A MUTATION IN CaV2.1 LINKED TO A SEVERE NEURODEVELOPMENTAL

DISORDER IMPAIRS CHANNEL GATING

by

SIDHARTH TYAGI

B.A, University of Colorado, 2017

A thesis submitted to the

Faculty of the Graduate School of the

University of Colorado in partial fulfillment

of the requirement for the degree of

Master of Science

Department of Integrative Physiology

2019

This thesis entitled:

A MUTATION IN CaV2.1 LINKED TO A SEVERE NEURODEVELOPMENTAL

DISORDER IMPAIRS CHANNEL GATING

Written by Sidharth Tyagi

Has been approved for the Department of Integrative Physiology

________________________________________

Roger Bannister, PhD

________________________________________

Roger Enoka, PhD

________________________________________

Robert Mazzeo, PhD

Date _________________

The final copy of this thesis has been examined by the signatories, and we find that

both the content and the form meet acceptable presentation standards of scholarly work

in the above-mentioned discipline.

iii

ABSTRACT

Tyagi, Sidharth (M.S., Integrative Physiology)

A mutation in CaV2.1 linked to a severe neurodevelopmental disorder impairs channel

gating

Thesis directed by Assistant Professor Roger A. Bannister

Ca2+ flux via voltage-gated Ca2+ channels is essential to the regulation of

membrane excitability, neurotransmission, and a variety of intracellular signaling

processes. CaV2.1 is the predominant Ca2+ channel present in the presynaptic terminals

of both neuromuscular junctions and many central synapses. Point mutations in CaV2.1

can drastically alter channel gating and expression, and indeed have been linked to two

paroxysmal disorders – Episodic Ataxia Type 2, and Familial Hemiplegic Migraine Type

1. With the novel whole-exome sequencing technique, mutations linked to a new class

of more severe disorders have been found with phenotypes like Episodic Ataxia Type 2

with an additional developmental component. One of these mutations is an arginine to

proline substitution in the S4 voltage-sensing region of the fourth membrane-bound

Repeat of CaV2.1 (R1673P). This mutation was proposed to cause a gain-of-function in

CaV2.1 based on the ability of the mutant channel to rescue the photoreceptor response

in CaV2.1-deficient Drosophila cacophony larvae. Here, I show that the R1673P

mutation actually results in a profound loss-of-channel-function. Voltage-clamp analysis

of tsA-201 cells expressing the mutant channel revealed a ~25 mV depolarizing shift in

the voltage-dependence of activation coupled with delayed activation kinetics. These

alterations in activation implies that a significant fraction of CaV2.1 channels resident in

presynaptic terminals are unlikely to open in response to an action potential, thereby

iv

increasing the probability of synaptic failure at both NMJs and central synapses.

Indeed, the mutant channel supported only minimal Ca2+ flux in response to an action

potential-like waveform. Application of the CaV2.1 agonist GV-58 shifted mutant

activation to more hyperpolarizing potentials and slowed deactivation. Consequently,

GV-58 was able to rescue some Ca2+ flux in response to an action potential-like

stimulus. My thesis suggests that therapeutic agents like GV-58 that increase channel

open probability may be effective in combatting this and other severe

neurodevelopmental disorders caused by loss-of-function mutations in CaV2.1.

v

ACKNOWLEDGEMENTS

I am forever grateful to my Thesis advisor, Roger Bannister, for his years of

support of my undergraduate and graduate research, as well as my non-research

related pursuits. Roger is an exceptional mentor whose understanding and patience

have allowed me to develop as a scientist. It was in his lab that I discovered my passion

for science and it was his advice and influence that have sparked my ambition to

become both a clinician and a bench scientist and pursue an MD-PhD. My life would

look quite different today had it not been for my time in the Bannister lab, and it saddens

me that I must leave.

Many thanks to Tyler Bendrick and Sara Beck-Pancer in the Bannister lab for

their company in the last year. Working with them never seemed like work. Additional

thanks to Tyler for her many hours spent transfecting and plating cells for me – this

thesis would not have been possible otherwise.

I also thank the Boettcher Foundation for both their monetary and systematic

support of this work. The Boettcher Scholarship has enabled me to pursue anything I

wish without the stress of financial obligation. I thank them for introducing me to Roger,

a development which, again, has altered the course of my life and career. The

Collaboration Grant and Educational Enrichment Grant programs have funded a

significant amount of the work in this thesis.

I would like to thank my parents for providing an environment in which I always

felt comfortable pursuing my dreams. Thank you for all your support in the past, and in

what’s to come.

vi

CONTENTS

CHAPTER

I. INTRODUCTION ........................................................................................... 1

Voltage-gated Ca2+ channels ................................................................... 1

Structure .................................................................................................. 3

Function ................................................................................................... 6

CaV Channelopathy ................................................................................. 7

II. CaV2.1 CHANNELOPATHIES ....................................................................... 9

Episodic Ataxia Type 2 ............................................................................ 10

Familial Hemiplegic Migraine Type 1 ....................................................... 11

CaV2.1 channelopathies affecting development ....................................... 12

III. CaV2.1 R1673P CAUSES A PROFOUND LOSS-OF-CHANNEL

FUNCTION .................................................................................................... 15

Introduction .............................................................................................. 15

Materials and Methods............................................................................. 17

Results ..................................................................................................... 23

Discussion ............................................................................................... 38

BIBLIOGRAPHY ........................................................................................................... 43

vii

FIGURES

I. INTRODUCTION

1-1. Ca2+ channel families .................................................................... 3

1-2. CryoEM structure and schematic of the rabbit CaV1.1 channel ..... 4

III. CaV2.1 R1673P CAUSES A PROFOUND LOSS-OF-CHANNEL-FUNCTION

3-1. Schematic diagram of V-CaV2.1 R1624P ...................................... 16

3-2. The R1624P mutation causes a profound depolarizing shift in Ca2+

current activation ........................................................................... 25

3-3. The R1624P mutation causes a profound depolarizing shift in Ba2+

current activation ........................................................................... 26

3-4. CaV2.1 R1624P deactivation kinetics ............................................ 27

3-5. Inactivation kinetics of CaV2.1 R1624P ......................................... 29

3-6. CaV2.1 and CaV2.1 R1624P are similarly inactivated following High

frequency stimulation .................................................................... 30

3-7. CaV2.1 and CaV2.1 R1624P demonstrate similar inactivation from

the closed state ............................................................................. 31

3-8. Roscovitine promotes Ca2+ flux via CaV2.1 R1624P at less

depolarizing test potentials ............................................................ 33

3-9. GV-58 promotes Ca2+ flux via both CaV2.1 and CaV2.1 R1624P .. 35

3-10. GV-58 increases Ca2+ flux via both CaV2.1 and CaV2.1 R1624P in

response to an action potential like waveform .............................. 37

1

INTRODUCTION

Voltage-gated Ca2+ channels

Ca2+ channels are found in every excitable cell (Hagiwara & Byerly, 1983). By

mediating Ca2+ flux in response to membrane depolarization, Ca2+ channels are able to

transduce electrical signals into chemical signals and regulate a litany of Ca2+-

dependent intracellular processes. Initial work on Ca2+ channels following the

development of patch-clamp methods revealed that two classes of channel could be

discriminated based on their response to changes in membrane potential. Broadly,

channels that opened at more positive potentials were deemed high-voltage activated

(HVA) Ca2+ channels and those that opened at more negative voltages were termed

low-voltage activated (LVA) Ca2+ channels (Carbone & Lux, 1984; Hagiwara, Ozawa, &

Sand, 1975; Llinás & Yarom, 1981).

The next stage of Ca2+ channel classification came from detailed

pharmacological and functional study. LVA current was found to inactivate quite rapidly

and would dissipate when a membrane was subjected to sustained depolarization.

These transient currents and tiny single channel conductances produced by the

formerly-named LVA channels spurred an additional naming of the current to “T-type”

(Nowycky, Fox, & Tsien, 1985). Contrastingly, HVA current lacked rapid inactivation,

and the long-lasting current and large single channel conductances produced by HVA

channels behooved an alternative naming of the channel to “L-type” (Tsien, Lipscombe,

Madison, Bley, & Fox, 1988). Further discrimination of T-type and L-type channels was

provided by the varying sensitivity of the channel subsets to a class of compounds

known as dihydropyridines (DHP), which contain molecules that can function as either

2

channel blockers (e.g., nifedipine) or channel agonists (e.g., BAY K 8644). L-type

channels were highly sensitive to these compounds, whereas T-type channels were not

(Bean, 1984).

