Nicholas E. V. Foster

Transcription

1 Whole cell and single channel analysis of cloned T=type calcium channel functionally expressed in HEK cells Nicholas E. V. Foster Department ofneurology and Neurosurgery McGill University, Montreal September 2002 A thesis submitted to the Faculty ofgraduate Studies and Research in partial fulfillment ofthe requirements ofthe degree ofmasters ofscience 2002 Nicholas Foster

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3 Abstract 1 examined the kinetic properties ofmacroscopic and single channel currents produced by an isoform ofthe rat CaV3.1 T-type voltage-gated calcium channel, expressed in HEK 293 ceus. Whole ceu currents were measured in 2 mm extracellular Ca 2 +, and single channel activity was recorded from cell-attached patches in 120 mm Ba 2 +. We found that macroscopic rates ofactivation, deactivation, inactivation, and recovery from inactivation each have voltage-dependent and voltage-independent components. Single channels opened and closed rapidly in bursts before inactivating, with a mean open tirne of 1 ms, a mean intra-burst close time of0.3 ms, and a mean burst duration of5.5 ms. The conductance ofthese openings was 4.4 ps. We also observed occasionallarger and smaller openings that may indicate subconductance states. These data will further contribute to the understanding ofthe biophysical function and diversity ofthis calcium channel isoform. 1

4 Résumé J'ai examiné les propriétés kinétiques des courrants macroscopics et de canaux uniques produits par un isoforme du cannai calcique voltage-sensible, CaV3.1 du rat exprimé dans des cellules HEK-293. Les courrants macroscopiques ont été enregistrés dans une solution extracellulaire de Ca 2 + à 2mM, tandis que l'activité de courrant unique, dans une solution de Ba mM. Nous avons trouvé que les vitesses d'activation, de désactivation, d'inactivation et de récupération de l'état inactivé, ont tous des composants voltage sensible et des composants indépendents du voltage. Les cannaux uniques s'ouvrent et se referment subitement à plusieurs reprises avant d'inactiver; le temps moyen d'ouverture est de lms, le temps moyen entre périodes d'activité, de O.3ms, et une période d'activité moyenne de 5.5ms. La conductance de ces ouvertures est de 4.4pS. Nous avons également observe des ouvertures de durées variées ce qui pourrait indiquer des états de sous-conductance. Ces résultats contribuent a la compréhension du fonctionnnement biophysique et de la diversité de cet isoforme du cannai calcique. II

5 Contents,, RESUME e1lllileltillilloeeeegiliogijeeeillgliiglilgliieeelllillgo'sollleeeaeeeillgclee0,,&ooe018llllllilldelloogo/il<llelitollolllllle0l00ltgooglileeegog000eeoogeeeeillgolilijltllllllilllileeeellllllgiilgolil" LIST OF FIGURES AND TABLES FREQUENTLY USED ABBREVIATIONS ACKNOWLEDGEMENTS V VI VII INTRODUCTIONolieeee"tilGoeee6eOIllGGGseeeeoeeeeeelllGGGeQe0GltIIl"IOlillllelilOllooooe"eeDooooe0eOOOO12l GeeeeOilleeellloeelllleeoGoeeeelil0GGGGCl 1 VGCCs AND CALCIUM FUNCTION 1 THE VOLTAGE-GATED CALCIUM CHANNEL FAMILY 2 CALCIUM CHANNEL S1RUCTURE 3 CALCIUM CHANNEL GENES 5 FUNCTIüNAL SIGNIFICANCE OF DIFFERENT T -TYPE CHANNEL SUBTYPES 6 T CHANNEL EXPRESSION 7 T CHANNEL ELECTROPHYSIOLOGY 8 MOLECULAR AND FUNCTIONAL DIVERSITY OF T CHANNELS 10 GOALS OF PRESENT RESEARCH Il METH0 DS 08111> CI III OOOGeee 1!I001llGOe00eGO Ille ".0111&0 elll 00l1li&00 1IlI0ee elll O ctollllilgo.'i"tlllleoeo.elll~u~ldeoooolleee0"iiilgoee00 (Il liloelloe IIIOee0Q8\11&00/l" 0 elllil 12 CELL CULTURE AND DNA TRANSFECTION 12 ELECTROPHYSIOLOGY 12 STIMULUS PROTOCOLS AND DATA ANALYSIS 13 Whole cel! activation threshold, activation kinetics, and inactivation kinetics 14 Window Current 14 Whole cel! deactivation kinetics 14 Whole cel! kinetics ofrecoveryfrom inactivation 15 Single channellatency to first opening and dwel! time 15 DATA FITTING AND CALCULATIONS 16 Boltzmann fits 16 Exponentialfits 16 Single channelfirst latency 17 Single channel open amplitude fits 18 Single channel dwel! time fits 18 WHOLE CELL ANALYSIS 19 SINGLE CHANNELANALYSIS 22 T CURRENT KlNETICS 26 LITERATURE COMPARISON 28 III

6 SUMMARY AND CONCLUSIONS 32 IV

7 List of Figures and Tables FIGURE 1. T CHANNELMOLECULAR STRUCTURE. 34 FIGURE 2. T CHANNEL ACTIVATION AND INACTIVATION THRESHOLDS. 36 FIGURE 3. VOLTAGE-DEPENDENCE OF ACTIVATION TIME-TO-PEAK. 38 FIGURE 4. VOLTAGE-DEPENDENCE OF MACROSCOPIC DEACTIVATION. 40 FIGURE 5. VOLTAGE DEPENDENCE OF MACROSCOPIC INACTIVATION. 42 FIGURE 6. VOLTAGE DEPENDENCE OF MACROSCOPIC RECOVERY FROM INACTIVATION. 44 FIGURE 7. SINGLE CHANNEL ACTIVITY FROM A CELL-ATTACHEDPATCH. 46 FIGURE 8. LATENCY TO FIRST OPENING. 48 FIGURE 9. SINGLE CHANNEL MEAN DWELL TIMES. 50 TABLE 1. ELECTROPHYSIOLOGICAL PROPERTIES OF CAV3.1 SPLICE ISOFORMS. 52 V

