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University of Calgary
PRISM: University of Calgary's Digital Repository
Graduate Studies The Vault: Electronic Theses and Dissertations
2013-12-04
The Role of L- and T-type Ca2+ Channels in Rat
Cerebral Arteries
Abd El-Rahman, Rasha
Abd El-Rahman, R. (2013). The Role of L- and T-type Ca2+ Channels in Rat Cerebral Arteries
(Unpublished doctoral thesis). University of Calgary, Calgary, AB. doi:10.11575/PRISM/28330
http://hdl.handle.net/11023/1172
doctoral thesis
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UNIVERSITY OF CALGARY
The Role of L- and T-type Ca2+ Channels in Rat Cerebral Arteries
by
Rasha R. Abd El-Rahman
A THESIS
SUBMITTED TO THE FACULTY OF GRADUATE STUDIES
IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE
DEGREE OF DOCTOR OF PHILOSOPHY
DEPARTMENT OF CARDIOVASCULAR AND RESPIRATORY SCIENCES
CALGARY, ALBERTA
November, 2013
Rasha R. Abd El-Rahman 2013
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Abstract
The overall goal of this thesis was to identify which voltage-gated Ca2+ channels are
expressed in rat cerebral arterial smooth muscle and to determine their contributions to
myogenic tone regulation. We began by exploring which voltage-gated Ca2+ channels are
expressed in cerebral arterial smooth muscle. A combination of molecular,
electrophysiological and functional measurements revealed the presence of L- (CaV1.2) and
T-type (CaV3.1 and CaV3.2) Ca2+ channel subtypes in rat cerebral arteries. Both types
contribute to arterial tone development, although the contribution of the L-type channels to
tone development is greater. We then investigated the role of a specific T-type Ca2+ channel
subtype, CaV3.2, in cerebral arterial smooth muscle by functional assessment and a structural
approach using immunohistochemistry, proximity ligation assay, electron-tomography, and
immunogold labeling, combined with computational modeling and electrophysiological
measurements. Results indicate that Ca2+ influx through CaV3.2 channels elicits dilation by
activating ryanodine receptors and inducing Ca2+ sparks, localized events that activate BKCa
channels. In conclusion this work provided evidence for the presence of different types of
voltage-gated Ca2+ channels and provided evidence of their diverse functional roles in
regulating myogenic tone in rat cerebral arteries. Overall, the conclusions indicate the
importance of the different functional roles of voltage-gated Ca2+ channels, which have
substantial physiological relevance to the function of the cerebral
vasculature.………………………………………..
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Acknowledgments
This thesis work would not have been possible without the help, support and
encouragement of many wonderful people. I would like to express my sincere gratitude to
my interim supervisor, Dr. Gary Kargacin, whose expertise, and patience, added
considerably to my graduate experience. I thank him for always being available when I
needed him the most. I am forever grateful to Dr. Meg Kargacin and Dr. Michael Walsh for
their invaluable guidance, help in editing my thesis, and encouragement. It is a pleasure to
thank those who made this thesis possible, and I owe my deepest gratitude to my supervisory
committee members, Dr. Michael Walsh, and Dr. William C. Cole, who have provided me
with valuable feedback and insight for my projects, my thesis and their tremendous input and
many stimulating discussions, whom without their help, I could not proceed with this
journey. All of my appreciation goes to Dr. Andy Braun for his great help and support to
make this thesis come together. Special thanks go to Dr. Ray Turner and his technician Mirna
Kruskic for their great help and support in teaching me and answering my questions. I would
like to extend my warm gratitude to my friend and colleague Dr. Rania Mufti for her great
support, advice and guidance. I also would like to thank my past and present colleagues Dr.
Kevin Luykenaar, Dr. Cam Ha Tran and Osama Harraz for providing me with their support
and suggestions during these years. Thank you for being there for me at all times and keeping
me motivated and inspired.
Lastly, I would not be where I am without the love and support of my family and words
cannot possibly capture my deep gratitude. First and foremost, I wish to thank my beloved
parents for their unconditional love, for raising me to be the person that I’m now, for giving
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me the freedom to pursue my dreams and for being there whenever I need guidance and
support. Mom, I wish you were here but I know that you are watching over me. Last, but by
no means least, I would like to thank my husband Ehab Ali whose love, support and
encouragement allowed me to finish this journey, and my lovely daughters Malak Ali and
Laila Ali, who helped and taught me a lot during this journey.