Continued pharmacological study allowed for further exploration and refinement

of the channel classes. HVA channels were discovered that were resistant to

dihydropyridines and had single channel conductances larger than T-type and smaller

than L-Type. These neuronal channels were named N-type, and were further

distinguished from T- and L-type channels by highly selective blocking by -conotoxin

(Fox, Nowycky, & Tsien, 1987; Nowycky et al., 1985). Continued toxin work then

showed that the predominant Ca2+ channel in cerebellar Purkinje cells reacted

pharmacologically differently from L-type and N-type channels. These new, “P/Q-type”

channels were sensitive to block by the spider toxin -Aga-IVA (Llinás, Sugimori, Lin, &

Cherksey, 1989; Mintz, Adams, & Bean, 1992; Randall & Tsien, 1995). HVA channels

that were resistant to both dihydropyridines and -toxins were named R-type (Randall &

Tsien, 1995).

In the late 1980’s, molecular cloning became more feasible and Shoshaku

Numa’s group became the first to clone an ion channel (Noda et al., 1984). As the

cloning revolution progressed, a more precise channel nomenclature schema was

developed on the basis of molecular sequence . Ca2+ channels are now divided into 3

gene families based on their sequence similarity – CaV1, CaV2, and CaV3. The L-type

channels comprise the CaV1.X group, while the T-type channels make up the CaV3.X

subfamily. P/Q-, N-, and R- type channels are known as CaV2.1, CaV2.2, and CaV2.3,

respectively (Ertel et al., 2000; Figure 1-1).

3

Figure 1-1. Ca2+ channel families.

Structure

On the molecular level, voltage-gated Ca2+ channels (CaV) are heteromultimeric

complexes composed of a principal 1 subunit and auxiliary and 2 subunits

(Catterall, 2010; Figure 1-2). The 1 subunit of a given CaV is the principal identifier of

the channel complex (Figure 1-1), possessing four major transmembrane repeats (I-IV),

each containing six membrane-spanning helices (S1-S6, Figure 1-2 B). The S5 and S6

helices are the pore-lining units of the channel, while the S5 and S6 linker regions (P-

4

Figure 1-2. CryoEM structure and schematic of the rabbit CaV1.1 channel. (A)

CryoEM structure of rabbit CaV1.1 (taken from Wu et al., 2016). (B) Schematic of CaV1, 2,

and

loops) contain 4 highly conserved glutamate residues that function as a selectivity filter

for divalent cations such as Ca2+ and Ba2+ (Simms & Zamponi, 2014; Yang, Ellinor,

Sather, Zhang, & Tsien, 1993). In the absence of divalent ions, Ca2+ channels conduct

large amounts of monovalent charge. The >500 fold selectivity for Ca2+ over Na+ in

these channels is conferred by two Ca2+ binding sites in the pore that allow Ca2+ to bind

and repel monovalent charge. The electrostatic repulsion between the two bound Ca2+

allows for one of the ions to overcome the tight binding to the channel and proceed

through the pore before the other (Sather & McCleskey, 2003; Tang et al., 2013). The

S4 helices of the 1 subunit are the voltage-sensors of the channel because they

contain a positively-charged face that controls voltage-dependent activation (Aggarwal

& MacKinnon, 1996).

The 1 subunits of HVA (CaV1.X and CaV2.X channels) are always coexpressed

with and 2 subunits (Buraei & Yang, 2013; Dolphin, 2013; Hoppa, Lana, Margas,

5

Dolphin, & Ryan, 2012). Though T-type (LVA) channels do not seem to be affected

much by interactions with these additional channel subunits (Arias, Murbartián, Vitko,

Lee, & Perez-Reyes, 2005; Bae, Suh, & Lee, 2010; Dubel et al., 2004), the subunit is

critical for membrane expression and proper gating of HVA channels. is a cytoplasmic

protein that associates with the CaV1 subunit at the linker of Repeats I and II (Buraei &

Yang, 2013; Figure 1-2). The subunit is thought to form a complex with CaV1 and

coordinate a switch in the signaling of the complex from endoplasmic reticulum (ER)

retention to ER export (Fang & Colecraft, 2011). In this way, is critically responsible

for membrane trafficking and expression of CaV1. Additionally, enhances channel

open probability via a 10-15 mV hyperpolarizing shift in the voltage-dependence of

channel activation (Gregg et al., 1996; Murakami et al., 2002). There are four known

genes that encode subunits (Buraei & Yang, 2013), and each subunit is expressed in

different tissues with different effects on channel biophysics and physiology (Simms &

Zamponi, 2014).

The third subunit in a neuronal CaV complex is the 2 class. The location of 2

has been a topic of debate, with early groups describing the extracellular subunit

binding to a transmembrane anchor (the subunit) (De Jongh, Warner, & Catterall,

1990; Jay et al., 1991). Both the 2 and subunits are products of a single gene

encoding the 2 protein, which undergoes post-translational cleavage before

reassembling at the membrane. Additional work and the solution of the CaV1.1

cryostructure revealed that the entire 2 subunit is an entirely extracellular protein that

associates with the plasma membrane through a glycosylphosphatidylinositol (GPI)

6

anchor rather than a transmembrane domain (Davies et al., 2010; Kadurin et al., 2012;

Wu et al., 2016). 2 appears to have a small effect on channel function, but its

expression has been shown to augment channel density on the cell surface (Simms &

Zamponi, 2014; Yasuda et al., 2004). 2 has particularly important effects on neuronal

CaV, where it has been shown to increase both channel abundance and release

probability at neuronal synapses (Hoppa et al., 2012).

Function

Voltage-dependent activation of the various CaV isoforms is coordinated by the

voltage-sensing domain (S1-S4) of the 1 subunit. The S4 helix is the voltage-sensor of

the channel because it contains 5-6 positively charged amino acids (R0-R5) at three

amino acid intervals that translocate across the interior of the plasma membrane

(Simms & Zamponi, 2014). To facilitate this movement across the membrane field, the

S1-S3 helices have several negatively charged residues that stabilize the positive

charges of S4 (Catterall, 2010). S4 translocation in turn induces a conformational

change of the S5 and S6 helices which allows for opening of the channel pore and

conduction of divalent ions (Palovcak, Delemotte, Klein, & Carnevale, 2014).

Neutralization or disruption of the R0-R5 residues has been shown to have significant

impacts on gating of CaV (Bannister & Beam, 2013; Hans et al., 1999; Stühmer et al.,

1989; Tottene et al., 2002; Wappl et al., 2002).

Though Ca2+ channel activation has been extensively studied and plausible

mechanisms found, knowledge surrounding voltage-dependent inactivation of the same

7

channels is less robust (Herlitze, Hockerman, Scheuer, & Catterall, 2002; Stotz, Jarvis,

& Zamponi, 2004; Zhang, Ellinor, Aldrich, & Tslent, 1994). It is clear that channel

inactivation is an intrinsic regulatory mechanism that can be pathological in its

dysfunction (Lorenzon & Beam, 2008; Splawski et al., 2004, 2005). Inactivation is

responsible for moving the channel into a non-conducting gating state, preventing ion

flux across the membrane even when the channel is subject to supra-threshold stimuli.

This is contrasted with channel deactivation, which involves closing of the channel pore

following removal of a gating stimulus. Multiple mechanisms of channel inactivation

exist. “Fast” inactivation of Ca2+ channels occurs on the timescale of tens to hundreds of

milliseconds and is thought to involve the Repeat I-II linker region of the 1 subunit

functioning as a “hinged-lid” which occludes the channel pore in response to repetitive

or sustained depolarization (Stotz et al., 2004). Since the subunit binds the Repeat I-II

linker, different subunits can moderate inactivation differentially. “Slow” inactivation

occurring in response to depolarizations of seconds to minutes has been characterized

in Na+ channels but is less understood in Ca2+ channels. Theories include changes in

the voltage-sensor domain or collapse of the channel pore (Zhu, McDavid, & Currie,

2015). Like fast inactivation, the subunit has also been shown to moderate slow

inactivation (Sokolov, Weiss, Timin, & Hering, 2000). Ca2+ channels can also inactivate

from the closed state, though mechanisms for this type of inactivation are even less

clearly understood (Bahring & Covarrubias, 2011; Catterall, 2010; Patil, Brody, & Yue,

1998)

8

CaV Channelopathy

Mutations in genes encoding CaV complexes, particularly those responsible for

the principal 1 subunits, have been linked to a variety of human diseases (Lorenzon &

Beam, 2000). Disease causing mutations in 7 of the 10 1 subunits in humans have

been identified (Lorenzon & Beam, 2008; Striessnig, 2016). Problems in Ca2+ flux

resulting from CaV mutation can be very problematic in vivo since Ca2+ is critical in its

role as both a charge carrier and an intracellular signaling molecule (e.g., activation of

calmodulin, nuclear factor of activated T-cells, protein kinase C, etc.).