8 freq~ently Used Abbreviations Ca 2 + CaV DRG HEK HVA IV Calcium Voltage-gated calcium channel Dorsal root ganglion Human embryonic kidney cell High-voltage activated CUITent-Voltage K+ Potassium LVA Mg 2 + Na+ V'li VGCC Low-voltage activated Magnesium Sodium Voltage ofhalf-activation or half-inactivation Voltage-gated calcium channel VI

9 Ack~owledgeme~ts Thanks to Dr. David Ragsdale for bis unending guidance, support, and optimism throughout the duration ofmy studies in bis laboratory. This thesis also owes much to my advisory cornrnittee-dr. Ragsdale, Dr. Wayne Sossin, and Dr. Angel Alonso-whose suggestions and feedback helped shape my project from the outset. Hong-Ling Li and Laurence Meadows both helped me leam the ropes in the laboratory; Laurence's generous help, advice, patience, and humor was especially important to my growth and enjoyrnent while working in the labo Summer student John Sussman perforrned the CaV3.1 subcloning described in the Methods section. Fellow graduate student Andrew Loukas graciously provided a French translation ofthe abstract for this thesis. FinaIly, thanks to ail the members ofthe Ragsdale Lab-Laurence, Hong-Ling, Takeshi, Andrew, Lindsey, John, David H., Jane, and Lyn-and my other colleagues at the Montreal Neurological Institute for their help and for making such a friendly place to work. VII

10 Introduction T-type voltage gated calcium (Ci+) channels are found in many regions ofthe brain, in the peripheral nervous system, and certain non-excitable tissues. Currents evoked by T channels are important for rhythmic and spontaneous neuronal activity in the CNS and the heart (reviewed in Huguenard 1996). The principal characteristics that distinguish T type channels from other voltage-gated calcium channels (VGCCs) are activation and inactivation at negative voltages-typicaily near the resting membrane potential-as weil as slow channel deactivation (Kostyuk 1999). VGCCS AND CALCIUM FUNCTION The study oflow-voltage activated calcium channels builds on decades ofion channel research, beginning with potassium (K+) and sodium (Na+) membrane conductances in neurons (Llinas 1988). VGCCs, voltage-gated Na+ channels, and voltage-gated K+ channels form a supergroup sharing certain molecular and functional similarities (Jan et al. 1990). Thus, while the molecular structure and function ofvgccs remains less understood than K+ and Na+ channels, similarities among the cation channels can give us an idea ofhow calcium channels may perform functions such as voltage sensing, channel gating, ion selectivity, and interaction with other molecules. The VGCC family comprises the high voltage activated (HVA) calcium channelsc1assified as L-type, N-type, P/Q-type, and R-type-and the low voltage activated (LVA), or T-type, calcium channels. As mediators ofinward calcium flux, VGCCs play a number ofimportant physiological roles. In addition to contributing to œil excitability, intracel1ular calcium signais can drive modulation ofintracellular molecules via factors such as calmodulin, control ofgene transcription via cyc1ic AMP response element 1

11 binding protein (CREB), neurotransmitter release and other secretion, initiation ofmuscle contraction, influence on synaptic plasticity, and promotion ofceu growth (reviewed in Bootman et al. 2001; Evenas et al. 1998). VGCCs can mediate rapid local calcium signais at the cell membrane on a millisecond timescale. The complex morphology ofneuronal dendrites makes this type oflocal, subceuular Ca 2 + signaling especially important (Bootman et al. 2001). Inward bursts ofcalcium such as these can travel by diffusion or may propagate in waves (Evenas et al. 1998) within or among ceus, amplified by regenerative processes (Bootman et al. 2001). THE VOLTAGE-GATED CALCIUM CHANNEL FAMILY T currents, so-named for their transient time-course and tiny single channel amplitude, were first observed in cerebellar Purkinje neurons as a negatively activated current distinct from L-type calcium currents (Llinas et al. 1981). Around the same time, similar currents were found in DRG neurons (Carbone et al. 1984) and starfish eggs (Hagiwara et al. 1975). This negatively activated current was mostly inactivated near the resting potential and was revealed following a briefperiod ofmembrane hyperpolarization. In comparison, L-type channels activate at potentials mv more positive, and generate longer, more slowly inactivating currents (Hermsmeyer et al. 1997). N-type currents were discovered in DRG neurons and have intermediate activation threshold and inactivation rate characteristics compared to L and T channels (Nowyckyet al. 1985). In the brain, T currents are often associated with generation ofrhythmic neuronal activity (Huguenard 1996). In thalamocortical relay neurons, interaction oft current and a hyperpolarization-activated cation current (Ih) underlies firing pattern changes from 2

12 tonie spikes to phasie high-frequeney bursts when the resting membrane potential is hyperpolarized (MeCormick et al. 1990; Jahnsen et al. 1984). This bursting activity is dependent on a low-threshold spike, now known to be a result oft-type channels (Sherman 1996; Suzuki et al 1989). T-type currents have been implieated in the function ofthe brainstem respiratory network, which govems the phases ofmuscular contraction that drive breathing (Ramirez et al. 1996). T ehannels in the heart are believed to be involved in pacemaking, calcium signaling, and cell growth (Triggle 1997). The influence oft currents on cellular proliferation seems to be especially important in pathologie conditions. In adult tissues, cellular remodeling ofthe type that follows heart failure can be inhibited by the T channel blocker mibefradil (reviewed in Giles 1997). T channels are also expressed at higher levels during embryonic development (Day et al. 1998; Rohwedel et al. 1994; McCobb et al. 1989). Expression oft channels appears to be related to the cell cycle in sorne cells, being more prominent during the G 1 /S transition (Kuga et al. 1996). Increasing T currents may not be sufficient in and ofitselfto increase cell proliferation, however (Chemin et al. 2000). CALCIUM CHANNEL STRUCTURE Voltage-gated calcium channels are comprised ofan al subunit ofabout 190 kd, which forms the ion-conductive pore and can associate with one or more auxiliary subunits (Takahashi et al. 1987). Most ofthe protein resides in the cytoplasm, with less than 20% in the lipid ofthe cell membrane (Perez-Reyes 1999). The topology ofvgccs is likely to resemble voltage-gated sodium and potassium channels, due to similarities in their overall peptide sequence and especially the high homology ofthe transmembrane regions (Lee et al. 1999; Perez-Reyes 1999; Durell et al. 1998). The al subunit contains 3

