1 channels respond to a +5 mV square test pulse with an initial rapid activation (-1-) followed by a slow phase of current increase as Ca2+ enters through the channel (-2-).  On the other hand, when preceded by a +20 mV prepulse, wild- type channels activate rapidly with a maximum current response (-3-) (also see ref (DeMaria et al., 2001)).  Lower panel; mechanistically this means that without a prepulse wild-type channels open with a normal gating mode (-1-) and slowly transition to a facilitated mode of gating as Ca2+ enters (-2-)(i.e. CDF).  As Ca2+ enters it binds CaM and the Ca2+/CaM complex enhances Po and shifts the equilibrium of channels to a facilitated mode of gating (see refs (DeMaria et al., 2001; Chaudhuri et al., 2007) for model and single channel analysis showing the mechanism is a true enhancement in Po).  Following a prepulse, channels open rapidly with the facilitated mode of gating (-3-).  B, upper panel; conversely, in the presence of the S218L mutation, channels appear to activate rapidly to the maximum current level (-3-) regardless of prepulse (similar results obtained for the R192Q mutation; data not shown).  Lower panel; a kinetic model whereby FHM-1 mutations cause an enhancement of Po that shifts the channel equilibrium toward a gating mode that resembles the one achieved during CDF.  At the macroscopic level, enhanced Po of Cav2.1 will result in faster activation kinetics.  C, In fact, we observe that both the S218L and R192Q mutations cause an increase in Cav2.1 activation kinetics across several membrane potentials measured at the macroscopic level during 90 ms square pulses from 0 to +25 mV, from a holding of -90 mV.  All representative traces and kinetics of activation recordings were measured using human recombinant channels expressed in HEK cells.  The tau activation was determined by fitting current traces with a single exponential.           121 3.2.4 FHM-1 mutant Cav2.1 channels appear to be in a basally facilitated state in PF boutons The geometry and spatial equilibration of Ca2+ in the PF boutons is ideal for measuring the role of Ca2+ dynamics on a tens-of-millisecond time scale using Ca2+-sensitive fluorescent dyes (Mintz et al., 1995; Atluri and Regehr, 1996).  To this end, we evoked a train of 5 PF responses (50 μs pulse durations at 20 Hz) in either wild-type or homozygous S218L mice and simultaneously monitored the fluorescent response of the Ca2+ indicator Rhod-2 in presynaptic boutons using two-photon microscopy in line scan mode.  We chose to look at the S218L mice because of the consistently larger effect on CDF.  Figure 3.6A shows an example of unstimulated (top) and stimulated (bottom) PFs in which the presynaptic boutons were detected as relatively bright regions (indicated by hashed line in stimulated image).  Ca2+ influx in boutons from S218L mice (Fig. 3.6B; grey diamonds) was clearly enhanced during the train of 5 PF responses compared to those in boutons of wild-type mice (Fig. 3.6B; black squares).  Additionally, the amplitudes of Ca2+ transients in the presynaptic boutons in S218L mice (Fig. 3.6C; pulse 1) were similar to those expected from Ca2+ currents through facilitated Cav2.1 channels (Fig. 3.6D).  The results represent a true gain in Cav2.1 channel function as there was no apparent compensation by other Ca2+ channel subtypes at these boutons in the S218L mice (Fig. 3.7).          122   123      Figure 3.6:  The kinetics of Ca2+ influx in S218L presynaptic terminals indicate facilitated Cav2.1 channels. A, PFs were stimulated with a silver wire electrode and two photon imaging was used to measure Ca2+ transients in presynaptic terminals of the PFs; post-synaptic activity was blocked with 100 μM MCPG, 20 μM CNQX and 50 μM APV. Line scans were used to measure individual terminals (representative terminal indicated by hashed line). Four sets of five 50 μs stimuli at 20 Hz were given and the Ca2+ transients were averaged for each terminal. B, The average Ca2+ influx at PF terminals (normalized to the background fluorescence) is enhanced in S218L mice relative to wild-type littermates; results are means ± s.e.m (arrows represent the five stimuli). A representative average Ca2+ transient response measured for one terminal in linescan mode is shown in the lower panel. C, An expanded scale of B to show Ca2+ transient elicited during the first AP. Ca2+ transients in mutant terminals are larger in magnitude than wild-type terminals (enhanced ~15% relative to wild-type), as expected if mutant channels are in a basally facilitated gating mode. Stimulus is shown by AP illustration. n refers to the number of presynaptic terminals recorded (from 5 wild-type and 5 S218L mice). D, Expanded view of CDF of recombinant channels during APW from Fig. 3.1 Ca2+ currents through wild-type recombinant Cav2.1 channels increases as Ca2+ influx causes CDF (Left panel), whereas recombinant Cav2.1 channels containing the S218L mutation have a maximal Ca2+ response regardless of APs (Right panel). Bottom panel, Ca2+ currents through facilitated P/Q-type channels (grey trace) are larger than through unfacilitated channels (black trace) during evoked APs (10-25% increase in Ca2+ current amplitude was the range obtained from recombinant Cav2.1 channels in HEK cells (Fig. 3.1) and endogenous P/Q-type currents in PCs (Fig. 3.3). The unfacilitated trace is the Ca2+ response of a wild-type channel obtained during the first AP, and the facilitated trace is the Ca2+ response obtained during the 10th AP (indicated by arrows in top left panel).          124  Figure 3.7:  Ca2+ transients in wild-type and S218L mice are identical in the presence of 500 nM ω- Aga-IVA. A, PFs were stimulated with a silver wire electrode and two photon imaging was used to measure Ca2+ transients in presynaptic terminals of the PFs.  Line scans were used to measure individual terminals. Four sets of five 50 μs stimuli at 20 Hz were given and the Ca2+ transients measured.  Plots are of average Ca2+ influx at PF terminals (normalized to the background fluorescence); results are means ± s.e.m.  B, is an expanded scale of A. to show Ca2+ transient elicited during the first of the five pulses.  In the presence of 500 nM concentration of the potent Cav2.1 channel specific blocker ω-Aga-IVA (saturating concentration for these terminals (Mintz et al., 1995)), the wild-type (open black squares) and S218L (open grey diamonds) terminals had identical presynaptic Ca2+ transients.  These results demonstrate that the gain-of-function effect of the S218L mutations is a true gain of Cav2.1 channel function and not channel compensation by N-type channels known to be present at this terminal (Mintz et al., 1995).  Blocked terminals where measured in slices incubated for 40 minutes in ACSF containing 500 nM ω-Aga-IVA at room temperature with continuous perfusion of 95% O2.  Slices from the same animal were incubated in parallel in ACSF without ω-Aga-IVA and measured on the same day. Experiments were performed on 23 and 20 terminals, from 2 animals each for wild-type and S218L, respectively.        125       3.3 Discussion 3.3.1 A role for Cav2.1 CDF in short-term synaptic plasticity at the PF-PC synapse It has been known for some time that there exist at least two mechanisms of facilitation at the PF-PC synapse: a well understood mechanism described by the residual Ca  hypothesis and an incompletely resolved mechanism driven by a CaS with high Ca  affinity that can detect modest, transient levels of Ca  likely near the pore of presynaptic Ca  channels (Atluri and Regehr, 1996).  The exact mechanisms and molecular players involved in the latter form of facilitation have not been previously reported, although a role for high affinity CaSs such as CaM has been predicted (Ullrich et al., 1994; Li et al., 1995; Atluri and Regehr, 1996).  In this chapter, we showed 2+ 2+ 2+ 2+ that the R192Q and S218L FHM-1 mutations occlude CaM mediated CDF of both human recombinant Cav2.1 channels (see Fig. 3.1) and endogenous Cav2.1 channels in dissociated mouse cerebellar Purkinje neurons (see Fig. 3.3). Further, homologous mutations introduced into mice by genetic knock-in resulted in a corresponding attenuation in short-term synaptic facilitation at the PF-PC synapse (see Fig. 3.4).  The evidence supports the hypothesis that, in addition to the traditional view that residual Ca2+ enhances vesicle fusion by binding to CaSs involved directly in vesicle release (Zucker and Regehr, 2002), an initial Ca2+ influx into PF boutons induces CDF of Cav2.1 channels and acts to enhance Ca2+ influx during subsequent action potentials resulting in synaptic facilitation at this central synapse.  3.3.2 Facilitated Cav2.1 channels may increase susceptibility to CSD and aura Our findings may also provide insight into the molecular mechanisms of FHM-1 pathophysiology.  