9

CaV2.1 CHANNELOPATHIES

P/Q-Type (CaV2.1) channels are located in presynaptic terminals throughout the

nervous system. Like other voltage-gated Ca2+ channels, CaV2.1 is composed of a

principal 1 subunit (1A) and auxiliary and 2 subunits (Catterall, 2010). In the 1A

subunit, there are 4 transmembrane repeats (I-IV) with six membrane-spanning helices

apiece (S1-S6). The S4 helices are the primary voltage-sensors of the channel, and this

voltage-sensing ability is conferred by a string of 5 positively charged amino acids

(Aggarwal & MacKinnon, 1996). The S1-S3 helices are necessary to stabilize the

positive charges of S4. S5 and S6 form the channel pore which eventually conducts

Ca2+ ions with high affinity (Neely & Hidalgo, 2014). Functional diversity of CaV2.1 is

generated by alternative splicing at multiple loci as well as 1A complexing with different

and 2 subunits. All members of the CaV2 subfamily have been shown to mediate

Ca2+ influx leading to neurotransmitter release. Of these, CaV2.1 is the most dominant

and most effective at releasing neurotransmitter in response to action-potential

stimulated membrane depolarization (Takahashi & Momiyama, 1993; Wheeler, Randall,

& Tsien, 1994; L. G. Wu, Westenbroek, Borst, Catterall, & Sakmann, 1999). Indeed,

density of functional CaV2.1 and preferential expression of CaV2.1 over other CaV2.X

isoforms is linked tightly to synaptic strength as examined at the Calyx-of-held synapse

(Lübbert et al., 2018).

Because CaV2.1 channels are the predominant Ca2+ channels present at the

neuromuscular junction and at most synapses of the central nervous system, Ca2+ flux

via CaV2.1 is critical for neurotransmitter release in these areas (Dunlap, Luebke, &

Turner, 1994, 1995; Ludwig, Flockerzi, & Hofmann, 1997). Mutations in CaV2.1 can

10

profoundly impact neuronal function; indeed, mutations in the 1A subunit of CaV2.1

have been linked to a multitude of neurological disorders in humans (Pietrobon, 2010).

Episodic Ataxia Type 2 (EA2) and Familial Hemiplegic Migraine Type 1 (FHM1) have

long been known to arise from point mutations in CaV2.1 (Jen et al., 2007; Pietrobon,

2007, 2010). The former disorder is generally caused by a loss-of-function mutation

while the latter is usually marked by channel gain-of-function (Jen et al., 2001; Tottene

et al., 2002). Specific amino acid substitutions in different regions of the channel can

affect channel functions in dramatically different ways. Some mutations can alter

channel gating, while others can cause haploinsufficiency and reduced channel

expression. Changes in channel function can affect Ca2+ flux at the neuromuscular

junction and central synapses.

Episodic Ataxia Type 2

EA2 is a rare neurological disease characterized by paroxysmal attacks of ataxia,

nystagmus, and vertigo. EA2 has been linked to loss-of-function mutations in CaV2.1 (J.

Jen et al., 2001; Sintas et al., 2017). The majority of EA2 causing disease mutations

disrupt the open reading frame, resulting in rapid degradation of truncated protein

products (Pietrobon, 2010). However, over 25 missense mutations have been identified,

most of which are substitutions of conserved amino acids that do not disrupt the reading

frame (Pietrobon, 2010; Sintas et al., 2017). The majority of these amino acid

substitutions are in the S5-S6 linker (P-loop) region or the S5 and S6 helices

themselves, suggesting that impaired ability to form a fully functional channel pore is the

likely pathophysiology of the EA2 phenotype in the majority of EA2 missense cases

11

(Jen et al., 2007; Sintas et al., 2017). In some cases, a complete loss-of-function was

observed, likely due to ER-associated degradation of the mutant channel and

subsequent lack of trafficking to the membrane (Page et al., 2004). Additionally, some

EA2 mutants seem to exert a dominant-negative effect since coexpression of mutant

channels with wild-type channels results in degradation of the wild-type channels and

expectedly diminished Ca2+ current under depolarization (Mezghrani et al., 2008). In

these cases, it is likely that misfolded mutant channels bind wild-type channels and

induce degradation (Page et al., 2010; Rajakulendran, Kaski, & Hanna, 2012).

Instances that have not completely abolished channel activity have also been observed.

In these mutations, the voltage-dependence of CaV2.1 activation shifts to significantly

more depolarizing potentials, decreasing channel open probability (Pietrobon, 2010).

Familial Hemiplegic Migraine Type 1

FHM1 is an inherited migraine condition that results in weakness of half the body

for prolonged periods of time. FHM1 is often accompanied by cerebellar degeneration

(Elliot, Peroutka, Welch, & May, 1996). In contrast to the etiology of EA2, FHM1 is most

often linked to gain-of-function mutations in CaV2.1 (Pietrobon, 2007; Tottene et al.,

2002). Thus far, over 25 missense mutations have been identified in cases of FHM1. All

of these mutations are substitutions, most commonly in the line pore, the S3-S4 or S5-

S6 linkers, or the S4 voltage-sensor (Pietrobon, 2010). Though the locations of the

mutations are variable, expression and analysis in heterologous systems has revealed a

persistent hyperpolarizing shift in channel activation for all studied mutants (Serra,

Fernàndez-Castillo, & Fernández-Fernández, 2009). Because these channels open at

12

lower voltages, channel open probability is greatly enhanced and an FHM1 CaV2.1

channel can support much greater Ca2+ influx than its wild-type counterpart. Mouse

knock-in models of FHM1-causing CaV2.1 channel mutations favor the initiation and

propagation of the electrophysiological marker of migraine aura, cortical spreading

depression (van den Maagdenberg et al., 2004, 2010).

CaV2.1 Channelopathies affecting Development

EA2 and FHM1 have long been known to be caused by point mutations in CaV2.1

(Jen et al., 2001; Pietrobon, 2007, 2010). However, biophysical effects on the channel

from these mutations are generally small and effects on phenotype are often

paroxysmal (Elliot et al., 1996; Jen et al., 2007; Sintas et al., 2017). With the innovative

whole-exome sequencing approach, a new class of more severe CaV2.1 linked

disorders with developmental components have been identified and linked to point

mutations in CaV2.1 (Blumkin et al., 2010; Luo et al., 2017; Romaniello et al., 2010;

Travaglini et al., 2017; Weyhrauch et al., 2016). These disorders share some

characteristics with EA2 but are distinct. These disorders are rare and thus are not

well-studied.

Romaniello et al. have described an A405T mutation in a 12-year-old girl with a

family history of CaV2.1 mutation-linked disorders (Romaniello et al., 2010). The patient

presented with persistent cerebellar signs (ataxia, dysmetria, hypotonia) and

developmental delay. A405T is a non-polar to polar substitution in the Repeat I-II linker

region of CaV2.1. The Repeat I-II linker is the site where the subunit binds the 1A

13

complex. A mutation like A405T could potentially disrupt subunit binding to 1A. This

disruption would substantially decrease surface expression of the channel by increasing

ER retention.

Blumkin et al. reported a R1350Q mutation in a 7-year-old male patient that

presented with cerebellar ataxia, developmental delay, and nonspecific dyskinesia

(Blumkin et al., 2010). This substitution changes a charged amino acid to a neutral one

in the S4 helix of CaV2.1 (the voltage-sensor). The loss of this charge may prevent

normal channel activation by disrupting the movement of the voltage-sensor through the

membrane field.

Travaglini et al. reported a pair of mutations (I1342T and V1396M) in two patients

with similar clinical phenotypes (Travaglini et al., 2017). Both cases involved congenital

ataxia, hypotonia, and intellectual disability. The I1342T mutation is a hydrophobic to

polar residue substitution in the intracellular loop between the S3 and S4 helix of 1A

(close to the first charged residue of the S4 voltage-sensor). It is hypothesized that this

substitution likely alters the conformation or mobility of the S4 segment, disrupting

normal voltage-sensing function of CaV2.1. The V1396M mutation is found in the S5

pore-forming domain of 1A. Valine-Methionine replacement could modify the helix

packing in the pore-forming domain by modifying hydrophobic interactions. Alteration of

the channel pore could result in decreased Ca2+ flux through CaV2.1.