My thanks to many of my friends Dr. Eman Afkar, Naglaa Ali, Rasha Saleh, Nermeen
Sharaf, and Abeer Ahmad, for cheering me up when I was down and for being my supporters
but also my critics and most of all for their friendship.
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Dedication
I dedicate this thesis to the soul of my beloved mom and my dad, my husband Ehab,
and my two lovely daughters (Malak & Laila), my brothers Mohamed and Ahmad, without
whom this journey would have been impossible.
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Table of Contents
Abstract ................................................................................................................................ ii
Acknowledgments ............................................................................................................... iii
Dedication ............................................................................................................................. v
Table of Contents ................................................................................................................. vi
List of Tables ....................................................................................................................... x
List of Figures and Illustrations .......................................................................................... xi
List of Symbols, Abbreviations and Nomenclature ........................................................... xiii
CHAPTER ONE: INTRODUCTION .................................................................................. 1
1.1 Overview .................................................................................................................... 1
1.2 Role of Ca2+ in smooth muscle (Ca2+ homeostasis). .................................................. 3
1.3 Ca2+ channels ............................................................................................................. 7
1.3.1 Non-voltage-gated Ca2+ channels ................................................................... 7
1.3.2 Voltage-gated Ca2+ channels (VGCCs) .......................................................... 9
1.3.3 General structure of VGCCs .......................................................................... 9
1.3.4 L-type channel structure ............................................................................... 10
1.3.5 T-type channel structure ............................................................................... 13
1.3.6 General functions of VGCCs ....................................................................... 14
1.3.7 L-type channel (CaV1.2) function................................................................. 15
1.3.8 T-type channel (CaV3.1 and CaV3.2) function ............................................ 15
1.4 Potassium channels in smooth muscle ...................................................................... 18
1.5 Sarcoplasmic reticulum (SR) .................................................................................... 19
1.5.1 Inositol 1,4,5-trisphosphate receptors (IP3R) .............................................. 19
1.5.2 Ryanodine receptors (RyR) .......................................................................... 20
1.5.3 Ca2+ sparks .................................................................................................. 22
1.6 Summary ................................................................................................................... 23
1.7 Hypothesis ................................................................................................................ 24
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CHAPTER TWO: IDENTIFICATION OF L- AND T-TYPE CA2+ CHANNELS IN RAT
CEREBRAL ARTERIES: ROLE IN MYOGENIC TONE DEVELOPMENT…………...25
2.1 Introduction ............................................................................................................... 26
2.2 Materials and Methods .............................................................................................. 28
2.2.1 Animal procedures ........................................................................................... 28
2.2.2 Vessel myography ........................................................................................... .28
2.2.3 Isolation of arterial smooth muscle cells .......................................................... 29
2.2.4 PCR analysis .................................................................................................... 30
2.2.5 Western blotting .............................................................................................. 31
2.2.6 Electrophysiology ............................................................................................ 32
2.2.7 Computational blood flow modeling ............................................................... 33
2.2.8 Statistical analysis ........................................................................................... 34
2.2.9 Solutions and chemicals .................................................................................. 34
2.3 Results ....................................................................................................................... 34
2.3.1 Ca2+ channel α1-subunit expression in middle and posterior cerebral
arteries…………………………………………………………………………….....34
2.3.2 Whole-cell Ba2+ currents in cerebral arterial smooth muscle cells .................. 35
2.3.3 Ca2+ channel blockers and myogenic tone ....................................................... 38