We find that FHM-1 mutations likely render Cav2.1 channels in a basally facilitated state and that the facilitated channels result in a larger Ca2+ influx during APs in PF boutons relative to wild-type channels (see Fig. 3.6).  Of note, in previous experiments using single channel recordings, some FHM-1 missense mutations (including R192Q and S218L) were shown to cause an overall gain-of- function Cav2.1 channel phenotype as a result of enhanced Po (Hans et al., 1999; Tottene et al., 2002).  126 Likewise, the mechanism that underlies CDF of Cav2.1 channels has been determined to be a Ca2+/CaM mediated transition of channels to a functional state with an enhanced Cav2.1 Po and facilitated mode of gating (Chaudhuri et al., 2007).  As such, at a mechanistic level, we predict that the R192Q and S218L mutations increase the likelihood of channels in a functional state with enhanced Po that is the same as (or analogous to) the facilitated state resulting from CDF and precludes further CDF-mediated facilitation.  In addition to directly affecting synaptic efficacy, this gain-of-function state is predicted to have important implications for migraine pathophysiology.  For example, increased Ca2+ influx during APs and increased glutamate release at pyramidal cell synapses are believed to be the underlying cause of both the lower threshold for stimulation of CSD and the increased velocity of propagation of CSD observed in FHM-1 knock-in mice.  CSD is believed to be the underlying mechanism of aura which precedes headache in some patients and likely triggers headache pain (van den Maagdenberg et al., 2004; Gherardini et al., 2006; Tottene et al., 2009; van den Maagdenberg et al., 2010).  3.3.3 Impairment of synaptic efficacy may underlie cerebellar ataxia in FHM-1 We provide the first evidence that FHM-1 mutations alter neurotransmission in the cerebellum. There has not been any explanation for the cerebellar ataxia sometimes associated with the FHM-1 phenotypes.  The cerebellum plays a central role in motor control, and alteration in neurotransmission at the PF-PC synapse has been correlated with aberrant motor phenotypes in other conditions involving ataxia (Zhou et al., 2003; Schmitt et al., 2009).  Here we showed evidence that the R192Q and S218L mutations alter short-term synaptic plasticity at the PF-PC synapse by attenuating CDF of Cav2.1.  We hypothesize that the significant effects on Cav2.1 CDF by the S218L mutation are sufficient to alter synaptic plasticity to an extent that ultimately leads to cerebellar motor deficits including ataxia under conditions associated with initiating factors of FHM-1 attacks.    127 3.3.4 Beyond the PF-PC synapse In summary, the data presented in this chapter provide evidence that Cav2.1 CDF plays a critical role in synaptic plasticity at the PF-PC synapse and provides the first direct correlation between Ca2+- dependent regulation of Cav2.1 and human disease.  Considering that Cav2.1 channels are the predominant Ca2+ channel underlying synaptic transmission at most other fast synapses in the mammalian CNS, and the PF-PC synapse is prototypical of other CNS synapses (Westenbroek et al., 1995; Evans and Zamponi, 2006), our findings support the notion that Cav2.1 CDF is an unrecognized but significant contributor to short-term synaptic facilitation at fast CNS synapses.  The disruption of Cav2.1 CDF may be an important factor in FHM-1 pathophysiology, although the precise effects of the FHM-1 mutations at other CNS synapses may vary depending on other factors including the nature of the Cav2.1 splice variants expressed within the synapse (Soong et al., 2002; Chaudhuri et al., 2004; Chaudhuri et al., 2005; Adams et al., 2009) as well as Cav2.1 interactions with auxiliary subunits (Mullner et al., 2004) and the expression of CaS proteins (Tsujimoto et al., 2002).       3.4 Experimental Procedures 3.4.1 Cell culture and transfection HEK 293 cells were grown in standard Dulbecco’s modified Eagle’s medium supplemented with 10% fetal bovine serum (heat inactivated) and 50 U/ml penicillin-50 ug/ml streptomycin.  Cells were incubated at 37oC in a humidified incubator with 95% air and 5% CO2 and grown to 10-20% confluency for transfection.  HEK 293 cells were transiently transfected with either wild-type, R192Q or S218L human Cav2.1 alpha subunits in combination with Ca2+ channel β2, α2δ1, and CD8 in a 1:1:1:0.25 molar ratio using Lipofectamine (Invitrogen, La Jolla, CA).  To ensure accurate comparisons, transfections were performed at the same time and electrophysiological recordings alternated within the same day for all channel types.  All clones used in the present chapter were generated as previously described (all clones are the Cav2.1 (Δ47) splice-variant) (Adams et al., 2009).  128  3.4.2 Dissociated PCs Mouse cerebellar PCs were enzymatically isolated using dissociation techniques previously described (Raman et al., 1997; Swensen and Bean, 2003, 2005).  Mice between postnatal days 15-25 were anesthetized using isoflurane (Baxter Corporation, Mississauga, ON), and decapitated according to protocols approved by the University of British Columbia animal care committee.  Cerebellum vermis were dissected out and cut into 1 mm cubes in ice cold dissociation solution containing (in mM): 82 Na2SO4, 30 K2SO4, 10 HEPES, 10 glucose, 5 MgCl2 and 0.001% Phenol Red, buffered to pH 7.4 with NaOH. Cerebellar cubes were transferred to 10 mL of room-temperature dissociation solution containing 3 mg/mL of protease XXIII (Sigma, St. Louis, MO) and incubated for 8-10 minutes at 32oC.  After incubation, tissue was transferred to ice-cold dissociation solution containing 1 mg/mL trypsin inhibitor and 1 mg/mL bovine serum albumin (Sigma, St. Louis, MO) and maintained on ice until trituration. Tissue was triturated with a series of three fire-polished Pasteur pipette to dissociate individual cells, and PCs were identified by their large diameter and pear-shaped appearance attributed to the stump of the dendritic tree.  3.4.3 Electrophysiological recording conditions in HEK and dissociated PCs Recombinant human Cav2.1 currents in HEK cells were recorded the second day after transfection, whereas mouse endogenous Cav2.1 currents in acutely dissociated PCs were recorded between 30 minutes and 4 hours after dissociation.  In both cases, macroscopic Ba2+ and Ca2+ currents were recorded at 19-23oC using the whole-cell patch-clamp technique (Hamill et al., 1981).  The internal pipette solution contained 135 mM Cs-MeSO3, 5 mM CsCl2, 0.5 mM EGTA, 1 mM MgCl2, 4 mM MgATP, and 10 mM HEPES (pH 7.4); external: 140 mM TEA-MeSO3, 10 mM HEPES, and 5 mM CaCl2 or BaCl2 (pH 7.3).  Patch pipettes (borosilicate glass BF150-86-10; Sutter Instrument Company, Novato, CA) were made using a horizontal puller (P-87; Sutter Instruments Company), fire polished using a microforge (Narishinge, Tokyo, Japan), and had resistances typically of 3 to 7 MΩ when  129 containing internal solution.  External solution bath was connected to ground with a 3M KCl agar bridge. Whole cell currents were recorded and filtered at 2-5 kHz bandwidth using an Axopatch 200A amplifier monitored and stored on a personal computer running pClamp software package version 9.  Recordings were analyzed using Clampfit 9 and figures, fittings and statistics (ANOVA) were made using the software program Origin version 7.5 (OriginLab Corp., Northampton, MA).  3.4.4 Protocols and data analysis for CDF and CDI  CDF was analyzed using a two-pulse protocol as previously described (Chaudhuri et al., 2004). In short, 50 ms test pulses to +5 mV were given subsequent to no prepulse or prepulses ranging from -90 mV to +100 mV.  All traces were normalized to the current at the end of the test pulse.  We integrated the current trace for the test pulse obtained with a prepulse and subtracted that integral from the integral for the test pulse obtained without a prepulse in order to obtain the variable delta Q.  To obtain the value of relative facilitation (RF) for all channels, we divided delta Q by the time constant obtained from fitting a single exponential to the trace obtained by subtracting the normalized test pulse current obtained without a prepulse from the normalized test pulse current obtained with a +20 mV prepulse from wild- type channels (see supplemental material in (Chaudhuri et al., 2004)).  Relative facilitation is plotted vs. prepulse membrane potential.  To index pure CDF we measured the difference in the relative facilitation values obtained in Ca2+ vs. Ba2+ with a prepulse potential of +20 mV.  