Weyhrauch et al. described a child with developmental delay, gross motor delay,

and congenital hypotonia linked to a P1353L mutation in CaV2.1 (Weyhrauch et al.,

2016). P1353L occurs in the S4 voltage-sensor of Repeat 3 of 1A. Electrophysiological

14

analysis of the mutation revealed near 100% ablation of CaV2.1-mediated Ca2+ current,

likely due to reduced expression at the cell surface. Loss-of-channel-function in this

case could be due to either dominant-negative effects or haploinsufficiency.

Study of CaV2.1 channelopathy is further complicated by the fact that FHM1 and

EA2 can have overlapping clinical presentations. For example, ~20% of FHM1 cases

have ataxic components (Elliot et al., 1996). Pharmacological intervention is an obvious

goal of channelopathy research. To this end, functional consequences of disease-

causing mutations in CaV2.1 are likely more relevant than the broad clinical phenotype

they elicit. Since both CaV2.1 agonists and antagonists exist, it is critical to identify if a

mutation is causing a gain-of-channel-function or a loss-of-channel-function as well as

the mechanism of how function is altered.

15

THE R1673P MUTATION CAUSES A PROFOUND LOSS-OF-CHANNEL-FUNCTION

Introduction

Recently, Luo and colleagues described an 8-year-old (now 11) female patient

with congenital ataxia, hypotonia, cerebellar atrophy, and global developmental delay.

Trio-based exome sequencing of this patient revealed a de novo missense mutation

(R1673P) in the gene for CaV2.1 being expressed heterozygously. The mutation results

in an arginine to proline substitution within the RIV S4 voltage-sensing helix of CaV2.1.

This mutation was predicted to be “probably damaging” by PolyPhen-2, a protein

prediction software. In an attempt to elucidate the molecular mechanism by which

R1673P resulted in the clinical phenotype, the group created transgenic, fly-based

constructs of both wild-type CaV2.1 and mutant CaV2.1 R1673P. Next, they expressed

these transgenes in CaV2.1 null drosophila larvae. The authors found that the mutant

CaV2.1 was able to rescue the photoreceptor response in these larvae but promoted

neurodegeneration. Because amplitudes of photoreceptor transients in larvae rescued

with the mutant channel were larger than those rescued with the wild-type channel, the

group hypothesized that increased Ca2+ influx via a gain-of-function mutation in CaV2.1

led to excitotoxicity and eventual neurodegeneration. On the basis of this gain-of-

function finding, the patient has been prescribed Ca2+ channel blockers (Luo et al.,

2017).

However, the nature of the drastic amino acid change and location of the

mutation warranted further study. Indeed, the mutation occurs in a location of the

channel that is critical for voltage-sensing function; additionally, proline can be a

16

particularly problematic substitute because of its tendency to induce tight turns in its

parent protein.

To study the molecular mechanisms by which the R1673P mutation impacts

CaV2.1 channel function, we expressed the rat version of the mutant channel (R1624P;

Fig. 3-1) in a heterologous system (tsA-201 cells). Then, using the whole-cell voltage-

clamp technique (Hamill et al., 1981), we investigated various properties of the mutant

channel and explored pharmacological compounds that could potentially correct defects

induced by the R1673P mutation. In this thesis, I describe that, in contrast to the

findings of Luo et al., the R1673P mutation causes a profound loss-of-channel-function

via a depolarizing shift in the voltage-dependence of channel activation and slowed

channel activation kinetics.

Figure 3-1. Schematic diagram of V-CaV2.1 R1624P.

17

Materials and methods

Molecular biology

The venus-CaV2.1 plasmid was a generous gift from Dr. P. J. Kammermeier

(University of Rochester). To derive the Venus-fused rat CaV2.1 R1624P (human

numbering: R1673P) construct, we induced a single G to C point mutation at bp 4871 of

V-CaV2.1 using the Q5 Site-Directed Mutagenesis Kit. The forward primer was 5’-

AAACTCCTCCcCCAGGGTTACAC-3’ and the reverse primer was 5’-

GATGAGTCGGGCAGCACG-3’. To verify that there were no off-target mutations in the

other voltage-sensing regions of the channel, we sequenced the four conserved repeats

of the mutant CaV2.1 construct.

Cell culture and transfection

tsA-201 cells were cultivated in culture medium containing 90% DMEM, 10%

fetal bovine serum, and 100 g/ml Penicillin-Streptomycin. Cells of low passage number

(<15) were plated on 35 mm culture wells. Lipofectamine 2000 was used to transfect

these cells with a mixture containing expression plasmids encoding rat V-CaV2.1 or V-

CaV2.1 R1624P, rabbit 2-1, and rat 4 subunits for a total of 1 g of each cDNA per

well. 24 hours following transfection, we trypsinized cells and replated them onto 35 mm

plastic culture dishes. We used successfully transfected cells (i.e., those exhibiting

Venus fluorescence) in experiments 24 hours after the replating step.

18

Confocal imaging

Images of live tsa-201 cells expressing either wild-type V-CaV2.1 or V-CaV2.1-

R1624P were acquired using a Zeiss LSM780 confocal laser scanning microscope.

Venus was excited with the 488 nm line of an Argon laser directed to the cell via a

488/543 nm dual dichroic mirror. Emitted Venus fluorescence was directed to a

photomultiplier equipped with a 500-550 nm band pass filter.

Solutions

We used solutions with the following compositions (concentrations in mM):

1. Internal solution: 140 Cs-aspartate, 10 Cs2-EGTA, 5 MgCl2, 10 HEPES, pH 7.4

with CsOH.

2. Ca2+ external solution: 145 Tetraethylammonium-Cl, 4 KCl, 2 CaCl2, 10 HEPES,

10 glucose, 1 4-aminopyridine, pH 7.4 with Tetraethylammonium-OH

3. Ba2+ external solution: 145 Tetraethylammonium-Cl, 4 KCl, 2 BaCl2, 10 HEPES,

10 glucose, 1 4-aminopyridine, pH 7.4 with Tetraethylammonium-OH

4. Roscovitine perfusion solution: 50 mM in 100% DMSO, diluted in the Ca2+

external solution to 12.5 M prior to experiments.

5. GV-58 perfusion solution: 50 mM in 100% DMSO, diluted in the Ca2+ external

solution to 12.5 M prior to experiments.

19

Whole cell electrophysiology

All experiments were performed at room temperature. Fabricated borosilicate

pipettes with resistances ranging from 3.0-4.5 M were filled with internal solution.

Electronic compensation was used to reduce the effective series resistance. Linear

components of leak and capacitive currents were reduced with P/4 leak subtraction.

Except where otherwise noted, filtering was done at 5 kHz and digitization performed at

10 kHz. Cell capacitance (Cm) was measured by integration of a transient from -80 mV

to -70 mV using Clampex 9.2. The time constant for decay of the whole-cell capacitance

transient (m) was reduced using the analog compensation circuit of the amplifier. Cm

was used to normalize current amplitudes to approximate cell size (pA/pF). The average

value of Cm was 30 ± 1 pF (n = 174 cells) and the average values of m and Ra were 279

± 9 s and 9.5 ± 0.3 M, respectively.

To analyze the voltage-dependence of activation of the wild-type and mutant

channels, we applied a series of test potentials to cells in the whole-cell configuration

and measured the resulting current using an Ag/AgCl patch electrode. Cells were

clamped at a holding potential of -90 mV, and 25 ms depolarizations ranging from -50 to

+90 mV were applied in 10 mV increments. Cells were repolarized to -40 mV before

returning to the holding potential. Current-voltage (I-V) curves were fitted according to:

I = Gmax(V-Vrev)/(1+exp[(V1/2-V)/kG]) (Equation 1)

Where I is the normalized current (pA/pF) corresponding to test potential V, Vrev is the

reversal potential, Gmax is the maximal channel conductance, V1/2 is the half-maximal

activation potential, and kG is the slope factor.