2.4 Discussion ................................................................................................................. 40
2.4.1 Background ...................................................................................................... 41
2.4.2 Molecular and electrical characterization ......................................................... 42
2.4.3 T-type Ca2+ channels and myogenic tone ......................................................... 44
2.4.4 Implications to blood flow control ................................................................... 46
2.4.5 Summary .......................................................................................................... 46
2.4.6 Limitations and future directions………………………………………….…...47
(i) Molecular and biochemical approach…………………...……………………...47
(ii) Patch-clamp electrophysiological approach..…………………………….……51
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(iii) Vessel myography approach……………………………………………….…..54
(iv) Computational approach………………………………………………….……55
2.5 Figures ....................................................................................................................... 57
CHAPTER THREE: CaV3.2 CHANNELS AND THE INDUCTION OF A NEGATIVE
FEEDBACK RESPONSE IN CEREBRAL ARTERIAL SMOOTH MUSCLE .............. .81
3.1 Introduction ............................................................................................................... 82
3.2 Materials and Methods .............................................................................................. 83
3.2.1 Animal procedures .......................................................................................... 83
3.2.2 Vessel myography and measurements of membrane potentials ....................... 84
3.2.3 Immunohistochemistry ..................................................................................... 85
3.2.4 Electron tomography ........................................................................................ 86
3.2.5 Immunogold-labeling ....................................................................................... 87
3.2.6 Isolation of arterial smooth muscle cells .......................................................... 88
3.2.7 Proximity ligation assay (PLA) ........................................................................ 88
3.2.8 Computational modeling .................................................................................. 89
3.2.9 Electrophysiology ............................................................................................. 90
3.2.10 Ca2+ sparks ..................................................................................................... 91
3.2.11 Statistical analysis .......................................................................................... 92
3.2.12 Solutions and chemicals ................................................................................. 92
3.3 Results ....................................................................................................................... 93
3.3.1 Vasomotor and electrical effects of CaV3.2 blockade ...................................... 93
3.3.2 Ca2+ channel localization .................................................................................. 93
3.3.3 Computational modeling ................................................................................. 97
3.3.4 BKCa currents and Ca2+ sparks ......................................................................... 97
3.3.5 Functional implications of CaV3.2 blockade .................................................... 98
3.4 Discussion ................................................................................................................. 99
3.4.1 Background ..................................................................................................... 99
3.4.2 Mechanism of CaV3.2-induced vasodilation .................................................. 100
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3.4.3 Summary ........................................................................................................ 103
3.4.4 Limitations and future directions ................................................................... 104
(i) Vessel myography approach ............................................................................ 104
(ii) Structural approach .......................................................................................... 105
(iii) STOCs and Ca2+ sparks ................................................................................. 109
(iv) Computational approach ................................................................................. 111
3.5 Figures ..................................................................................................................... 112
CHAPTER FOUR: DISCUSSION ................................................................................... 128
4.1 Overview ................................................................................................................. 128
4.2 Objective #1: Identification of L- and T-type Ca2+ channel subtypes in rat
cerebral arterial smooth muscle cells and their roles in myogenic tone
development ........................................................................................................... .129
4.3 Objective #2: CaV3.2 and the induction of a negative feedback response in
cerebral arterial smooth muscle ......................................................... ……………..131
4.4 Future directions, experimental considerations, clinical relevance, and
conclusion…………………………………………………………………….........134