13 four homologous domains (repeats I-IV), each having six transmembrane alpha-helices (S I-S6; see diagram in Figure la) (Catterall 2000). In HVA calcium channels, the linker sequences between domains contain regulatory sites where molecules sueh as auxiliary {3 subunits or G proteins can associate (Walker et al. 1998). For example, the I-II linker of HVA channels contains a {3 subunit binding site, which is absent in T channels studied to date (Lambert et al. 1997; Pragnell et al. 1994). It has been proposed that one or more of the domain linkers may also serve as inactivation gates, as with the Na+ channel III-IV linker (Miller et al. 1995). The six transmembrane segments in each domain are believed to cluster together with the S4 segment in the center (Cattera1l2000). The four domains come together so that a pore is formed inside, lined by the S5 and S6 segments. In L-type channels, binding sites for a number ofcalcium antagonists have been found in the S6 segment, consistent with those sites residing within the channel pore (Catterall 2000). The segments form the outside ofthe subunit (Perez-Reyes 1999). As in the voltage-gated Na+ channel, sensing ofmembrane potential is believed to reside in 84. This transmembrane segment contains a series ofpositively charged residues in a regular arrangement (every third amino acid) that probably results in their alignment within the alpha-helix and allows 84 to respond to the cell membrane potential, sliding and rotating out in the extracellu1ar direction at depo1arizing potentia1s (Durell et al. 1998). The conformational changes invo1ved in this movement wou1d underlie voltage-dependent open gating (activation) (Catterall 2000). Between the 85 and 86 segments in each domain is a membrane-associated pore loop ("P 100p"), which is believed to form the ion se1ectivity filter (Catterall 2000). The 4

14 channel pore is selective for divalent cations sueh as calcium and barium, and is blocked by a number ofmultivalent metal ions. These pore loop regions are the only part ofthe al protein predicted to extend into the extracellular space (perez-reyes 1999). Specifie amino acid residues in the pore loop vary with channel type and appear to be critical for detennining ion selectivity. A site within each ofthe four VGCC pore loops has been confinned by site-directed mutagenesis to be essential for calcium selectivity (Parent et al. 1995). In L-type channels ah four ofthese residues are glutamate (EEEE), whereas in the T-type channels two are aspartate (EEDD) (McRory et al. 2001; Perez-Reyes 1999). Furthennore, mutation ofthe residues in sodium channels from the nonnal DEKA to DEEE confers Ca 2 + penneability, underlining the specifie influence ofthese residues upon ion selectivity in voltage-gated ion channels (Heinemann et al. 1992). In T-type channels there is an additional extracehular loop (107 a.a.) between the S5 segment and P loop that is not present in HVA channels (Lee et al. 1999). Its proximity to the P loop may indicate that this extra loop influences ion selectivity. CALCIUM CHANNEL GENES There are at least 10 VGCC genes known at present, which code for the L-type (a1c, D, F, and S), non-l-type HVA (a1a, B, and E), and T-type (a1g, H, and 1) channels (Perez-Reyes 1999). The firstal subunit believed to fonn LVA channels was ale, which was cloned by Soong et al. (1993) and fonns channels having a lower threshold ofactivation than L-, N-, and P/Q-type channels. Later discovery ofalg (Perez-Reyes et al. 1998) and, subsequently, alh (Williams et al. 1999; Cribbs et al. 1998) and ah (Lee et al. 1999) demonstrated that these three channel subunits are responsible for T CUITent in the CNS, whereas ale (now known to fonn R-type channels) 5

15 has activation properties intermediate to T channels and RVA channels (Bean et al. 1998). alg, air, and ali each produce robust LVA currents when expressed in cultured ceu hnes without auxiliary subunits. A new nomenclature for VGCCs has recently been adopted, under which the alg, H and l subtypes are designated CaV3.l, CaV3.2 and CaV3.3, respectively, L-type channels (ais, alc, ald, alf) are classified as CaVI, and ala, alb, and ale form the CaV2 group (Perez-Reyes 1999). A human gene, CACNAlG, on chromosome 17q22, likely encodes the human CaV3.l (a1g) T channel, and a putative C. elegans calcium channel gene, c54d2.5, is related to the cloned mammalian CaV3 channels (Perez-Reyes et al. 1998). FUNCTIONAL SIGNIFICANCE OF DIFFERENT T-TYPE CHANNEL SUBTYPES a1g and air channels are similar in many oftheir electrophysiological properties (Klockner et al. 1999). Because they inactivate quickly and recover from inactivation slowly, T channels formed from the alg or air subunits are more tuned for low frequency repetitive activity «20 Rz) (Kozlov et al. 1999). ail channels have more distinct properties, including higher single channel conductance (Monteil et al. 2000b), faster deactivation (Klockner et al. 1999), and slower activation and inactivation (Monteil et al. 2000b). a 11 channels also have the fastest recovery from long-term inactivation (Klockner et al. 1999). These inactivation properties allow them to continue activating in neurons during higher frequency stimuli (Kozlov et al. 1999). The distinct properties oft channels compared to other VGCCs appear to be a result ofearly divergence from the RVA calcium channels, judging by analysis of sequence homology (Cattera1l2000; Perez-Reyes 1999). Many regions that are conserved among the HVA calcium channel al subunits are different in T channels, despite the 6