20

The tail currents measured during these recordings were used to construct

conductance-voltage (G-V) relationships for the channels. Tail current amplitudes were

measured immediately after the onset of repolarization from a given test pulse to -40

mV. These amplitudes were normalized to the tail current amplitude produced by

repolarization from +90 mV to -40 mV. Normalized tail current values were fit with the

equation:

G/Gmax = 1/(1+exp[-(V1/2act-V)/k]), (Equation 2)

Where G is the tail current amplitude evoked by repolarization from a given test

potential V back to -40 mV, Gmax is the conductance for repolarization from +90 mV to -

40 mV, V1/2act is the half-maximal activation potential and k is the slope factor.

The above steps for construction of I-V and G-V relationships were repeated in

Ba2+ external solution. Additionally, to measure the activation kinetics of wild-type and

mutant channels in Ba2+, we applied single-exponential fits to the recordings using the

function:

I(t) = I0[exp(-t/act)], (Equation 3)

Where I(t) is the current at a given time t, I0 is the initial current, and act is the time

constant for activation.

To elucidate differences in voltage-dependent deactivation, we evaluated

deactivation kinetics following 25 ms depolarizations to voltages corresponding to peak

activation of either the wild-type (+10 mV) or the mutant (+30 mV) channels. Following

this maximal activation, cells were repolarized to a range of voltages in 10 mV

increments (-90 mV to +30 mV) for 60 ms. Only tail currents which successfully

21

repolarized to baseline and were not complicated by repolarization potentials with

probable channel opening were fit with the following equation:

Itail(t) = Itail0 [exp(-t/deact)], (Equation 4)

Where Itail(t) is the amplitude of the tail current at a time t, Itail0 is the maximal tail current

amplitude, and deact is the time constant for deactivation.

To assess differences in voltage-dependent current decay, we measured current

decay kinetics after 500 ms test pulses to voltages corresponding to maximal activation

of wild-type and mutant channels. Both the holding and repolarization potentials were

-90 mV. Half-times of total current decay were measured (t1/2decay).

To measure how the channels reacted to a high-frequency stimulus, we

measured Ba2+ currents from cells exposed to a 25 ms depolarization before and after a

200 ms, 100 Hz conditioning train. Both the test and the train potentials corresponded to

the peak of the Ba2+ I-V relationships for the wild-type and mutant channels.

To measure closed-state inactivation, we elicited Ba2+ currents using 100 ms

depolarizations to +40 mV following 5 s prepulses ranging from -100 to +20 mV and a

35 ms repolarization to -90 mV. Current was normalized to the current produced by the

depolarization following the -100 mV prepulse. Normalized steady-state inactivation

curves were fit by the equation:

I/Imax = 1/(1+exp[-(V1/2inact – V)/k]) (Equation 4)

Where I is the Ba2+ current corresponding to a depolarization to +40 mV following a

prepulse to potential V, Imax is the current produced in response to depolarization to +40

22

mV following the -100 mV prepulse, V1/2inact is the half-maximal inactivation potential,

and k is the slope factor.

Ramp waveforms were used to mimic neuronal action potentials. Cells were

depolarized by a 1 ms ramp from -80 mV to +30 mV and then immediately repolarized

from +30 mV to -80 mV by another 1 ms ramp. Currents were acquired at 33 kHz and

filtered at 10 kHz. Peak currents were measured during the action potential like

waveforms and expressed in pA/pF. Charge flux was calculated as the integral of the

evoked current bounded by the baseline during the ramp protocol.

Roscovitine and GV-58 (both Alomone) perfusion solutions were applied through

a manually operated, gravity-driven global perfusion system. I-V and G-V relationships

before and after application of the drugs were constructed and fit with equations 1 and 2

as discussed previously. Action potential like waveforms were used on cells before and

after drug application as well and quantified as mentioned earlier.

Statistics

All data are presented as mean ± SEM. All statistical comparisons were by

unpaired, two-tailed t-test, with the exception of drug perfusion data, which were

compared by paired, two-tailed t-test. All figures were made using SigmaPlot software

(version 14.0). p<0.05 was considered statistically significant.

23

Results

The R1624P mutation causes a profound depolarizing shift in CaV2.1 activation

We used the whole-cell voltage-clamp technique to test whether the CaV2.1

R1624P mutation altered channel gating. Indeed, with 2 mM Ca2+ serving as the charge

carrier, tsA-201 cells expressing V-CaV2.1 produced Ca2+ currents peaking around +10

mV (Fig. 3-2 A, C). Cells expressing the mutant V-CaV2.1 R1624P channel also

supported Ca2+ current, but only at more depolarizing test potentials, peaking near +35

mV (Fig. 3-2 B, C). This substantial depolarizing shift in channel activation was clear in

the I-V relationships (V1/2 = 18.2 ± 1.6 mV, n = 23 vs. -0.89 ± 1.02 mV, n = 24,

respectively; p < 0.0001; Fig. 3-2 C). Gmax values in the I-V fits of V-CaV2.1 and V-

CaV2.1 R1624P were not significantly different (690 ± 30 and 560 ± 70 pS/pF,

respectively; p > 0.05), indicating that differences in current density were results of the

shift in voltage-dependence of activation and not due to variable channel expression.

An important caveat to these data is that the V1/2 values obtained from fitting the

I-V relationships of both the wild-type and mutant channel were complicated by an

outward Cs+ conductance at more depolarizing test potentials. The equation of the I-V fit

requires the reversal potential of the channel, but the potential refers to a single ion

only. Since there is some Cs+ efflux at more depolarizing potentials, the experimental

reversal potential instead reflects the voltage at which Ca2+ influx equals Cs+ efflux and

is dependent on multiple ions. This results in some error in the values obtained from the

I-V fit. To more clearly elucidate the nature of the shift in activation, we analyzed tail

currents from the same voltage-clamp recordings and constructed G-V relationships for

both V-CaV2.1 and V-CaV2.1 R1624P. The profound depolarizing shift in channel

activation of V-CaV2.1 and V-CaV2.1 R1624P persisted in these G-V relationships (V1/2

24

= 4.3 ± 1.2 mV vs. 27.8 ± 2.1 mV, respectively; p < 0.001; Figure 3-2 D). The same

depolarizing shift was retained when 2 mM Ba2+ was used as the charge carrier (V1/2

20.8 ± 2.7 and -8.93 ± 2.3 mV, respectively; p < 0.001; Fig. 3-3 A-C).

The R1624P mutation slows the activation kinetics of CaV2.1

Current traces from wild-type and mutant channels appeared to have different

channel activation kinetics. To assess this possibility, we fit Ba2+ current activation over

a range of test potentials (+10 mV to +30 mV) with a single exponential function.

Indeed, activation kinetics were significantly slower for V-CaV2.1 R1624P than wild-type

V-CaV2.1 at all test potentials examined (all sets p < 0.05, Fig. 3-3 D).

The R1624P mutation minimally affects the voltage-dependence of CaV2.1 deactivation

Most of the Ca2+ influx at the neuromuscular junction occurs during the falling phase of

the action potential (Llinás & Yarom, 1981). Therefore, changes in CaV2.1 deactivation

could more profoundly impact action potential induced Ca2+ influx than changes in

channel activation alone. For this reason, we investigated deactivation kinetics evoked

by repolarization from peak activation (+10 mV for V-CaV2.1 and +30 mV for V-CaV2.1

R1624P) to a range of less depolarizing test potentials. In these experiments, we

demonstrated that the wild-type V-CaV2.1 channel deactivates at voltages ~20 mV more

hyperpolarized relative to V-CaV2.1 R1624P (Fig. 3-4). However, considering the 20 mV

difference in the preceding depolarization, it appears that there is not a significant

difference in the voltage-dependence of channel deactivation in that a similar magnitude

25

Figure 3-2. The R1624P mutation causes a profound depolarizing shift in Ca2+ current

activation. Ca2+ current families recorded from tsA-201 cells expressing V-CaV2.1 (A) or V-

CaV2.1 R1624P (B) with 4 and 2-1. Currents were elicited by 25 ms step depolarizations

from -90 mV to the indicated test potentials; the repolarization voltage was -40 mV. (C)

comparison of V-CaV2.1 (●; n = 24) and V-CaV2.1 R1624P (○; n = 23) average peak I-V

relationships. Currents were evoked at 0.1 Hz by test potentials ranging from -50 mV through

+90 mV in 10 mV increments. Amplitudes were normalized by capacitance (pA/pF). I-V curves

are plotted according to Equation 1 with the following respective parameters for V-CaV2.1 and

V-CaV2.1 R1624P: Gmax = 690 ± 30 and 560 ± 70 pS/pF, V1/2 = -0.9 ± 1.0 and 18.2 ± 1.6 mV,

Vrev = 71.1 ± 2.6 and 77.7 ± 2.0 mV and k = 3.4 ± 0.8 and 6.2 ± 0.9 mV. (D) normalized G-V

relationships were fit with Equation 2 with the following respective parameters for V-CaV2.1 and

V-CaV2.1 R1624P: V1/2 = 2.3 ± 1.20 and 27.8 ± 2.1 mV; k = 4.3 ± 0.8 and 8.2 ± 1.2 mV.