CDF was measured in PCs using both the rectangular two pulse protocol and the APW described above, except the holding potentials were changed form -90 mV to -60 mV to eliminate any contribution of T-type Ca2+ currents that may have been present in some cells (Chaudhuri et al., 2005). CDI was measured using a square pulse protocol including a 1 s test depolarization ranging from -10 mV to +30 mV, in 10 mV increments, from a holding of -90 mV (-60 mV holding in PCs).  The fraction of current remaining at 800 ms after the initiation of the test pulse, r800, was plotted as a function of test pulse potential.  To index pure CDI we measured the difference in the r800 values obtained in Ca2+ vs. Ba2+ with a test potential of +10 mV.  130  3.4.5 Cerebellar slices For both extracellular field recording and two-photon imaging experiments, mice between postnatal days 25-35 were anesthetized using isoflurane (Baxter Corporation, Mississauga, ON) and decapitated according to protocols approved by the University of British Columbia animal care committee.  The Cerebellum was rapidly removed and placed in ice-cold dissection solution containing (in mM): 87 NaCl, 25 NaHCO3, 25 D-glucose, 75 sucrose, 2.5 KCl, 2 NaH2PO4, 0.5 CaCl2, 7 MgCl2. Cerebellum was cut into 300 μm transverse slices using a vibrating tissue slicer (Vibratome, St. Louis, MO) and maintained for 1-6 hours at room temperature in artificial cerebral spinal fluid (aCSF) containing (in mM): 119 NaCl, 2.5 KCl, 1.3 MgSO4, 26 NaHCO3, 2.5 CaCl2, 10 Glucose, 1.4 NaH2PO4 and aerated with 95% O2 / 5% CO2.  For experiments slices were at 22–24 °C and perfused at ~2 ml/min.  3.4.6 Extracellular Field Recordings fEPSPs were evoked by orthodromic stimulation of the PFs in the molecular layer of the cerebellar vermis using a bipolar tungsten stimulating electrode.  Glass micropipettes filled with aCSF (resistance, 1–3M) were used to measure PC fEPSPs.  PPF was measured by invoking two extracellular stimuli of ~15V, 180 μs delivered at 50 ms intervals and the PPR quantified by dividing response two by response one.  Similarly, synaptic responses were also evoked by five extracellular stimuli of ~15V, 180 μs delivered at 20 Hz and relative facilitation measured by dividing the peak response from each stimulus by the response obtained from the first stimulus. fEPSP signals were amplified 1000 times with an AC amplifier (AM Systems, Sequim, WA) , bandpass filtered at 1 kHz, digitized at 10 kHz using a Digidata 1322A interface board (Molecular Devices, Foster City, CA), and transferred to a computer for analysis.  Data were analyzed using Clampfit 9.0 (Molecular Devices, Foster City, CA).   131 3.4.7 Two-photon microscopy: Ca2+ Imaging Neurons in cerebellar slices (300 µm) were bulk loaded with Rhod-2 AM (Invitrogen) using a modified Chremophor loading technique adapted from (Trevelyan et al., 2006).  Slices were pre- incubated in 3 mL ACSF and 8 uL Chremophor EL solution (0.5% in DMSO) at 34ºC for 5 mins.  50 μg Rhod-2 AM mixed with 8 µL DMSO and 2 µL pluronic F-127 solution (10% in DMSO) was then added, and slices were allowed to incubate for an additional 45minutes.  Slices were then allowed to recover at room temperature for at least 30 min.  This resulted in robust loading of PFs in addition to staining of other cell types in the tissue.  In order to isolate pre-synaptic Ca2+ signals, post-synaptic and glial signals were blocked with a cocktail of glutamate receptor blockers; 20 µM CNQX, 50 µM DL-APV, 100 µM MCPG (all purchased from Tocris). We performed imaging with a two-photon laser scanning microscope (Zeiss LSM510-Axioskop- 2 with a 40X/1.0 numerical aperture objective lens) directly coupled to a 10 W Chameleon ultrafast laser (Coherent).  PFs were stimulated with a silver wire electrode and two-photon imaging used to measure resulting Ca2+ transients in presynaptic terminals of the PFs.  Line scans were performed at 325 Hz, and individual terminal responses were measured in response to four sets of five 50 μs duration stimuli applied at 20 Hz.  Images were obtained from between 40 and 80 μm deep into the slice. Rhod-2 was excited at 835 nm ( 3 mW after the objective) and fluorescence detected with a PMT after passing through a 605-nm (55-nm band-pass) emission filter.  Images were collected and analyzed using Zeiss LSM software.        132      3.5 Acknowledgements We thank Drs. David Yue, Bruce Bean and Amy Lee for protocols and advice on optimizing PC dissociation procedures; Dr. David Parker, Luke Materek and Paul Lam for providing human Cav2.1 α1 subunit constructs; Drs. Patrick Francis and Charles A. 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Alternative splicing of the CACNA1A gene has the potential to produce numerous functionally distinct Ca 2.1 splice variants that are differentially expressed regionally within the nervous system in discrete cell types and subcellular compartments and at different developmental stages.  T v he first major objective of this thesis was to explore whether Cav2.1 splice variants respond differently to FHM-1 mutations and to perhaps contribute toward understanding the localized and episodic nature of FHM-1. Previous studies exploring FHM-1 effects on Cav2.1 channels have used either a single recombinant Cav2.1 channel variant expressed in Xenopus oocytes or HEK cells, or endogenous Cav2.1 channels in neurons which contain numerous Cav2.1 splice variants associated with various auxiliary subunits (Tsunemi et al., 2002; Kanumilli et al., 2006; Richards et al., 2007).  Whether the effects of FHM-1 mutations are contingent on the Cav2.1 splice-variant has not been discernible.  In Chapter 2, we utilized an experimental design that allowed us to specifically compare how two human recombinant Cav2.1 splice variants expressed in a standardized system (HEK cells co-expressing the same VGCC auxiliary subunits) respond to three different FHM-1 mutations.  We found that all three FHM-1 mutations induced significant changes in channel basic biophysical properties and that the effects were in fact contingent upon the nature of the Cav2.1 splice-variant. The three FHM-1 mutations all caused greater gain-of-function effects regarding channel activation when expressed in Cav2.1 (Δ47) channels relative to Cav2.1 (+47) variant channels; mutant  138 Cav2.1 (Δ47) channels activate at lower membrane potentials relative to mutant Cav2.1 (+47) channels (Fig 2.2 and Table 2.1).  Such gain-of-function effects in Cav2.1 channels are predicted to increase Ca2+ influx during APs and enhance susceptibility to CSD and the initiation of migraine (van den Maagdenberg et al., 2004; Pietrobon, 2007; Tottene et al., 2009).  The results in Chapter 2 support the hypothesis that Cav2.1 splice variants (e.g., Cav2.1 (Δ47)) with greater sensitivity to these and other FHM-1 mediated gain-of-function effects might result in cortical regions (or cell types) with a greater susceptibility to CSD and/or migraine initiation under some conditions, while the effects of FHM-1 mutations on other Cav2.1 splice variants (e.g., Cav2.1 (+47)) expressed elsewhere may be below the threshold to initiate CSD and/or other pathological effects. Further, we found evidence that FHM-1 mutations differentially affect Cav2.1 channel splice variants under different firing conditions.  For example, during tonic depolarizations the R192Q mutation had no effect on the Cav2.1 (+47) channel, yet during bursts of depolarizations the mutation caused a significant enhancement in channel activity relative to wild-type.  On the other hand, when expressed in the Cav2.1 (Δ47) variant, the R192Q mutation decreased channel activity during both tonic and bursts of depolarizations relative to wild-type (Figs 2.4 and 2.5).  Interestingly, neuronal firing patterns in the brain can be altered by events such as emotional stress (Weiss and Simson, 1988; McEwen, 2007), a common initiating factor of FHM-1 attacks (Ducros et al., 2001).  As such, although the exact firing conditions directly associated with the precipitating factors of migraine are unknown, our results support the conclusion that the episodic nature of the FHM-1 phenotype may, in part, be associated with changes in neuronal firing patterns and/or frequency that affect certain Cav2.1 splice variants localized to specific brain regions.  4.1.2 FHM-1 mutations occlude Ca2+-dependent modulation of Cav2.1 and synaptic plasticity CDF and CDI are robust forms of Cav2.1 Ca2+-dependent modulation, yet whether they are altered by FHM-1 mutations has not been reported.   Furthermore, Cav2.1 CDF and CDI are predicted to  139 have important roles in neurotransmission and synaptic plasticity (Tsujimoto et al., 2002; Chaudhuri et al., 2005; Mochida et al., 2008), however their relevance within intact neuronal circuits is uncertain.  