16 similarities in general structure and hydropathy pattern. T channel sequences cloned to date have about 25% homology with the L-type and non-l-type HVA channels (Catterall 2000). Regions having important amino acid similarities between T-type and non-t-type VGCCs include the extracellular pore loop and the S4 segment (Bean et al. 1998). general, the transmembrane segments are the most conserved regions among the LVA and HVA calcium channels (Lee et al. 1999). HVA calcium channels associate in vivo with one or more auxiliary subunits, designated {3, 'Y, and d2ô, which modulate channel behavior and in sorne cases are required for formation offunctional channels. However, the typical subunit arrangement for T channels is unknown. Initial studies suggested that T channels do not form functional associations with calcium channel auxiliary subunits (e.g. Lambert et al. 1997). More recent results have revealed that certain auxiliary subunit subtypes, including {31b, a2o-2, and 'Y-S, do modulate the expression and electrophysiology of specifie T-type al channels in vitro (Hobom et al. 2000; Klugbauer et al. 2000; Dolphin et al. 1999). The a2o-2 subunit increases alg current density, but decreases the contribution ofindividual alg channels by speeding current inactivation and shifting the steady-state inactivation threshold in the positive direction (Hobom et al. 2000). The 'Y-5 subunit speeds activation and inactivation ofalg (Klugbauer et al. 2000). These data suggest that known or yet to be identified auxiliary subunits may modulate T-type channel function in vivo. T CHANNEL EXPRESSION T channels transcripts are found in a wide range ofbrain tissues. The expression of alg, alh, and ali is often complementary (Talleyet al. 1999), suggesting that they play 7

17 different roles depending on brain area and cell type. algis expressed at the greatest levels in thalamic relay neurons, cerebellar Purkinje cens, in the inferior olive, and the stria terminalis. In sorne tissues the expressionofa1g is graded, such as the 1ateral habenula, where expression is greater rostrally and low in the caudal region. Within neurons, T channels are localized to the soma and dendrites (Huguenard, 1996). T currents are more prevalent and are found in a greater range oftissues during fetal stages, implying developmental control ofchannel expression (Day et al. 1998; Rohwedel et al. 1994; McCobb et al. 1989). T CHANNEL ELECTROPHYSIOLOGY One ofthe key distinguishing properties oft channels is their low threshold of activation (Huguenard 1996). Unlike HVA calcium channels and voltage-gated sodium channels, T-type and other LVA channels are typically activated near the neuronal resting membrane potential, opening after depo1arizations as small as 10 mv. Sodium channels require greater depolarizations (about -25 mv) (e.g. Qu et al. 2001), and HVA calcium channe1s have higher thresholds still, as positive as 0 mv (Hermsmeyer et al. 1997). Ionie currents near the resting membrane potential can affect the firing pattern in neurons (Kozlov et al. 1999). The lower threshold oft channels allows them to gate the function ofmore positively-activated channels. This can result in a lowered spike threshold, and is important for the function ofneurons in the thalamus and in olfactory receptor cells (Huguenard 1996). In addition to their LVA characteristics, T channels were originally distinguished on the basis oftheir transient, fast-inactivating currents, and steady-state inactivation at low potentials. The latter properties are no longer considered strict criteria for T-type 8

18 channels, after the discovery of slowly inactivating LVA currents in GABAergic neurons ofthe thalamic reticular nucleus (Huguenard et al. 1992) and ofthe CaV3.1 (a1i) isofonn, which inactivates at more positive potentials (Monteil et al. 2000b). Studyof whole-cell currents suggests that channel inactivation takes place primari1y from the open state (Serrano et al. 1999), and is mostly voltage-independent. Recovery from inactivation, in contrast, does not oœur via the open state (Serrano et al. 1999), and is dependent on channel deactivation (Kuo et al. 2001). A 'ball-and-chain' mechanism, similar to that ofshaker K+ channels, is believed to underlie fast inactivation ofthe T channel, based on study ofalc (L-type)/alG chimeras by Staes et al. (2001). In the Na+ channel, the domain III-IV linker is believed to function as an inactivation gate; this may a1so be true for T channe1s (Miller et al. 1995). T channels, like Na+ and Shaker K+ channels, display different modes of inactivation (Bossu et al. 1986). Fast inactivation iseasily observable in T channels as it causes the rapid decay ofcurrent during a prolonged depolarization. Slow inactivation, from which there is a correspondingly slow recovery, operates on a scale ofseconds and results in a pool ofinactive channels which can become available ifthe membrane potential remains hyperpolarized for a relatively long period. Considerable functional diversity has been found among the T channels studied to date, though certain criteria are still accepted as distinguishing T-type channels from HVA calcium channels: slow current deactivation, leading to a prominent taii current following depolarizing test pulses (Perez-Reyes 1999; Randall et al. 1997; Huguenard 1996); and a small unitary channel conductance, averaging around 8 ps in isotonie Ba 2 + or Ca 2 +(Perez-Reyes 1999). Now that several T-type channel genes have been cloned, 9

19 sequence homology can be used to detennine whether newly discovered"channels can be considered as T-type and thereby to distinguish which functional characteristics are unique to T channels. MOLECULAR AND FUNCnONAL DIVERSITY OF T CHANNELS Beyond the three types oft channel (alg, alh, and au) there is a furtherrange of channel diversity due to alternatively spliced channel isofonns. For each LVA channel type multiple splice variants have been found, which can vary both in their pattern of expression and their biophysical properties (Chemin et al. 2001). This range ofvariation could explain the electrophysiological disparity observed among sorne native T currents, especial1y in the kinetics and steady-state threshold ofinactivation (Huguenard 1996). CaV3.l channels have at least 38 exons, with alternative splicing identified at six sitesexons 14, 25B, 26, 34,35, and 38B (Mittman et al. 1999) (Figure 1). Mittman et al. predict that as many as 24 CaV3.1 isofonns could result from alternative splicing at six sites within the sequence, producing proteins between amino acids in size (Mittman et al. 1999). So far at least 7 ofthese putative CaV3.1 isofonns have been identified in neuronal tissues to date, in human (Chemin et al. 2001; Cribbs et al. 2000; Monteil et al. 2000a), rat (McRory et al. 2001; Zhuang et al. 2000), and mouse (Satin et al. 2000; Klugbauer et al. 1999). Functional differences among the isofonns inciude activation threshold, steady state inactivation threshold, and voltage dependence ofthe activation and inactivation rates. A study ofseveral alg splice variants demonstrated that T channel activation threshold and kinetics are greatly determined by amino acid residues in the III-IV linker, where the sequence varies due to alternative splicing (Chemin et al. 2001). Variation in inactivation properties is due in part to alternative 10