Throughout, error bars represent ± SEM.

26

Figure 3-3. The R1624P mutation causes a profound depolarizing shift in Ba2+ current

activation. Ba2+ current families recorded from tsA-201 cells expressing V-CaV2.1 (A) or V-

CaV2.1 R1624P (B) with 4 and 2-1. Currents were elicited by 25 ms step depolarizations

from -90 mV to the indicated potentials; the repolarization voltage was -40 mV. (C) normalized

G-V relationships were fit with Equation 2 with the following respective parameters for V-CaV2.1

(●; n = 16) and V-CaV2.1 R1624P (○; n = 10): V1/2 = -8.9 ± 2.3 and 20.8 ± 2.7 mV; k = 6.9 ± 0.7

and 9.7 ± 0.4 mV. (D) comparison of activation kinetics from tsA-201 cells expressing either V-

CaV2.1 (●; n = 16) or V-CaV2.1 R1624P (○; n = 10). For both channels, the activation phase

was fit by Equation 3. Only test potentials in which a substantial amount of current was present

are shown (i.e., ranging from 0 mV through +30 mV for V-CaV2.1 and +10 mV through +30 mV

for V-CaV2.1 R1624P). Significant differences by two-tailed, unpaired t-test are indicated (*, p <

0.05).

27

Figure 3-4. CaV2.1 R1624P deactivation kinetics. Ba2+ tail currents were elicited by 60 ms

repolarizations to test potentials ranging from -90 mV to +10 mV following a 25 ms

depolarization from -90 mV to either +30 mV or +10 mV (A). Representative Ba2+ tail currents

recorded from tsA-201 cells expressing V-CaV2.1 (B) or V-CaV2.1 R1624P (C) following

depolarization from either +10 mV for V-CaV2.1 or +30 mV for V-CaV2.1 R1624P to the indicated

test potentials. (D) comparison of deactivation kinetics from tsA-201 cells expressing either V-

CaV2.1 (●; n = 10) or V-CaV2.1 R1624P (○; n = 7). For both channels, deactivation was fit by

Equation 4. Significant differences by two-tailed, unpaired t-test are indicated (*, p < 0.05).

28

of repolarization from peak activation is required for both wild-type and mutant channel

closure. The shift in activation of the mutant channel relative to wild-type makes

assessing deactivation at the same depolarization voltage difficult.

Inactivation kinetics of CaV2.1 are accelerated by the R1624P mutation

Alterations in Ca2+ channel inactivation kinetics have been identified as the basis

for multiple devastating disorders (e.g., Timothy syndrome, Splawski et al., 2004, 2005).

To investigate the effects of the R1624P mutation on CaV2.1 inactivation, we measured

Ba2+ current decay kinetics during 500 ms depolarizations to the voltage of peak

channel activation (+10 mV for wild-type V-CaV2.1 and +30 mV for V-CaV2.1 R1624P,

Figure 3-4). In these experiments, we observed that the V-CaV2.1 R1624P channel

inactivates significantly faster than the wild-type V-CaV2.1 channel (t1/2decay = 110 ± 14

ms, n = 13, vs. 270 ± 39 ms, n = 10, respectively; p < 0.001; Fig. 3-5 C). Though the

R1624P mutation destabilizes the open state, the acceleration of inactivation occurs at

timescales that are unlikely to be relevant in vivo.

CaV2.1 and CaV2.1 R1624P demonstrate similar levels of inactivation following a high-

frequency stimulus protocol and in the closed state.

To assess how the channel might respond to more physiological inactivation

conditions, tsA-201 cells transfected with either the wild-type or the R1624P channel

were depolarized following a 200 ms, 100 Hz stimulus train (Fig. 3-6). Indeed, there was

no appreciable difference in the amount of decay of the peak current between V-CaV2.1

and V-CaV2.1 R1624P (P2/P1 = 0.958 ± 0.007, n = 10 vs. 0.946 ± 0.006, n = 10,

29

Figure 3-5. Inactivation kinetics of CaV2.1 R1624P. Representative Ba2+ currents recorded from tsA-201 cells expressing V-CaV2.1 (A) or V-CaV2.1 R1624P (B) near the peak of the I-V relationship for either channel. Currents were elicited by 500 ms depolarizations from -90 mV to +10 mV (A) and +30 mV (B). In both (A) and (B), half-times of decay are indicated by the red dot. (C) summary of half-times of current decay for V-CaV2.1 (●; n = 10) and V-CaV2.1 R1624P (○; n = 13). Means and medians are indicated by the dashed and solid black lines of the boxes, respectively. Boxes represent the 25th/75th percentiles. Bars represent the 5th/95th percentiles. A significant difference by two-tailed, unpaired t-test is indicated (p < 0.001).

30

Figure 3-6. CaV2.1 and CaV2.1 R1624P are similarly inactivated following High frequency

stimulation. Ba2+ currents from tsA-201 cells expressing V-CaV2.1 (A) or V-CaV2.1 R1624P

(B) with 4 and 2-1 were elicited by 25 ms depolarizations before (left panels) and following a

200 ms, 100 Hz conditioning train (right panels). The test and train potentials (2 ms) correspond

to the peak of the Ba2+ I-V relationships for V-CaV2.1 (+10 mV) and V-CaV2.1 R1624P (+30

mV). The membrane potential was returned to -90 mV for 10 ms following the conditioning

train. The protocols illustrated at the top are not drawn to scale. Filtering was at 2 kHz and

digitization was at 5 kHz. (C) summary of results with number of experiments indicated. Means

and medians are indicated by the dashed and solid lines of the boxes, respectively. Boxes

represent the 25th/75th percentiles. Bars represent the 5th/95th percentiles. No significant

differences were observed.

31

Figure 3-7. CaV2.1 and CaV2.1 R1624P demonstrate similar inactivation from the closed state. (A) Ba2+ currents were elicited by 100 ms depolarizations to +40 mV following 5 s prepulses ranging from -100 mV to +20 mV and a 35 ms repolarization to -90 mV. The voltage protocol is not drawn to scale. Representative Ba2+ currents recorded from tsA-201 cells expressing V-CaV2.1 (B) or V-CaV2.1 R1624P (C) using the voltage protocol illustrated in panel (A). Filtering was at 2 kHz and digitization was at 5 kHz. (D) voltage-dependence of inactivation for V-CaV2.1 (●; n = 13) and V-CaV2.1 R1624P (○; n = 8). Normalized steady-state inactivation curves were fit by the equation I/Imax = 1/(1 + exp[-(V1/2inact - V)/k]), with the following respective parameters for V-CaV2.1 and V-CaV2.1 R1624P: V1/2inact = -33.3 ± 2.2 mV and -29.1 ± 3.4 mV; k = -8.1 ± 1.1 mV and -9.6 ± 1.2 mV. No significant differences were observed.

32

respectively; p > 0.05; Fig. 3-6 C). Additionally, there was no significant difference in

inactivation in the closed state, assessed by depolarizations to the peak of mutant

channel activation following a 5 second conditioning prepulse (V1/2inact = -33.3 ± 2.2 mV,

n = 13 vs. -29.1 ± 3.4 mV, n = 8, respectively; p > 0.05; Fig. 3-7).

Roscovitine promotes Ca2+ flux via CaV2.1 R1624P at less depolarizing test potentials

Roscovitine is a cyclin-dependent kinase inhibitor that has demonstrated CaV2.1

agonist ability (Yan, Chi, Bibb, Ryan, & Greengard, 2002). On the basis of Roscovitine’s

ability to promote Ca2+ flux in the wild-type channel, we tested the idea that it could

rescue some Ca2+ current in V-CaV2.1 R1624P as well. Indeed, the drug significantly

slowed the rate of tail current decay upon repolarization from +10 mV back to -40 mV

(deact = 0.62 ± 0.05 ms vs. 1.07 ± 0.14, before and during Roscovitine application,

respectively; p < 0.01, Fig. 3-8 A, D) Additionally, the drug also increased Ca2+ flux via

V-CaV2.1 R1624P at less depolarizing potentials corresponding to peak activation of

wild-type V-CaV2.1 (Fig. 3-8 A-B). This increase in current amplitude at +10 mV was a

consequence of a >10 mV hyperpolarizing shift in the voltage-dependence of CaV2.1

R1624P activation. (V1/2 = 19.6 ± 0.9 mV vs. 5.3 ± 2.6 mV, before and during

Roscovitine application, respectively; p < 0.001, Fig. 3-8 C).