The second major objective of this thesis was to determine whether CDF and CDI of Cav2.1 are altered by FHM-1 mutations, and to explore any physiological implications toward normal synaptic efficacy and also the pathophysiology associated with migraine using transgenic mouse models. In Chapter 3, we first measured CDF and CDI of recombinant Cav2.1 channels transiently expressed in HEK cells (along with auxiliary subunits β2a and α2δ1).  This recombinant system is well characterized (Lee et al., 1999; Lee et al., 2000; DeMaria et al., 2001; Chaudhuri et al., 2005; Kreiner et al., 2010) and allowed us to obtain clear isolation and measurement of CaM mediated CDF and CDI for both wild-type and R192Q and S218L Cav2.1 channels.  We further tested the FHM-1 mutations in the context of endogenous Cav2.1 channel modulation by measuring P/Q-type currents in acutely dissociated cerebellar PCs from wild-type, R192Q and S218L knock-in mice (van den Maagdenberg et al., 2004; Eikermann-Haerter et al., 2009; Tottene et al., 2009).  The use of both recombinant and endogenous experimental paradigms provided strong corroborative evidence that the R192Q and S218L FHM-1 mutations occlude CDF of Cav2.1 channels.  These findings are the first evidence that mutations associated with human disease alter Cav2.1 Ca2+-dependent modulation. A correlation between such Cav2.1 CDF mechanisms and short-term synaptic plasticity had previously been achieved using recombinant Cav2.1 channels expressed in a model system (Mochida et al., 2008).  In order to establish the relevance of this Cav2.1 CDF in synaptic plasticity at an intact central synapse, we compared fEPSPs at the PF-PC synapse in wild-type and R192Q and S218L mice.  We found that the occlusion of Cav2.1 CDF by the mutations corresponded with an attenuation in short-term synaptic plasticity at the PF-PC synapse.  This is the first direct evidence that initial Ca2+ influx at these central presynaptic terminals induces CDF of Cav2.1 channels and acts as a means to enhance Ca2+ influx during subsequent action potentials to achieve synaptic facilitation.  In Chapter 3 we also used two-photon microscopy to measure Ca2+ transients during APs at PF boutons in wild-type and FHM-1 mice.  With this high-resolution technique, we obtained evidence that  140 FHM-1 mutations render Cav2.1 channels toward a basally facilitated state, and that the facilitated channels allow larger Ca2+ influx during evoked APs in presynaptic terminals that rely predominantly on Cav2.1 channels.  The findings suggest that a similar mechanism of gain-of-function may contribute to the observed increased Ca2+ influx during APs and increased glutamate release at pyramidal cell synapses believed to cause both lower threshold for stimulation of CSD and increased velocity of propagation of CSD across the cortex (van den Maagdenberg et al., 2004; Gherardini et al., 2006; Tottene et al., 2009; van den Maagdenberg et al., 2010). Lastly, the findings outlined in Chapter 3 provide initial evidence for a possible mechanism underlying cerebellar dysfunction in severe FHM-1 phenotypes.  The S218L mutation is associated with a phenotype that includes episodes of cerebellar ataxia.  Our data show that the S218L mutation enhances presynaptic Ca2+ influx in PF boutons and synaptic efficacy at the PF-PC synapse is significantly compromised.  The cerebellum plays a central role in motor control and alteration in neurotransmission at the PF-PC synapse has been correlated with aberrant motor phenotypes in other conditions involving ataxia (Zhou et al., 2003; Schmitt et al., 2009).       4.2 FHM-1 mutations differentially affect Cav2.1 splice variants: a role in the localized, episodic nature of the FHM-1 phenotype 4.2.1 Working hypothesis Over the years a number of studies have shown that alternative splicing of Cav2.1 and other Cav channels can be integral to the localized phenotype of human disease, although critically, in ways distinct from that described in Chapter 2 (reviewed in (Adams and Snutch, 2007; Liao et al., 2009)).  The most apparent is the use of alternative exons by Cav channels (see Chapter 1, section 1.3.3).  If a disease- causing mutation occurs within an alternative exon that is only expressed in a subset of tissues and/or cells, then the phenotype is localized and the severity of the disease is correlated with the exon expression level.  For example, in the L-type Cav1.2 channel, two de novo mutations in one of two  141 mutually exclusive exons (exon 8 and 8a) have been associated with Timothy syndrome (TS) (Splawski et al., 2005)(and reviewed in (Adams and Snutch, 2007); Appendix 1).  Exons 8 and 8a are preferentially expressed in smooth muscle and cardiac muscle, respectively (Welling et al., 1997; Liao et al., 2005).  A TS mutation in exon 8 is associated with a mild phenotype and patient longevity is not compromised (Splawski et al., 2004), whereas a mutation in exon 8a is associated with severe cardiac arrhythmia and patient death between two and three years of age (Splawski et al., 2005). Similarly, three mutations causing premature stop codons in the mutually exclusive exon 37a of Cav2.1 were identified in several EA2 patients.  Exons 37a and 37b are used mutually exclusively in humans and rodent (Bourinet et al., 1999; Soong et al., 2002; Kanumilli et al., 2006).  37a is preferentially expressed in cerebellar cortex, predominantly in Purkinje neurons (Bourinet et al., 1999), and EA2 mutations in exon 37a thus correlate well with cerebellar dysfunction as the primary EA2 phenotype (Ophoff et al., 1996).  Also, in humans there is a developmental switch from 37a to 37b in the fourth decade of life (Chang et al., 2007a), which may contribute to the age-dependent decrease in frequency and severity of attacks in some EA2 patients (Jen et al., 2007). In another example, it has long been understood that expansion of the CAG trinucleotide repeat in exon 47 of Cav2.1 causes SCA6 (Zhuchenko et al., 1997).  Importantly, the trinucleotide expansion has strong effects on voltage and time-dependent properties of the Cav2.1 (+47) splice-variant containing exon 47, whereas the Cav2.1 (Δ47) variant lacking exon 47 is unaffected (Restituito et al., 2000; Toru et al., 2000).  Interestingly, recent analysis of human cerebellum revealed that the expression profile of Cav2.1 variants is altered in patients with the SCA6 CAG expansion.  Cav2.1 (+47) is found nearly exclusively in PCs from SCA6 patients, whereas in normal cerebellum Cav2.1 (+47) and Cav2.1 (Δ47) are expressed in equal proportion (Tsunemi et al., 2008).  The authors argue that the higher levels of Cav2.1 (+47) in PCs in SCA6 patients causes specific PC degeneration and cerebellar ataxia, perhaps due to Cav2.1 (+47) C-terminal fragments containing the SCA6 CAG expansion forming cytoplasmic and nuclear aggregates (Ishiguro et al., 2010).  142 There are other examples of Cav mutations that have been identified in alternative exons or within conventional splice-sites at intron-exon boundaries that are predicted to contribute to the localization of disease, however, the majority of identified mutations associated with Ca2+ channelopathies are missense mutations in coding sequences other than splice-sites and alternate exons (reviewed in (Adams & Snutch, 2007); Appendix 1).  Our results presented in Chapter 2 are therefore important evidence toward what may prove a more common mechanism underlying the localized episodic phenotype associated with the majority of Ca2+ channelopathies. The work in Chapter 2 has proven particularly relevant toward the interpretation of recent results obtained from FHM-1 studies.  Two research groups explored synaptic transmission in cortical neuron cultures and cortical slices from FHM-1 knock-in mice in order to gain insight into the mechanisms underlying the observed lower threshold for stimulation of CSD depression and increased velocity of propagation across the cortex (van den Maagdenberg et al., 2004; Gherardini et al., 2006; van den Maagdenberg et al., 2010).  In the cortex, layer 2/3 pyramidal cells have glutamatergic synapses onto fast spiking (FS) inhibitory interneurons which in turn make GABAergic inhibitory synapses onto pyramidal cells.  Using paired recordings in acute somatosensory cortical slices, it was shown that there is an increased probability of glutamate release from 2/3 pyramidal cells in R192Q mice relative to wild-type, but surprisingly, no change in GABAergic transmission at FS synapses, despite the fact that Cav2.1 channels are the predominant VGCCs in both types of terminals (Tottene et al., 2009; Inchauspe et al., 2010).  