20 splicing within the II-HI linker; an insertion, designated e, shifts the steady-state inactivation curve negative and speeds the inactivation rate at higher potentials (Chemin et al. 2001). GOALS OF PRESENT RESEARCH Measurements ofthe gating kinetics ofc10ned T-type channel isoforms have focused almost exc1usively on macroscopic currents, whereas single channel characterization ofc10ned T channel subunits has thus far been limited. In particular, currents ofthe rat alg isoform c10ned by Zhuang et al. (2000) have not been studied in mammalian cells on the macroscopic or single channellevel. Our goal was to describe the kinetic parameters oftms T channel isoform in cultured heterologous cells. We have examined the rates ofactivation, deactivation, inactivation, and recovery from inactivation from recordings ofwhole cell and single channel currents. We used HEK 293 cells for channel expression due to their negligible levels ofendogenous currents which could interfere with the measurement oft currents. The behavior ofcalcium currents is affected by extracellular calcium levels (Wilson et al. 1983), so we performed our whole-cell recordings at physiological (2mM) extracellular calcium concentrations. Our results give insight into the biophysical function and roles ofthis T channel isoform, and help to fill in a picture ofthe electrophysiological variability among CaV3.1 channels. 11

21 Methods CELL CULTURE AND DNA TRANSFECTION Human Embryonic Kidney cells (HEK-293, from American Tissue Culture, Manassas, Virginia, USA) were grown in plastic dishes in DMEM and fetal bovine serum (10%), penicillin ( unitshtl) and streptomycin ( units/ill). Trypsin (0.25%) with 1 mm EDTA was used to passage the cens when confluent. AH cell culture products were purchased from Gibco-BRL (Burlington, ON, Canada). The rat CaV3.1 isoforrn cdna (Zhuang et al. 2000) was obtained from Ming Li and subcloned into the pcdna hygro(-) vector (Invitrogen, Burlington, Ontario, Canada). Proper insertion ofthe gene was verified by sequencing across ligation sites. For single-channel recording, cells were transiently transfected with a pcdna3.1 hygro(-) vector containing the rat CaV3.1 gene subunit and a second vector containing the green fluorescent protein (GFP) gene (pegfp-l, from Clontech, Palo Alto, CA, USA) in an 8:1 ratio. Transfections were perforrned using the FuGene kit (Roche Diagnostics Canada, Laval, Quebec, Canada) according to the manufacturer's instructions. For whole-ceh recording a stably-transfected Hne ofhek-293 cells was prepared using the pc-dna3.l/cav3.1 construct and selecting with Ilg/mL Hygromycin B (Gibco-BRL). ELECTROPHYSIOLOGY Whoie-cell experiments were perforrned using an extracellular bath solution containing 2mM Ca 2 + as the charge carrier. This solution contained (in mm): CaCh, 2; TEA-Cl, 140; CsCI, 6; HEPES, 10; ph 7.2. The intracellular (pipette) solution contained (in mm): NaCl, 5; MgClz, 2; L-aspartic acid, 120; EGTA, 10; HEPES, 10; ph 7.2. Single 12

22 channel expenments were performed using a 120 mm BaH intracellular solution containing (in mm): BaCh, 120; HEPES, 10; ph 7.4. A high-potassium (140 mm) extracellular solution was used during cell-attached patch recordings. It contained (in mm): KAc, 140; NaCl, 5; MgCh, 4; CdClz, 0.2; HEPES, 10; glucose, 25; ph 7.4. AH recordings were performed at room temperature following incubation ofcells at 37 C for hours. Glass electrodes were puhed from borosilicate micro-hermatocrit capillary tubes (Fisherbrand, Fisher Scientific, Ontario, Canada) using a Sutter P-97 puller and frre-pohshed using a Micro Forge MF-830. For recording in the cell-attached patch configuration, the electrodes were coated with Sylgard (Dow Coming, Midland, MI, USA) in order to reduce noise. Electrodes generally had a resistance in the range of 2-4 Mühms. Currents were recorded under voltage clamp conditions using a CV203BU headstage (Axon Instruments, Foster City, CA, USA) and amplified and anaiog filtered at 5 khz using an Axopatch 200B amplifier (Axon Instruments). Cell-attached patch recordings from one cell were filtered at 2 khz. Linear capacitance currents were canceled using the internai voltage clamp circuitry. ResiduaIIinear currents were subtracted using the P/4 procedure (Armstrong and Bezanilla, 1977) during whole-cell recordings. SignaIs were Iow-pass filtered at 5 khz (-3 db) using the amplifier's internai circuitry, digitized with a Digidata or BOO-series, and acquired using pciamp software (Axon Instruments) at a 10 khz sampling rate. STIMULUS PROTOCOLS AND DATA ANALYSIS AH whoie-ceh traces were Gaussian filtered with a -3 db cutoffof1500 Hz in software with Clampfit (Axon Instruments) and anaiyzed using CIampfit, Origin (MicrocaI, Northampton, MA, USA), and SigmaPIot (SPSS Ine., Chicago, Illinois, USA) 13