33

Figure 3-8. Roscovitine promotes Ca2+ flux via CaV2.1 R1624P at less depolarizing

test potentials. (A) Ca2+ currents recorded before (1) and during (2) application of 12.5 M

Roscovitine to a tsA-201 cell expressing V-CaV2.1 R1624P, 4, and 2-1 (holding potential = -

90 mV; test potential = +10 mV with repolarization to -40 mV). Roscovitine (Alomone) was

dissolved in 100% DMSO to make 10 mM stock solution and then diluted in the external

recording solution just prior to experiments. During experiments, the Roscovitine working

solutions was applied through a manually-operated, gravity-driven global perfusion system. (B)

time-course of step current amplitude before (○) and during (●) Roscovitine application.

Numbers correspond to traces shown in panel (A). Currents were evoked at 0.1 Hz. Filtering

was at 2 kHz and digitization was at 5 kHz. (C) normalized G-V relationships before and during

application of Roscovitine (n = 9). Currents were evoked at 0.1 Hz by test potentials ranging

from -50 mV through +90 mV in 10 mV increments. G-V curves are plotted according to

Equation 2 with the respective parameters for control and Roscovitine: V1/2 = 30.3 ± 1.9 and

16.8 ± 1.9 mV and k = 7.7 ± 0.6 and 12.5 ± 0.8 mV. (D) comparison of deact of tail currents

evoked by repolarization from +10 mV to -40 mV in the absence (○) or presence (●) of

Roscovitine. Tail currents were fit by Equation 4. Means and medians are indicated by the

dashed and solid lines of the boxes, respectively. Boxes represent the 25th/75th percentiles.

Bars represent the 5th/95th percentiles. A significant difference by two-tailed, paired t-test is

indicated (p < 0.01).

34

The Roscovitine derivative GV-58 promotes Ca2+ flux via both CaV2.1 and CaV2.1

R1624P

Though Roscovitine’s effects were encouraging, its therapeutic potential is

dampened somewhat by its wide-ranging effects on ion channels other than CaV2.1

(Buraei, Schofield, & Elmslie, 2007; Yarotskyy & Elmslie, 2007). For this reason, we

decided to test its higher-affinity derivative, GV-58, to see if the agonist effects of its

parent compound persisted in a more promising clinical candidate. To parse out if GV-

58 targeted the wild-type and mutant channel differentially, both cells transfected with V-

CaV2.1 and V-CaV2.1 R1624P were perfused with the drug and tested. When applied to

the wild-type channel, GV-58 expectedly slowed deactivation (deact = 0.69 ± 0.07 ms vs.

5.79 ± 0.48 ms, before and during GV-58 application, respectively; p < 0.001; Fig. 3-9 A,

C). Step current was also slightly augmented due to a small hyperpolarizing shift in the

voltage-dependence of V-CaV2.1 activation (V1/2 = 1.7 ± 1.3 mV vs. -5.7 ± 2.8 mV,

before and during drug application, respectively; p < 0.001, Fig. 3-9 B, D). We observed

similar effects on deactivation when GV-58 was applied to cells expressing V-CaV2.1

R1624P (deact = 1.02 ± 0.03 ms vs. 4.41 ± 0.45 ms, before and during GV-58

application, respectively; p < 0.001, Fig. 3-9 E, G). Additionally, the effects of the drug

on step current were significantly more prominent in the mutant experiments. In

particular, the hyperpolarizing shift in activation of the mutant channel elicited by GV-58

was ~3 times larger than shift in the wild-type V1/2 = 27.8 ± 2.2 mV vs. 10.8 ± 1.9 mV,

before and during drug application, respectively; p < 0.001, Fig. 3-9 F, H)

35

Figure 3-9. GV-58 promotes Ca2+ flux via both CaV2.1 and CaV2.1 R1624P. (A) Ca2+

currents recorded before (1) and during (2) application of 12.5 M GV-58 to a tsA-201 cell

expressing V-CaV2.1, 4 and 2-1. (B) time-course of step current amplitude before (●) and

during (●) GV-58 application. Currents were evoked by the protocol illustrated in (A) at 0.1 Hz.

Numbers correspond to traces shown in panel (A). (C) comparison of deact of V-CaV2.1 tail

currents evoked by repolarization from +10 mV to -40 mV in the absence (●) or presence (●) of

GV-58 (n = 8). (D) normalized G-V relationships before and during application of GV-58.

Currents were evoked at 0.1 Hz by test potentials ranging from -50 mV through +90 mV in 10

mV increments. G-V curves are plotted according to Equation 2 with the respective parameters

for control and GV-58: V1/2 = 1.7 ± 1.3 and -5.7 ± 2.8 mV and k = 4.8 ± 0.4 and 5.6 ± 0.8 mV.

(E) Ca2+ currents recorded before (1) and during (2) application of 12.5 M GV-58 to a tsA-201

cell expressing V-CaV2.1 R1624P, 4 and 2-1. The corresponding time course is shown (F).

(G) comparison of V-CaV2.1 R1624P deactivation upon repolarization from +10 mV to -40 mV in

the absence (○) or presence (●) of GV-58 (n = 7). (H) normalized G-V relationships for V-

CaV2.1 R1624P before and during application of GV-58. The respective G-V fit parameters for

control and GV-58 were: V1/2 = 27.9 ± 2.2 and 10.8 ± 1.9 mV and k = 8.3 ± 0.4 and 8.6 ± 0.5

mV. For reference, the G-V curve for wild-type V-CaV2.1 in the absence of GV-58 is shown as a

dashed black line. As in Figure 3, tail currents were fit by Equation 4. Means and medians in

panels (C) and (G) are indicated by the dashed and solid lines of the boxes, respectively.

Boxes represent the 25th/75th percentiles. Bars represent the 5th/95th percentiles. Significant

differences by two-tailed, paired t-test are indicated.

36

GV-58 augments Ca2+ flux via both CaV2.1 and CaV2.1 R1624P in response to an

action potential like waveform

The depolarizing shift combined with the prolonged activation kinetics of V-

CaV2.1 R1624P relative to V-CaV2.1 support the idea that the mutant channel will only

be minimally activated by a neuronal action potential. To further probe this idea, we

evoked current using an action potential like ramp waveform (Bahamonde et al., 2015).

Briefly, the protocol consisted of a 1 ms ramp from -80 mV to +30 mV followed by a 1

ms ramp back to -80 mV (Fig. 3-10 A, D). This method enabled significant Ca2+ influx

via wild-type V-CaV2.1 as measured by both amplitude (-4.4 ± 0.7 pA/pF; Fig. 3-10 B)

and total flux (-2.7 ± 0.3 nC/F; Fig. 3-10 C). The same protocol applied to cells

expressing mutant V-CaV2.1 R1624P resulted in virtually no current amplitude (-1.4 ±

0.2 pA/pF, n = 7; p < 0.001 vs. that for V-CaV2.1; Fig. 3-10 E) or total flux (-0.7 ± 0.2

nC/F; p < 0.001 vs. that for V-CaV2.1; Fig. 3-10 F)

GV-58 increased CaV2.1 open probability by both shifting the voltage-

dependence of activation and delaying channel closure. We perfused GV-58 onto cells

expressing either V-CaV2.1 or V-CaV2.1 R1624P to see if GV-58 could increase Ca2+

flux evoked by an action-potential like waveform. Indeed, in V-CaV2.1, GV-58 robustly

increased both current amplitude (+272.5 ± 33.3%) and charge flux (+282.6 ± 29.0%) (n

= 8; both p < 0.001; Fig. 3-10 A-C). Similar results were observed in cells transfected

with V-CaV2.1 R1624P, where GV-58 augmented both current amplitude (+315.6 ±

59.8%) and charge flux (+288.5 ± 34.7%) (n = 7; both p < 0.001; Fig. 3-10 D-F).