The authors hypothesize that pyramidal cells and FS interneurons normally coordinate a balance between excitation and inhibition during cortical activity, and that a compromise in the balance by FHM- 1 mutations under some conditions (i.e. migraine triggers that alter firing patterns and/or frequencies in the cortex) could cause an increase in extracellular K+ above the threshold for CSD initiation (Pietrobon, 2010b, a). Based on our results, it is possible that the differential effects of the R192Q mutation on the two cortical neuron types is due to the expression of different Cav2.1 splice variants.  For example, we showed that the R192Q mutation caused a greater hyperpolarizing shift in the activation threshold when  143 expressed in the Cav2.1 (∆47) variant relative to Cav2.1 (+47) (Fig. 2.2); perhaps 2/3 pyramidal neurons express Cav2.1 (∆47) and FS neurons Cav2.1 (+47).  Further, under a particular frequency of tonic depolarization, the R192Q mutation had a loss-of-function effect in one Cav2.1 variant and no detectable effect in the other variant (Fig. 2.4).  Also, during certain burst firing conditions, the R192Q mutation had a loss-of-function effect in one Cav2.1 variant and a gain-of-function effect in another (Fig. 2.5). These differing, or even opposing, effects by the R192Q mutation are potential ways in which the mutation could disrupt the balance between cortical excitation-inhibition in favor of excitation under certain firing conditions if pyramidal and FS neurons in fact express different Cav2.1 splice variants. Similar differential effects by FHM-1 mutations may be relevant to migraine pathophysiology within the trigeminovascular pain pathway where Cav2.1 channels are integral for both excitatory and inhibitory neurotransmission (see Chapter 1, section 1.4.4 for details on FHM-1 mechanism).  Cav2.1 channels are known to be involved in facilitating pain through their role in the release of both vasoactive neuropeptides from perivascular terminals of meningeal nociceptors (Hong et al., 1999; Akerman et al., 2003) and glutamate from trigeminal ganglion neurons (Xiao et al., 2008).  On the other hand, Cav2.1 channels are also involved in suppressing pain through their role in transmitter release from inhibitory projections from the dura and PAG onto the TNC (Knight et al., 2002; Ebersberger et al., 2004).  It is possible that differential or opposing effects of FHM-1 mutations on Cav2.1 splice variants expressed in these excitatory and inhibitory neurons disrupt the balance between excitation and inhibition within the neuronal network of the trigeminovascular system.  For example, gain-of-function in excitatory pathways and loss-of-function in inhibitory pathways could ultimately enhance neuronal activity in the trigeminal pain pathway leading to sensitization and migraine pain (Pietrobon and Striessnig, 2003; Moskowitz et al., 2004; Burstein and Jakubowski, 2005; Edvinsson and Uddman, 2005; Pietrobon, 2005; Sanchez-Del- Rio et al., 2006; Goadsby et al., 2009)). Similar to FHM-1, many other Ca2+ channelopathies exhibit episodic and/or developmentally specific phenotypes localized to a subset of regions or tissues, and like Cav2.1, other Cav channels also undergo alternative splicing that generates functionally distinct channel variants (reviewed in  144 (Lipscombe et al., 2002; Lipscombe and Castiglioni, 2004)).  I hypothesize that the differential effects of FHM-1 mutations on Cav2.1 splice variants represents a central mechanism relevant in the localized and episodic phenotype associated with all Ca2+ channelopathies.  In support, recent work performed in the Snutch lab showed that a missense mutation in Cav3.2 associated with a rat model of absence epilepsy differentially affected two distinct thalamic Cav3.2 T-type channel splice variants (Powell et al., 2009). Specifically, alternative splicing in exon 25 determined the effect of the mutation on recovery from inactivation and charge movement during high-frequency bursts.  4.2.2 Possible limitations The use of human recombinant Cav2.1 channels in HEK cells provided a highly controlled environment to specifically address how alternative splicing at a single locus can determine the functional consequences of FHM-1 mutations.  Although these experiments offer a powerful proof of principle, in reality, neurons in the brain likely express numerous Cav2.1 channels consisting of various combinations of splicing at the 7 known splice-sites (Soong et al., 2002; Tsunemi et al., 2002; Chaudhuri et al., 2004; Chaudhuri et al., 2005; Kanumilli et al., 2006; Richards et al., 2007).  As such, the overall functional effects of a given FHM-1 mutation in a particular neuron will likely depend on both the combination of Cav2.1 channel splicing at the seven loci, as well as the relative proportion of each Cav2.1 variant.  In future experiments it will be crucial to identify full-length alternative splice variants from brain regions and specific cell types implicated in FHM-1 pathophysiology. Another potential limitation in the HEK cell-based recombinant studies is that we only examined the auxiliary subunits β4 and α2δ1 coexpressed with Cav2.1.  Endogenous Cav2.1 channels in neurons can be associated with various β (β1-4) and α2δ subunits (α2δ1-4) which can both modify Cav2.1 channel biophysical properties (reviewed in (Dolphin, 2009)) and alter how the channels respond to FHM-1 mutations (Mullner et al., 2004).  As such, the effects of FHM-1 mutations on Cav2.1 splice variants within neurons may also be contingent upon the auxiliary subunit isoforms expressed and their relative proportions.  For example, the differential effects induced by the R192Q mutation in cortical 2/3  145 pyramidal neurons and FS interneurons may be independent of Cav2.1 alternative splicing and rather due to differential expression of β-subunits, or some combination of both.  For instance, the K1336E, W1684R and V1696I FHM-1 mutations all show varying functional effects on a single recombinant Cav2.1 variant channel when coexpressed with either β1, β3 or β4.  Most notably, the V1696I mutation causes a significant hyperpolarizing shift in the voltage dependence of activation of Cav2.1 when coexpressed with β1, but has no effect when coexpressed with either β3 or β4 (Mullner et al., 2004).  If FS interneurons express either β3 or β4, and pyramidal neurons express β1, FHM-1 mutations may have significant gain-of-function affects on pyramidal neurons with little effect on FS interneurons and ultimately lead to enhanced glutamate release and increased extracellular K+ above the threshold for CSD initiation.  All four β-subunit isoforms are expressed in mammalian cortex (Schlick et al.), although isoforms expressed specifically in 2/3 pyramidal and FS neurons has yet to be determined.  4.2.3 Future directions The identity and functional characterization of full-length Cav2.1 splice variants and auxiliary subunits expressed in layer 2/3 pyramidal cells, FS interneurons, meningeal nociceptors, trigeminal nuclei and cerebellar granule and Purkinje neurons will be necessary before a full appreciation of the role of Cav2.1 alternative splicing in the localized and episodic nature of the FHM-1 phenotype can be realized.  The first step would be to utilize single cell quantitative RT-PCR to amplify full-length Cav2.1 mRNA transcripts from these neurons in FHM-1 KI mouse models and wild-type littermates; it would be necessary to look at splice variants in both WT and mutant mice as mutations may alter the expression of Cav2.1 variants (Tsunemi et al., 2008).  It would also be relevant to perform similar Cav2.1 splice-variant analysis in brain regions of the KI mice that are known to have no involvement in FHM-1 pathology for comparison (for example hippocampus, thalamus and amygdala) (Pietrobon, 2005).  Cav2.1 splice variants identified as unique to brain regions associated with FHM-1 pathology could be investigated further as variants specifically involved in the disease.  Experimental procedures outlined in Chapters 2 and 3 in addition to experiments using APWs relevant to neurons implicated in FHM-1 could be  146 effectively used to establish any differential effects of FHM-1 mutations in the context of the disease variants vs. variants from unaffected brain regions.       4.3 A role for Cav2.1 CDF in synaptic plasticity 4.3.1 Working hypothesis In one aspect of Chapter 3 we provide evidence that Cav2.1 CDF is a required component of short-term synaptic facilitation at the PF-PC synapse.  The proportion of facilitation attributed to Cav2.1 CDF at this synapse can be deduced from experiments in which presynaptic terminals were treated with high (100 μM) concentrations of EGTA-AM (Atluri and Regehr, 1996a).  In these experiments, EGTA- AM sped the decay of intracellular Ca  to the extent of removing the component of synaptic facilitation dependent upon residual Ca  in the traditional sense.  