23 software. Recordings from untransfected HEK cells were made to verify the absence of endogenous calcium CUITents. WHOLE CELL ACTIVATION THRESHOLD, ACTIVATION KINETICS, AND INACTIVATION IaNETICS The activation threshold, activation rate (time to peak) and inactivation time course ofwhole cell CUITents were measured using depolarizing pulses over a range of potentials. Pulses of75 ms duration were made from a holding potential of-90 mv to potentials between -70 and 35 mv in 5 mv increments. These stimulus sweeps were delivered at a rate ofone every 7 seconds. We determined the whole-cell conductance from these recordings by dividing CUITent peakvalues bythe driving force. The driving force was calculated by fitting the linear portion ofthe IV curve with a linear function. WINDOW CURRENT We estimated the whole-cell window CUITent (i.e. steady-state CUITent) from the steadystate inactivation and macroscopic conductance curves, and the driving force. The normalized steady-state inactivation curve in Figure 2C was multiplied by the normalized macroscopic conductance to calculate steady-state conductance over the same range of potentials. This curve was multiplied by the driving force (see above) to obtain steadystate CUITent values. WHOLE CELL DEACTIVATION KINETICS The macroscopic deactivation rate was measured from tail CUITents evoked by repolarization ofthe cell membrane potential from 0 mv to more negative potentials. From a holding potential of-90 mv, the cell membrane was first depolarized to 0 mv for 14

24 15 ms, then repolarized to a potential ofbetween -20 and -140 mv for 60 ms. Stimulus sweeps were delivered at a rate ofone every 7 seconds. WHOLE CELL KINETICS OF RECOVERY FROM INACTIVATION The recovery ofwhole eeu CUITent from inactivation was measured using a stimulus protocol that first inactivated virtuauy ah the whole ceu current with a depolarizing pulse, then auowed recovery from inactivation with a variable-duration repolarization to the holding potential, then assessed recovery with a second depolarizing pulse. The first depolarizing pulse was to -30 mv, for 161 ms; this duration was long enough that the whole ceu CUITent appeared to decay to zero, and there was no observable tail CUITent upon the subsequent repolarization. The duration ofthe delay between the two depolarizing pulses was varied among sweeps in a trial, increasing from 5 ms to 455 ms. The holding potential, applied between sweeps and between the depolarizing pulses, was varied among trials from -100 mv to -60 mv. At the more hyperpolarized voltages we tested holding potentials at 5 mv intervals. Above -70 mv we tested some potentials at 2 mv intervals due to the greater difference in recovery time in this voltage range. Stimulus sweeps were delivered at a rate ofone every 10 seconds. SINGLE CHANNEL LATENCY TO FIRST OPENING AND DWELL TIME Single channel analysis was performed using Fetchan and pstat (Axon Instruments), Excel (Microsoft, Redmond, WA, USA), and Origin (Microcal) software. Single channel recordings were filtered at 1000 Hz (-3dB) in Clampfit. Currents were evoked with 60ms pulses from a holding potential of-100 mv to -50, -40, and -30 mv, at a rate ofone sweep every 3 seconds. Leak CUITents and capacitive transients were subtracted off-line using averaged sweeps or segments ofsweeps containing nuu-events. 15

25 Baseline drift among sweeps was minimized by manual adjustment in Clampfit when necessary. Opening and closing events were detected using a standard "50% threshold" method. Events were omitted from analysis when noise, superimposed openings, or drift ofthe baseline made opening and closing events ambiguous. First latency distributions were corrected for the number ofchannels in a patch (n) by taking the n-th root ofeach data point. DATA FITTING AND CALCULATIONS BOLTZMANN FITS The steady-state inactivation and whole cell conductance data were fit with the Boltzmann equation: In this equation, Xo is the midpoint ofthe curve (Vy,) and k is the slope factor. Because the data were normalized before fitting, the values ofal and Az were AI=1 and A 2 =0 for steady-state inactivation, and AI=O and Az=1 for the whole cell conductance. EXPONENTIAL FITS Most voltage-dependent and time-dependent data were fit with exponential equations, as noted in the text. Data that increased with x were fit with one ofthe fol1owing equations: Single exponential: Biexponential: F(x)- A. e(xjt,) + A. e(xjtz) + y

26 where t values are time constants, A values are scaling factors, and Yo is the y offset. Data that decreased withx were fit with one ofthe fol1owing equations: Single exponential: F(x) =A e(-x/t) + Yo Biexponential: F(x) =Al. e(-x/t 1 ) + A z e(-x/t 2 ) + y 0 using the same variables as above. SINGLE CHANNEL FIRST LATENCY The number ofchannels in each patch was determined from the greatest number of superimposed openings observed while recording from that patch. Because having more than one channel in a patch will skew the distribution offirst latencies, the first latency histogram data were corrected for the presence ofn number ofchannels in a patch by taking the Nth root-e.g. for a patch having 4 channels, each bin was transformed using the foilowing equation: bincount corrected= Vbin count The frequency distribution offirst latencies was plotted as a cumulative histogram for each ceil, and was fit with an exponential function using the Simplex least squares method: where P is the proportion for each term, and T is the time constant. 17

27 SINGLE CHANNEL OPEN AMPLITUDE FITS The distribution ofsingle channel opening amplitudes was fit with a Gaussian function, using the Simplex least squares method: where Il is the mean amplitude, and a is the standard deviation. SINGLE CHANNEL DWELL TIME FITS The frequency distributions ofsingle channel opening, c1osing, and burst durations were fit with an exponential decay function using the Simplex least squares method: where P is the proportion for each term, and 7 is the time constant. 18

28 Results WHüLE CELL ANALYSIS We began our analysis ofthis T channel isofonn by assessing the properties of macroscopic currents in HEK-293 ceus expressing the cloned CaV3.1 cdna. Figure 2A shows typical whole-cell currents recorded in 2mM Ca 2 + using an IV stimulus protocoi. The T currents became active at about -60 mv and reached a peak amplitude around -30 mv. The current-voltage (IV) relationship ofcurrents from the same ceu is shown in Figure 2B. The activation curve in Figure 2C reflects the averaged IV curves from 25 ceus, divided by the driving force to obtain whole cell conductance. A Boltzmann fit to the activation curve gives a VYz value of-42.9 mv with a slope factor (k value) of4.6 mv. The steady-state inactivation level was measured from the peak current during a -25 mv test pulse fouowing a 220 ms prepulse over a range ofpotentials. The resulting curve in Figure 2C, averaged from 19 celis, reflects the fraction ofchannels available in a noninactivated state at each membrane potential The inactivation VYz value was mv with a kvalue of6.0 mv. The overlap ofthe steady-state inactivation curve with the activation curve around -50 mv indicates the presence ofwindow current, a steady-state equilibrium ofactivated channels that can be important in detennining neuronal excitability near the resting membrane potential (Hennsmeyer et al. 1997). We estimated the window current amplitude from the inactivation and activation curves (Figure 2D). The predicted peak window current is about -78 pa, at -47 mv. We characterized the kinetic behavior ofthis T channel by examining the rates of current activation, deactivation, inactivation, and recovery from inactivation from whole- 19