37

Figure 3-10. GV-58 increases Ca2+ flux via both CaV2.1 and CaV2.1 R1624P in

response to an action potential like waveform. Ca2+ currents recorded before (1) and

during (2) application of 12.5 M GV-58 to tsA-201 cells expressing either V-CaV2.1 (A) or V-

CaV2.1 R1624P (D). In both cases, Ca2+ currents were evoked by an action potential-like

waveform similar to that used by Bahamonde et al. (2015). Specifically, this stimulus consisted

of a 1 ms rising phase from -80 mV to +30 mV followed by an 1 ms decline back to -80 mV

(illustrated at the top of panels A and D). Comparison of current amplitudes (B) and total charge

flux (C) for cells expressing V-CaV2.1 in the absence (gray box) or presence (black box) of GV-

58 (n = 8). Comparison of current amplitudes (E) and total charge flux (F) for cells expressing

V-CaV2.1 R1624P in the absence (gray box) or presence (black box) of GV-58 (n = 7). Means

and medians are indicated by the dashed and solid lines of the boxes, respectively. Boxes

represent the 25th/75th percentiles. Bars represent the 5th/95th percentiles. Significant

differences by two-tailed, paired t-test are indicated.

38

Discussion

V-CaV2.1 R1624P expressed in tsA-201 cells was efficiently trafficked to the

membrane and conducted both Ca2+ and Ba2+ current in response to membrane

depolarization (Fig. 3-2, 3-3). However, the R1624P mutation generated a ~25 mV

depolarizing shift in the voltage-dependence of channel activation in both Ca2+ and Ba2+

and slowed channel activation kinetics. The mutation also did not affect the voltage-

dependence of channel deactivation, though the shift in activation made these effects

difficult to parse out. Over extended timescales, the mutant channels also inactivated

more rapidly, an attribute that has more biophysical relevance than physiological since

the channels did not inactivate differently in response to repetitive or closed-state

stimulation. The implications of the shifts in voltage-dependence of activation and in

opening kinetics of the channel lend support to the notion that an action potential would

gate significantly fewer CaV2.1 R1624P channels than wild-type channels. Indeed, upon

depolarization with an action potential-like stimulus, CaV2.1 R1624P did not produce a

significant inward current while its wild-type counterpart did.

CaV2.1 is the predominant mediator of synaptic vesicle release and

neurotransmission at many central synapses and the neuromuscular junction. An

impairment in the ability of CaV2.1 channels to conduct Ca2+ in response to a neuronal

action potential would have profound effects on nervous communication. Indeed, in the

case of the patient with CaV2.1 R1673P, decreased responsiveness to an action

potential because of a depolarizing shift in activation and delayed activation kinetics

may explain some of the symptoms of the condition. The hypotonia that is associated

with the disorder could be linked to synaptic failure at the neuromuscular junction.

39

Similarly, the ataxia, cerebellar atrophy, and global developmental delay observed in

patients with the R1673P mutation are likely due to synaptic failure within central

synapses.

Though we find the R1673P mutation to elicit a profound loss-of-channel

function, our results differ with a previous group that characterized the mutation as a

gain-of-function on the basis of the ability of the mutation to rescue the photoreceptor

response in CaV2.1-null larvae (Luo et al., 2017). There are several potential

explanations for the seemingly dichotomous results of the two studies. One, the mutant

channel may have had higher levels of expression than the wild-type in the fly larvae, or

other CaV isoforms may have compensated for lack of wild-type CaV2.1 expression. In

any case, further investigation into why the results of the two approaches were different

is certainly warranted.

Loss-of-function mutations in CaV2.1 have classically been linked to Episodic

Ataxia Type 2. The R1673P mutation generates a similar phenotype to EA2, though the

disorder is more severe and persistent rather than paroxysmal. In both cases, the

impairment of CaV2.1 function likely weakens neurotransmission and thus produces

mobility issues. Another mutation which produces a depolarizing shift in CaV2.1

activation (Y1662N) was used to generate a zebrafish line expressing the mutant

CaV2.1 (Wen et al., 2013). In this model, both reduced end-plate currents and

significantly impaired mobility were observed, thought the larvae were viable.

Researchers administered Roscovitine, a compound previously shown to possess

CaV2.1 agonist properties (Buraei et al., 2007; Wen et al., 2013). Roscovitine was able

to rescue both end-plate currents and swimming behavior. Intrigued by the agonist

40

action of Roscovitine, we examined whether the drug could augment Ca2+ flux via

CaV2.1 R1624P. Like in the zebrafish study, Roscovitine slowed channel deactivation

and shifted the voltage-dependence of activation to more hyperpolarized potentials.

Roscovitine is a cyclin-dependent kinase inhibitor used as an experimental

chemotherapeutic (Yan et al., 2002). As such, the drug has several off-target effects

that dampen its clinical promise. Namely, Roscovitine both inhibits and stabilizes the

open state of N- and R-type Ca2+ channels, inhibits L- and T-type Ca2+ channels and

blocks KV4.2, KV2.1 and KV1.3 K+ channels (Buraei, Anghelescu, & Elmslie, 2005;

Buraei et al., 2007; Yan et al., 2002; Yarotskyy & Elmslie, 2007, 2012). Another

confounding factor in the action of Roscovitine is that it decreases Ca2+ flux through

CaV2.1 R1624P at more depolarizing potentials (Fig. 3-8 C).

Because of the problems associated with Roscovitine, we decided to test its

higher affinity analog, GV-58, to see if it had similar effects on both wild-type CaV2.1

and CaV2.1 R1624P. In these experiments, GV-58 slowed deactivation and did not

significantly decrease conductance at more depolarizing potentials. Additionally, GV-58

elicited more of a hyperpolarizing shift in the voltage-dependence of CaV2.1 R1624P

activation than Roscovitine did (Fig. 3-9 H vs. Fig. 3-8 C). The changes that GV-58

confers to both CaV2.1 activation and deactivation enhances the channel’s ability to

conduct Ca2+ in response to a neuronal action potential. Though GV-58 is more

effective than Roscovitine in its CaV2.1 agonist action, more work must be done to see if

the off-target effects of Roscovitine persist in GV-58.

Nevertheless, it is clear that CaV2.1 agonists may play in an important role in

treating diseases like the one caused by R1673P that result in a loss-of-channel

41

function. EA2 patients have been treated with the carbonic anhydrase inhibitor

acetazolamide, which has had some moderate success in blunting paroxysmal attacks.

Acetazolamide increases cycling of synaptic vesicles, but the patient with the CaV2.1

R1673P mutation is unresponsive to acetazolamide therapy. Since acetazolamide acts

on vesicle cycling, it may be possible that the R1673P mutation precludes the action of

the drug by not allowing adequate Ca2+ flux into the presynaptic terminal. Thus, drugs

like GV-58 which seek to increase CaV2.1 open probability at the synapse or those that

increase the magnitude/duration of the action potential may be more effective therapies

for disorders resulting from mutations like R1673P. The K+ channel blocker 3,4-

diaminopyridine (amifampridine) is an FDA approved treatment for Lambert-Eaton

Myasthenic Syndrome, an autoimmune disorder characterized by autoantibodies

against CaV2.1 (Wen et al., 2013). By blocking K+ current before the synaptic bouton,

3,4-diaminopyridine can increase the duration and magnitude of a neuronal action

potential, thus gating more CaV2.1 channels at the presynaptic terminal. Intriguingly,

3,4-diaminopyridine rescued some of the impaired motility and synaptic function present

in zebrafish with the Y1662N mutation (Wen et al., 2013). Clearly, more research is

necessary to determine if a combination of synaptic vesicle cyclers, CaV2.1 agonists,

and K+-channel blockers can alleviate some of the dysfunction present in patients with

the R1673P mutation or a related disorder.

A limitation of our heterologous expression system is that we were unable to

investigate the effects of different drug regimens on a level beyond that of the

presynaptic terminal. It would be useful to examine how these potential therapies impact

both synaptic function and motor behavior. Though this preliminary work in tsA-201 cells

42

is essential in developing a thorough understanding of the biophysical and

pharmacological properties of the CaV2.1 mutation, expressing the channel in a

neuronal context may yield different results that further inform study of the channel and

treatment of the linked disease.

The next steps in researching this channel involve generation of a humanized

zebrafish model of the R1673P mutation. A zebrafish line expressing CaV2.1 R1673P

would open the door to electrophysiological, behavioral, and pharmacological study of

the mutation in a vertebrate model of the disorder.

43

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