The remaining “residual-Ca -independent” form of synaptic facilitation exhibited a short intrinsic time constant, rapid kinetics and high Ca  affinity all similar to that described for CaM mediated Ca 2.1 CDF (Atluri and Regehr, 1996b; Lee et al., 2000; DeMaria et al., 2001).  During paired APs (100 Hz) in absence of EGTA-AM, both forms of facilitation were present and produced 125% facilitation of the second post-synaptic response relative to the first, but in the presence of 100 μM EGTA-AM only 25% facilitation of the post-synaptic response remained (Atluri and Regehr, 1996b).  Taken together, this suggests that Ca 2.1 CDF likely accounts for approximately 25% of the facilitation observed at the PF-PC synapse during paired APs. 2+ 2+ 2+ 2+ v v The coupling between Ca  influx through Ca 2.1 channels and neurotransmitter release within PF boutons, and the intrinsic properties of Ca 2.1 CDF observed in recombinant and endogenous channels both support a value of around 25% synaptic facilitation mediated by Ca 2.1 CDF.  Although PF presynaptic terminals express Ca 2.1, Ca 2.2 and Ca 2.3 channels (Randall and Tsien, 1995), Ca 2.1 channels are responsible for approximately 60% of presynaptic Ca  influx during APs, are the most effective VGCC at triggering transmitter release from PF terminals, and are thus responsible for nearly 93% of the synaptic response (Mintz et al., 1995).  There is a strong correlation between Ca  influx in 2+ v v v v v v v 2+ 2+  147 PF boutons and transmitter release such that the relationship between Ca  influx through Ca 2.1 and transmitter release is well approximated by a power law, n ≈ 2.3 (release = k(Ca )  ; k is a constant) (Mintz et al., 1995; Sabatini and Regehr, 1997).  Thus, even the relatively small enhancement and broadening of APs predicted to result from Ca 2.1 CDF (Chaudhuri et al., 2007) could significantly contribute to short-term facilitation at this synapse.  For example, in Figure 3.6 panel D, the peak Ca currents through recombinant Ca 2.1 channels increase during repetitive APs (100 Hz) such that the peak current elicited by a second AP is facilitated by ~10% relative to the first (or ~5% facilitation in Ca 2.1 channels in dissociated mouse Purkinje neurons (Fig. 3.3 D)).  With a power law relationship between Ca  influx and neurotransmission of ~2.3, a 5 to 10% increase in the maximum Ca 2.1 Ca  influx due to CDF within PF terminals during paired APs could account for ~12 to 25% facilitation in the post- synaptic response at the PF-PC synapse.  In support, in a model system Ca 2.1 CDF was shown to account for approximately 30% of synaptic facilitation during paired APs (Mochida et al., 2008).  As such, I hypothesize that within PF presynaptic terminals 2+ v 2+ influx n v 2+ v v 2+ v 2+ v residual Ca2+ can control Ca2+ influx through Cav2.1 during an action potential to shape the local Ca2+ transient at the active zone and cause between 12 and 25% facilitation during paired APs and likely more during trains of APs through the process of Ca 2.1 CDFv . In addition to an important role in synaptic plasticity, our results in Chapter 3 indicate that Cav2.1 CDF is likely involved in the pathophysiology of FHM-1.  Disruption of Cav2.1 CDF and synaptic efficacy at the PF-PC synapse may explain the cerebellar ataxia often associated with FHM-1. PCs provide the sole output of cerebellar cortex and are integral in providing signals required for motor planning, execution and coordination (Voogd and Glickstein, 1998; Ito, 2000, 2002; Boyden et al., 2004) (Chapter 1, section 1.6.1).  The signals from PCs are encoded in their rate of firing and pattern of activity.  PCs have regular intrinsic pacemaking that is shaped by various inhibitory and excitatory inputs onto PCs, and fine-tuning (increase or decrease) of the inter-spike interval from that of intrinsic pacemaking conveys information to the DCN relevant to motor coordination (Ebner, 1998).  As such, the significant increase in presynaptic Ca2+ influx and presumably enhancement in glutamate release from PF induced by the S218L FHM-1 mutation (Fig. 3.6) may alter the precise balance of excitatory and  148 inhibitory inputs onto PC under some conditions.  In so doing, the fine-tuning of PCs firing and information conveyed to DCN may be compromised sufficiently to lead to cerebellar motor deficits including ataxia.   In fact, other mutations in the Cav2.1 channel that increase, decrease and/or alter the dynamics of excitatory inputs to PC have been correlated with aberrant motor phenotypes in several other animal models involving episodic ataxia.  For example, the tg and tgrol mice are established animal models of cerebellar ataxia caused by point mutations in the mouse cacna1a gene encoding Cav2.1. Similar to the severe form of FHM-1 associated with the S218L mutation, the tg and tgrol phenotypes include episodic attacks of cerebellar ataxia.  The cerebellar motor deficits in these mice have been attributed to decreased excitatory transmission at the PF-PC synapse (Matsushita et al., 2002; Zhou et al., 2003).  Although opposite in effect to the enhanced PF-PC transmission in S218L mice, the end result is disruption in the precise balance of excitatory and inhibitory inputs onto PCs that likely alters the fine- tuning of PC intrinsic firing and information processing. Considering that Cav2.1 channels are the predominant VGCC underlying synaptic transmission at most fast synapses in the mammalian CNS and that the PF-PC synapse is prototypical of other CNS synapses (Westenbroek et al., 1995; Evans and Zamponi, 2006), I predict that Cav2.1 channel CDF is a largely unrecognized but significant contributor to short-term synaptic facilitation at other fast synapses in the CNS (Fig. 4.1; proposed model).  At terminals expressing Cav2.1, Ca2+ influx during a single AP likely binds both CaS molecules directly associated with vesicle fusion (such as synaptotagmin) and CaS molecules that cause CDF of Cav2.1 (such as CaM) (Fig. 4.1A-C).  As a result, during subsequent APs, Cav2.1 channels mediate a greater influx of Ca2+ that combines with residual Ca2+ and causes a larger synaptic response (Fig. 4.1D-F).  However, not all Cav2.1 splice variants exhibit the same level of CDF and synapses with different Cav2.1 splice variants may have a different reliance on Cav2.1 CDF.  For example, the use of the EFa and EFb alternative exons and the Δ47 and +47 variants in the C-terminus of Cav2.1 affects the magnitude of CaM mediated Cav2.1 CDF (Chaudhuri et al., 2004).  Cav2.1 channels containing the EFa exon exhibit robust CDF (in combination with either Δ47 or +47 variants), whereas Cav2.1 channels containing the EFb exon exhibit CDF only when in combination with the Δ47 variant. There is approximately equal expression of the 37a (40.5%) and 37b (59.5%) variants in mammalian  149 cerebellum, hippocampus and cerebral cortex, a large preferential expression of EFb in amygdala, and predominant expression of EFa in the substantia nigra and thalamus (Soong et al., 2002; Chaudhuri et al., 2004).  It is unknown what the expression of each variant is within specific cell types in these various brain regions (Kanumilli et al., 2006; Richards et al., 2007).  An additional consideration is that there are profound changes in the expression pattern of the splice variants during development within various brain regions and also a clear gender bias in rodent and human brain (Chaudhuri et al., 2005; Chang et al., 2007b).  In addition to Cav2.1 alternative splicing, coexpression with various beta subunits, AP frequency and the types of CaS proteins expressed within terminals likely determine the magnitude of Ca 2.1 CDF and its contribution to synaptic facilitation in central synapses. v                150   Figure 4.1: Schematic model outlining the role of CDF of Cav2.1 channels as a key component of short-term synaptic facilitation. (A) A resident CaS (such as calmodulin; CaM) is bound to the Cav2.1 channel in a Ca2+ free form (ApoCaM).  (B) During an action potential (AP1), Ca2+ enters synaptic terminals primarily through Cav2.1 channels.  (C) Ca2+ ions bind to CaS molecules (such as synaptotagmin) which are directly associated with vesicle fusion and transmitter release functions.  Transmitter release causes a modest post-synaptic potential (PSP 1).  In addition, according to our model, Ca2+ ions near the pore of the channel also bind to a CaS such as ApoCaM causing a conformational change in the Cav2.1 channel rendering it in a facilitated state; a state of higher open-channel probability.  (D) Tens of milliseconds later, some residual Ca2+ (Ca2+ with blue rings) remains in the terminal and Cav2.1 channels are in a facilitated state.  (E) Upon arrival of a second AP at the terminal (AP 2), facilitated Cav2.1 channels mediate a greater influx of Ca2+ which combines with the residual Ca2+.  (F) The combination of residual Ca2+ and the greater Ca2+ influx through facilitated Cav2.1 channels result in more vesicles fusing with the membrane and greater transmitter release which is detected as a larger PSP (PSP 2).   151 If Cav2.1 CDF is relevant to other CNS synapses besides the PF-PC synapse, then this phenomenon may also be important in FHM-1 pathophysiology beyond cerebellar dysfunction.  For example, disruption in the balance between excitation and inhibition within neuronal networks in the cerebral cortex and trigeminovascular pain pathways likely contribute to lower threshold for CSD and sensitization and migraine pain, respectively (see discussion in section 4.2.1).  By rendering channels in a constitutively facilitated state, FHM-1 mutations may disrupt the balance between excitation and inhibition.  Of note however, preliminary evidence examining the –NP and +NP Cav2.1 splice variants indicates that the effect of FHM-1 mutations on Cav2.1 CDF can be splice-variant dependent (Fig 4.2). As such, only cells expressing certain Cav2.1 variants will likely be affected.  I hypothesize that a combination of the differential effects of FHM-1 mutations on CDF of specific Cav2.1 splice variants combined with differences in the basal level of Cav2.1 CDF at various CNS synapses will determine the role of CDF toward FHM-1 mediated pathophysiology.              152  A B Figure 4.2: R192Q and S218L FHM-1 mutant effects on Cav2.1 CDF are splice-variant dependent. Using the paired-pulse protocol as described (Chaudhuri et al., 2004), both the R192Q and S218L FHM- 1 mutations are found to reduce CDF across several prepulse potentials shown as relative facilitation vs. prepulse potential (results are means ± s.e.m).  Both FHM-1 mutations tested have a differential effect on CDF of two predominant human Cav2.1 splice variants.  Both mutations have a larger effect on CDF when expressed in the Cav2.1 (–NP) variant, A, relative to when expressed in the Cav2.1 (+NP) variant, B.  Constructs are human Cav2.1 (Δ47) transiently transfected with β2 and α2δ-1 in HEK 293 cells.  The effect of the mutations on CDF was similar in both the Cav2.1 (Δ47) and Cav2.1(+47) variants (data not shown).  4.3.2 Possible limitations In Chapter 3 we provided empirical evidence that FHM-1 mutations disrupt Cav2.1 CDF and cause a corresponding change in short-term synaptic facilitation at the PF-PC synapse.  We discuss several lines of evidence from other CNS synapses, a model system and properties of the PF-PC synapse that support the conclusion that Cav2.1 CDF is likely an important mechanism of short-term facilitation at the PF-PC synapse.  However, we have not yet shown that Cav2.1 CDF exists under paired APs in wild-type terminals.  It would be ideal to demonstrate under paired APs in wild-type animals that Ca2+  153 influx through Cav2.1 channels in PF boutons is facilitated 5-10% during the second AP.  However, there are several practical and technical challenges.  For one, Cav2.1 channels account for ~60% of Ca2+ influx in PF boutons whereas the remainder is mostly through Cav2.2 and some Cav2.3 channels (Mintz et al., 1995).  Although Cav2.2 and Cav2.3 channels do not undergo CDF, they do undergo various forms of VD facilitation (Liang et al., 2003) and to properly isolate Cav2.1 CDF it would be necessary to block these channels.  Due to the small size of mouse PF terminals, with only 60% of the Ca2+ signal remaining the Ca2+ signal elicited relative to background fluorescence would be very low and a 5-10% change above the residual Ca2+ signal would likely not be resolved.  In fact, when describing the two components of facilitation involved within the PF boutons, Atluri and Regehr recognized that the smaller “residual- Ca2+-independent” component could not be adequately resolved using Ca2+-sensitive fluorescent indicators (Atluri and Regehr, 1996b).  The direct measurement of Cav2.1 CDF in presynaptic terminals will require higher resolution techniques. An additional area of potential concern is that both the R192Q and S218L mutations cause a hyperpolarizing shift in the current-voltage relationship under square-pulse depolarizing voltage steps (Chapter 2, and other authors cited elsewhere).  It could be argued that the shift to more negative activation voltages is responsible for the increased Ca2+ influx at PF terminals in S218L mice; assuming that the kinetics of presynaptic Ca2+ currents through Cav2.1 channels can be modeled by Hodgkin- Huxley equations, a shift to more negative activation voltages theoretically could generate larger Ca2+ currents during APs (Borst and Sakmann, 1999).  In fact, a hyperpolarizing shift by the R192Q mutation appears to be responsible for increased Ca2+ influx during APs in pyramidal neurons in cortical slices from R192Q knock-in mice (Tottene et al., 2009; Inchauspe et al., 2010).  However, at the calyx of Held in the same R192Q knock-in mice, even though the mutation caused a similar 6 mV hyperpolarizing shift in the Cav2.1 current-voltage relationship under square-pulse depolarizing voltage steps, the shift did not alter presynaptic Ca2+ influx through Cav2.1 channels during normal evoked APs (Inchauspe et al., 2010).  It was determined that the difference in effects between pyramidal cells synapses and the calyx of Held synapse in the R192Q mice were due to AP duration.  During short 1 ms AP durations typical of the calyx of Held, a hyperpolarizing shift in the activation voltage was insufficient to alter presynaptic  154 Ca2+ influx.  On the other hand, when the AP duration was prolonged by 1-2 ms to match those typical of cortical pyramidal neurons, the shift in the activation by the R192Q mutation was sufficient to cause greater presynaptic Ca2+ influx (Inchauspe et al., 2010).  Importantly, the AP duration at PF terminals closely resembles the normal, 1 ms APs at the calyx of Held (Sabatini and Regehr, 1996, 1997).  In this regard, a shift to more negative activation voltages observed under square pulse depolarizations does not likely translate to an increase in Ca2+ influx during the normal, short APs at PF terminals.  In contrast, Cav2.1 channels in the facilitated state have an enhanced Ca2+ influx during APs with durations of 1 ms or less (our results in Chapter 3 as well as (Chaudhuri et al., 2004; Chaudhuri et al., 2005; Chaudhuri et al., 2007; Kreiner et al., 2010)).  Overall, if the S218L mutation renders channels into a facilitated state resembling that achieved during CDF (as proposed in Chapter 3, see discussion in section 3.2.3), an enhancement of Ca2+ influx during APs at PF boutons in S218L mice relative to wild-type would be expected. A further potential limitation relates to our hypothesis that Cav2.1 CDF is a likely mechanism of short-term synaptic facilitation at other central synapses and that changes in Cav2.1 CDF by FHM-1 mutations likely alters synaptic efficacy at these synapses.  In the large calyx of Held, Ca2+-dependent enhancement of presynaptic Ca2+ currents through Cav2.1 accounts for upwards of 40% of the total facilitation of synaptic response during repetitive APs (Forsythe, 1994; Borst and Sakmann, 1998; Cuttle et al., 1998; Forsythe et al., 1998; Inchauspe et al., 2004; Xu and Wu, 2005; Muller et al., 2008), however, as mentioned, Ca2+ influx through Cav2.1 channels at the calyx of Held is unchanged in R192Q knock-in mice under conditions of normal, short AP firing (Inchauspe et al., 2010).  Furthermore, the authors showed that facilitation of synaptic transmission was not altered in the R192Q mice relative to wild-type under these same conditions.  This indicates that perhaps Cav2.1 CDF is fundamentally different in terms of the mechanism of facilitation at this particular synapse.  In fact, while there is some evidence that the CaS NCS-1 may mediate facilitation of P/Q-type currents at this synapse (Tsujimoto et al., 2002), a role for CaM is uncertain (Sakaba and Neher, 2001; Xu and Wu, 2005; Nakamura et al., 2008).  Further, it is possible that CaM mediated Cav2.1 CDF is a minimal component of facilitation due to the expression of Cav2.1 variants that do not exhibit this type of Cav2.1 CDF (e.g. Cav2.1 EFb/Δ47).  155 As such, the R192Q may not affect facilitation at the calyx of Held due to different mechanims of Cav2.1 facilitation at this synapse or perhaps due to the expression of Cav2.1 splice variants that have CDF unaffected by the R192Q mutation (such as the Cav2.1 –NP variant; Fig. 4.2).  In future studies it will be interesting to see whether the more severe effects of the S218L mutation on Cav2.1 CDF alter facilitation at the calyx of Held.  4.3.3 Future directions An important future study will be to identify the CaSs responsible for mediation of Cav2.1 CDF in PF terminals.  