29 celi CUITents. The time-to-peak (Figure 3) reflects how fast channels open upon membrane depolarization. The timing ofthe CUITent peak is affected to a 1esser extent by rates ofchannel inactivation and deactivation, but provides a good measure ofthe rate of channel activation. We applied depolarizing pulses using the same stimulus protocol as described above for measuring the IV and activation curves. Activation was slow at negative potentials and became progressively faster at progressively more depolarized potentials. A monoexponential fit to the data gave an e-fold increase in activation rate per 14.0 mv (Figure 3B). The time-to-peak reached an asymptote of3.1 ms above about 10mV. T channels characteristically produce a prominent tail CUITent upon repolarization from a depolarizing pulse, indicating slow closing (deactivation) ofopen channels. The macroscopic deactivation rate can be examined by measuring the rate ofcuitent relaxation following a hyperpolarizing step in membrane potential. We elicited tail currents by stepping the membrane potential from 0 mv to voltages between -20 and -140 mv. Tail current amplitude decayed exponentially and was weil fit by a single exponential function at eachpotentia1 (Figure 4A). The first 1-2 ms ofthe tai1 CUITent was not included in the fit to avoid eitors caused by filtering or voltage clamp settling time. The mean time constants (taus) from 16 cells are plotted in Figure 4B. Between -135 and -60 mv the deactivation rate slows exponentially at e-fold per 32 mv, then appears to approach a constant 13 ms above about -30 mv. This leveling offat higher potentials probably is due to CUITent inactivation, which reaches similar rates in this voltage range (see be1ow). 20

30 Whole-cell inactivation is conventionally measured from the CUITent decay during a depolarizing voltage step. The rate ofcuitent decay also depends on the rates ofchannel activation and deactivation, which are voltage-dependent over a similar range of potentials as channel inactivation. For measurement ofthe macroscopic inactivation rate we used the same stimulus protocol as described above for the IV and activation time to peak. CUITent decay followed a monoexponential function, giving a single time constant (Figure SA). Above -40 mv the inactivation rate remained constant at about 15 ms, and decreased and more negative potentials. The voltage dependence ofthe inactivation rate was approximately exponential, increasing e-fold per 7 mv. Inactivation is thus the most voltage-dependent ofthe macroscopic kinetic rates we measured for this T channel. Recovery from inactivation is a measure ofhow fast channels leave the inactivated state to enter a closed, available state. This process becomes active below the activation threshold. We measured the. rate ofrecovery from inactivation using a two-pulse protocol. First, a 160 ms depolarizing pulse to -30 mv was applied, which inactivates virtually all the channels. The membrane is then held at a more negative potential for a duration ofbetween 5 and 255 ms; during this period the recovery process is active because ofthe hyperpolarizedpotential. A secqnd pulse to -30 mv is then applied. If there has been no recovery ofinactivated channels to a closed state then no channels will be available to open during the second depolarization and no CUITent is observed. At increasingly longer interpulse durations a greater fraction ofthe cel1's channels is able to recover from inactivation, and the CUITent response becomes greater. The relationship of the CUITent peak during the second depolarizing pulse to the interpulse duratiotl follows an exponential function (Figure 6B). We repeated this protocol using inter-pulse holding 21

31 potentials between -100 and -60 mv to measure the voltage dependence of recovery from inactivation, which follows a monoexponential curve (Figure 6C). The recovery time constant slowed e-fold per 16 mv from a voltage-independent asymptote of ms below OOmV. SINGLE CHANNEL ANALYSIS Recording in the cell-attached patch configuration allowed us to measure the currents ofindividual T channel openings. Using 120 mm Ba 2 + as the charge carrier, we applied 60 ms pulses to -50, -40, and -30 mv from a holding potential of-100 mv. We obtained recordings from four patches among four different cells. Each patch contained between 4 and 6 channels. At ail three potentials channels tended to open in bursts, flickering closed one or more times before inactivating (Figure 7A-B; -50 mv not shown). Openings were occasional at -50 mv and robust at -30 mv. At -30 mv, superimposed openings by multiple channels were common. Single openings ranged from about 0.37 pa at -30 mv to 0.46 pa at -50 mv. The slope conductance was 4.37 ps (Figure 7E). We also observed occasional openings to a smaller amplitude (about hait) and a larger amplitude (about double); see Figure 7C. We expect these openings correspond to difference conductance states, as observed in T channels by others (Chen et al. 1990; Carbone et al. 1984). We chose to focus our single channel analysis on the 4.4 ps conductance state because these openings were most abundant in our recordings. The kinetics ofnative T channels have been characterized in a number ofneuronal cell types (e.g. Chen et al. 1990; Carbone et al. 1984). We examined the kinetic rates of channel opening, closing, and inactivation in the present channel isoform by measuring the mean latency to first opening, open dwell time, closed dwell time, and burst durations 22