CaM is a primary candidate as its kinetic properties as a CaS correlate with the temporal profile of facilitation at this synapse (Atluri and Regehr, 1996b).  The involvement of CaM in short-term facilitation of PF-PC synapses would be best determined by recording EPSCs in PCs during paired stimuli of PFs in transverse slices of the cerebellar vermis.  During paired stimuli of PF, PC EPSCs show robust PPF (Atluri and Regehr, 1996b).  Recordings at wild-type synapses could be compared in the presence and absence of a potent CaM blocker such as N-(6-aminohexyl)-5-chloro-1- naphthalenesulfonamide (W7) (Tanaka and Hidaka, 1980).  If CaM is the dominant mediator of Cav2.1 CDF in presynaptic terminals of parallel fibres, then in the presence of W7, PPF of EPSCs would be expected to be reduced by approximately 25%. Another future study relevant to the work presented here would be to explore the effect of the R192Q and S218L mutations on LTP.  LTP is a use-dependent form of synaptic plasticity that has long been considered to play a critical role in motor learning (Teyler and Discenna, 1984).   There is evidence that LTP at the PF-PC synapse involves presynaptic processes that overlap with those underlying PPF (Schulz et al., 1995; Salin et al., 1996).   Since both the R192Q and S218L mutations reduce PPF at this synapse (Fig 3.4), if the underlying mechanisms of PPF and LTP overlap, then LTP may also be altered in these mice.  Furthermore, LTP at the PF-PC synapse is typically induced by eliciting high-frequency stimulations consisting of 10 trains (100 Hz stimulations given for 50 ms) delivered at 200 ms intervals for a total of 50 stimulations over 2 s (Schulz et al., 1994).  The periods of AP firing required to induce  156 LTP are long enough that CDI of Cav2.1 may also be relevant.  Strong CDI of Cav2.1 is evident during prolonged APs that last beyond 400 ms and in Chapter 3 we show CDI of Cav2.1 channels is significantly reduced by the S218L mutation (but not R192Q) during a 1 second train of APs delivered at 100 Hz (Fig. 3.1 and 3.2).  As such, if Cav2.1 CDI is also relevant to synaptic plasticity at the PF-PC synapses under conditions of prolonged AP firing as predicted by a model system (Mochida et al., 2008), then additional changes may be observed in LTP at PF-PC in S218L mice compared to R192Q and wild- type mice.  LTP in wild-type and mutant KI mice at the PF-PC synapse can be measured using extracellular field recordings in transverse cerebellar slices.  EPSPs can be evoked by orthodromic stimulation of the parallel fibres in the molecular layer of the cerebellar vermis.  Understanding the effects of FHM-1 mutations on LTP may provide valuable information regarding the role of cerebellar development and motor coordination in ataxia.  Further, elucidating the role of Cav2.1 channels in LTP would be an important advancement toward understanding the basic molecular mechanisms of long-term synaptic plasticity and motor learning in the cerebellum.       4.4 Conclusions 4.4.1 General Conclusions Overall, we have shown that Cav2.1 splice variants respond differently to FHM-1 mutations.  We showed that a single point mutation can cause significant gain-of-function effects in one Cav2.1 variant and have little or no effect (or even loss-of-function effects) in another Cav2.1 variant.  We also showed that under different firing conditions, Cav2.1 variants respond differently to FHM-1 mutations.  I conclude that the phenomenon of differential effects of missense mutations on Cav splice variants is a major underlying theme in the localized and episodic nature of FHM-1 and other Ca2+ channelopathies. Furthermore, we showed for the first time that FHM-1 mutations alter CaM mediated Cav2.1 CDF.  We showed that the occlusion of Cav2.1 CDF by two FHM-1 mutations results in a corresponding attenuation in short-term synaptic plasticity at an intact CNS synapse.  As such, I conclude that Cav2.1  157 CDF is an important mechanism of short-term synaptic plasticity at the PF-PC synapse, and perhaps other CNS synapses.  Furthermore, significant changes in PF-PC synaptic transmission by some FHM-1 mutations likely alter the precise balance between excitatory and inhibitory inputs onto PCs (and thus the fine-tuning of PC intrinsic pacemaking), and as a result, lead to cerebellar motor deficits including ataxia under some conditions.  4.4.2 Relevance to treatment of human disease Migraine headaches affect approximately 15% of the Western population (Stewart et al., 1992; Lipton and Stewart, 1994; Lipton et al., 2001; Henry et al., 2002).  However, the complicated genetics and physiology of migraine have slowed both the development of adequate treatments and our understanding of underlying disease mechanisms (Montagna, 2004).  FHM-1 however, is an autosomal- dominant inherited subtype of migraine with similar clinical features to typical migraine, and has served as a useful model to study pathogenic mechanisms of common forms of migraine (Ducros et al., 2001; Thomsen et al., 2002).  As such, understanding the mechanisms of FHM-1 pathophysiology provides information concerning the disease mechanisms that can hopefully be used to develop more effective migraine treatments.  The involvement of the Cav2.1 channel in migraine pathology make it a potential therapeutic target, although to date its ubiquitous expression and function in the CNS and PNS have led to concerns for the high risk of undesired side-effects.  Our work suggests that targeting specific Cav2.1 splice variants directly involved in migraine pathology could minimize undesired effects.  For example, we demonstrate above that CDF of Cav2.1 (-NP) and Cav2.1 (+NP) variants are differentially affected by FHM-1 mutations (Fig. 4.2).  These same channel variants show discrete pharmacological sensitivities to peptide toxins that differ by more than 100 fold (Mintz et al., 1992; Wheeler et al., 1994).  Taken together, this suggests splice variants differentially affected by FHM-1 mutations may also be differentially targeted pharmacologically as a therapeutic approach.  This type of targeted approach to disease treatment could be a successful means of obtaining optimal positive results with minimal adverse reactions.  In fact, targeting specific Cav2.2 channel splice variants has become a leading strategy in  158 chronic pain therapy even though Cav2.2 channels are also ubiquitously expressed in the CNS (Snutch, 2005; Altier et al., 2007; Swayne and Bourinet, 2008).                      159      4.5 Bibliography Adams PJ, Snutch TP (2007) Calcium channelopathies: voltage-gated calcium channels. Sub- cellular biochemistry 45:215-251. Akerman S, Williamson DJ, Goadsby PJ (2003) Voltage-dependent calcium channels are involved in neurogenic dural vasodilatation via a presynaptic transmitter release mechanism. British journal of pharmacology 140:558-566. 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Zhuchenko O, Bailey J, Bonnen P, Ashizawa T, Stockton DW, Amos C, Dobyns WB, Subramony SH, Zoghbi HY, Lee CC (1997) Autosomal dominant cerebellar ataxia (SCA6) associated with small polyglutamine expansions in the alpha 1A-voltage- dependent calcium channel. Nature genetics 15:62-69.    165 APPENDIX 1: Ca2+ CHANNELOPATHIES: VOLTAGE-GATED Ca2+ CHANNELS*    _______________________________ * A version of this appendix has been published. Adams, P.J. and Snutch, T.P. (2007). Chapter 8: Calcium channelopathies: voltage-gated calcium channels. Subcell Biochem 45:215-251. Reprinted with kind permission of Springer Science and Business Media. All rights reserved.   166  167  168  169  170  171  172  173  174  175  176  177  178  179  180  181  182  183  184  185  186  187  188  189  190  191  192  193  194  195  196  197  198  199  200   201   202  APPENDIX 2: OTHER PHD PUBLICATIONS   Besides my above publication that related directly to Cav2.1 alternative splicing and modulation, I also have contributed significantly to another publication during my PhD studies (see below). I helped design and performed all PCR and Genescan experiments and participated in the writing of the paper by Spacey et al. (2006). This study revealed a second gene locus for the autosomal dominantly inherited Paroxysmal nonkinesigenic dyskinesia (PNKD).  These results are the first to suggest that there are at least two different genes responsible for PNKD, implying its genetic heterogeneity.  S.D. Spacey, MD; P.J. Adams, BSc; P.C.P. Lam; L.A. Materek, BSc; A.J. Stoessl, MD; T.P. Snutch, PhD; and G.-Y.R. Hsiung, MD. (2006). Genetic heterogeneity in paroxysmal nonkinesigenic dyskinesia. Neurology. 2006 May 23;66(10):1588-90.   203  APPENDIX 3: UBC RESEARCH CERTIFICATES OF APPROVAL        204    205

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