32 at -40 and -30 mv. Above -30 mv, superimposition ofchannel openings combined with the smaller opening amplitudes made disceming individual channel events too difficult for analysis. Openings at -50 mv were too infrequent foranalysis at that potential. The latency to first opening is a measure ofhow fast channels open upon suprathreshold depolarization. It is analogous to the macroscopic activation rate measured by the time-to-peak; however, the latency to first opening is a more direct measurement ofthe state transitions involved single channel gating. The distribution of frrst latencies at -30 and -40 mv is shown in Figure SA-B. Because there were multiple channels in each ofthe four patches in our recordings, we corrected the distribution data as described in Methods. The distributions at both potentials were best fit by biexponential functions. The two time constants probably reflect rates ofdifferent state transitions between the closed (resting) channel state and the open state. At both potentials the faster time constant had the greatest contribution to the first latency distribution. Both time constants show voltage dependence (Figure SC); the faster time constant decreases from about 2.6 ms at -40 mv to 1.S ms at -30 mv, and the slower component decreases from 339 ms at -40 mv to 10.S ms at -30 mv. The slow time constant at -40 mv is considerably slower than the other three time constants, and may reflect a less-accurate fit due to fewer openings at this potential. The rates oftransitions between closed, open, and inactivated states can be measured from single channel open durations, burst durations, and intra-burst closing durations. Measurements ofail three duration types formed exponential distributions, allowing estimation ofmean dwell times using exponential fits to the data (Figure 9A-C). The mean open time was about 1 ms at both -40 and -30 mv (0.94 ms at -30 mv,

33 ms at -40 mv). This duration reflects the transition rate from the open channel state to the nearest closed state. Likewise, the durations ofclosings within bursts is dependent on the rate ofthe closed-to-open state transition. Because our closed time measurements included both intra-burst closings and inter-burst closings, we fit the closed time distribution with a biexponential function in order to separate the intra-burst closings (fast exponential term) from those between bursts (slow exponential term). The closed time within bursts was considerably shorter, making this method ofdata separation practical. The mean closing time within bursts was about 0.35 ms at both -40 and -30 mv and did not differ significantly between the two potentials (0.34 ms at -30 mv, 0.37 ms at-40 mv). The duration ofbursts gives an indication ofhowfast the open-to-inactivated transition is relative to open-to-closed-a slower rate for the inactivating transition would result in bursts oflonger duration, but with unchanged durations ofthe intra-burst closings and openings. The bursting activity oft channels implies that the transition from open to closed has a greater probability (and hence rate) than the open-to-inactivated transition. It also suggests that inactivation from closed states, or at least from the last closed state, is slower than the closed-to-open transition (ifthe rates were equal, over half ofail bursts would terminate after a single opening). In order to measure the durations of bursts we chose a threshold closing time of3.0 ms (10 times the mean intra-burst closing time, to minimize the chance oflonger bursts being counted as multiple, shorter bursts). Closings ofa duration above this 3 ms threshold were considered to separate bursts, while those below the threshold were considered intra-burst closings. This threshold worked weil to resolve bursts ofambiguous demarcation. The mean burst length was 24

34 about 5.4 ms for -30 and -40 mv (5.41 ms at -30 mv, 5.46 ms at -40 mv). Statistical comparison ofopen, closed, and burst durations between -40 and -30 mv did not reveal a dependence on membrane potential (Student's t-test; open durations, P=0.79; closed durations (fast tau), P=0.80; burst durations, P=0.92). 25

35 In this thesis, 1have explored the macroscopic and single channel kinetics ofa rat T-type calcium channel isoform expressed inhek-293 cells. This is the first characterization ofthis CaV3.1 isoform in HEK cells and on a single channellevel. The currents we observed from expression ofthis channel in HEK cells are similar to those reported by others for other CaV3.1 isoforms, and to native T currents in the nervous system. T CURRENT KINETICS Three ofthe macroscopic rates we examined-activation, inactivation, and recovery from inactivation-were voltage-dependent over a range ofpotentials, but approach voltage-independent asymptotes at either positive or negative potentials. In the case of both activation and recovery from inactivation, the whole cell current refiects channels that have gone through multiple state transitions before opening-for example, in the case ofactivation, from a resting closed state, through intermediate closed states, then to an open state. At positive or negative voltage extremes, a voltage-independent transition can become rate limiting. Thus we believe the rate limits indicate that among the series of single channel state changes that underlie macroscopic currents are both voltagedependent and voltage-independent transitions. Our single channel results agree with this interpretation ofchannel activation. The latency to fifst opening, which depends on the rates ofan transitions between the resting closed state and channel opening, was faster at -30 mv than -40 mv and had two rate constants. On the other hand, the intra-burst closing times, which refiect just the final closed-to-open state transition rate, were not dependent on voltage, even though the whole cell time-to-peak decreases by almost 50% 26

36 at the same potentials. Therefore, as others have reported (Chen and Hess, 1990), our data suggest that the final transition to the open state is voltage independent while one or more ofthe intermediary c1osed-state transitions are voltage independent. Inactivation ofcav3.1 channels is believed to occur primarily from the open state (Serrano et al. 1999). The macroscopic inactivation rate, unlike activation and recovery from inactivation, is probably the result ofa single transition, i.e. from the open state to a single absorbing inactivated state. A pure measurement ofthe inactivation rate should therefore reveal either a voltage-dependent or voltage-independent relationship, but not both. On the single channellevel, we did not find voltage-dependence in the burst duration between -40 and -30 mv, implying that the inactivation rate is the same over this voltage range. Another study examining whole cell currents using different stimulus protocols also conc1uded that channel inactivation is voltage-independent (Serrano et al. 1999). We believe the voltage-dependence seen in inactivation ofmacroscopic currents is a result ofvoltage-dependent activation, as suggested by Chen and Hess (1990). One complication in this interpretation, however, is the fact that the macroscopic inactivation rate in CaV3.1 channels is more dependent on membrane potential than activation time, regardless ofwhether the macroscopic activation rate is measured by time-to-peak (as we have done) or with exponential fits (see Table 1). For example, we found that below -40 mv the macroscopic inactivation rate slows at about e-fold per 7 mv. Activation rate slows by onlyel14 mv. Since the rate at which CUITent rises after depolarization depends not only on channel activation time but also on how fast open channels deactivate or inactivate, the voltage-dependence observed in the macroscopic inactivation curve may 27