4.4.1 Future directions ............................................................................................. 134
4.4.2 Clinical relevance .......................................................................................... 144
4.4.3 Conclusion ...................................................................................................... 145
APPENDIX ....................................................................................................................... 146
REFERENCES ................................................................................................................. 153
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List of Tables
Table 1. IC50 values for inhibition of VGCCs by various blockers………………………....17
Table 2. Immunogold-labelling for RyR and CaV3.2……………………………………….96
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List of Figures and Illustrations
Figure 1. Structure of arteries…………………………………………………………….…2
Figure 2. Regulation of smooth muscle contraction through the phosphorylation
and dephosphorylation of the myosin II regulatory light chain (LC20) by
MLCK and MLCP, respectively……………………………………………….....5
Figure 3. The topology of the high voltage-gated calcium channels.……………………...11
Figure 4. mRNA expression of the α1-subunits of voltage-gated Ca2+ channels….….......57
Figure 5. Protein expression of the α1-subunits of voltage-gated Ca2+ channels……….....59
Figure 6. Inward Ba2+ currents in cerebral arterial smooth muscle cells……………….....61
Figure 7. Nifedipine-sensitive and -insensitive Ba2+ currents in cerebral
arterial smooth muscle cells……………………………………………………..63
Figure 8. Effects of mibefradil on the nifedipine-insensitive component of the
inward Ba2+ current……………………………………………………………...65
Figure 9. Effects of kurtoxin on the nifedipine-insensitive component of the
inward Ba2+current….………………………………………………………......67
Figure 10. Effects of efonidipine on the nifedipine-insensitive component of the
inward Ba2+ current…………….…………………………………..…………….69
Figure 11. The effects of nifedipine and mibefradil on myogenic tone……………..….…..71
Figure 12. The effects of nifedipine and kurtoxin on myogenic tone…………….…..….…73
Figure 13. The effects of nifedipine and efonidipine on myogenic tone…………………...75
Figure 14. The effects of nifedipine and mibefradil on myogenic tone through a full
range of intravascular pressures……………………………………………...….77
Figure 15. Computational modeling predicts that T-type Ca2+ channel modulation
alters tissue blood flow……………………………………………………….....79
Figure 16. Effect of Ni2+ on pressure-induced tone and membrane potential (Vm)
in rat cerebral arteries.……………………………………………………........112
Figure 17. Ca2+ channel localization in rat cerebral arteries…..…………………………....114
Figure 18. Controls of smooth muscle actin, CaV3.2, RyR, and CaV1.2……...……………116
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Figure 19 Proximity ligation assay and the colocalization of CaV3.2 and RyR in
rat cerebral arterial smooth muscle cells……………………………………….118
Figure 20. Electron microscopic imaging of rat cerebral arterial smooth muscle cells…..120
Figure 21. A microdomain model of smooth muscle Ca2+ dynamics……………………..122
Figure 22. Spontaneous transient outward currents (STOCs) and Ca2+ spark
production in cerebral arterial smooth muscle cells……………………………124
Figure 23. Effect of Ni2+ and paxilline on myogenic tone in rat cerebral arteries………..126
Figure 24. Diagram highlighting the proposed function of CaV3.2 channels in
the regulation of arterial tone………………………………………………....133
Figure 25. Diagram highlighting the different proposed possible functions
of CaV3.2 channels in the regulation of arterial tone………………………....142
Figure 26. Diagram highlighting the possible separate functions of CaV3.1 and
CaV3.2 channels in the regulation of arterial tone…………………………….143
Figure 27. CaV3.1 protein expression in vascular smooth muscle cells……………………147
Figure 28. BKCa protein expression in vascular smooth muscle cells……………………..149
Figure 29. IP3R protein expression in vascular smooth muscle cells………………….......151
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List of Symbols, Abbreviations and Nomenclature
ATP Adenosine 5´-triphosphate
BKCa Large conductance calcium-activated potassium channels
[Ca2+]i Cytosolic free Ca2+ concentration
CaM Calmodulin
cAMP Cyclic adenosine 3´:5´-monophosphate
cGMP Cyclic guanosine 3´:5´-monophosphate
CICR Ca2+-induced Ca2+ release
CNS Central nervous system
DAG Diacylglycerol
DMSO Dimethyl sulfoxide
DTT Dithiothreitol
EGTA Ethylene glycol tetraacetic acid
EM Electron microscope
FRET Fluorescence resonance energy transfer
GPCR G protein-coupled receptor
HEK Human embryonic kidney cells
HRP Horseradish peroxidase
IC50 The half maximal inhibitory concentration
IP3 Inositol 1, 4, 5-trisphosphate
IP3R IP3 receptor
KATP ATP-sensitive potassium channel
KIR Inward rectifier potassium channel
KV Voltage-gated potassium channel
MLC20 20-kDa myosin regulatory light chain
MLCK Myosin light chain kinase
MLCP Myosin light chain phosphatase
MYPT1 Myosin phosphatase targeting subunit 1
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NCX Na+/Ca2+ exchanger
PBS Phosphate-buffered saline
PIP2 Phosphatidylinositol-4,5-bisphosphate
PKA cAMP-dependent protein kinase (protein kinase A)
PKC Protein kinase C
PKG cGMP-dependent protein kinase (protein kinase G)
PLC Phospholipase C
PLCγ1 Phospholipase Cγ1 isoform
PP1c Phosphatase type I catalytic subunit
PSS Physiological salt solution
ROK Rho-associated protein kinase
ROCs Receptor-operated channels
RyR Ryanodine receptor
SACs Stretch-activated channels
SMC Smooth muscle cells
SDS-PAGE Sodium dodecyl sulfate-polyacrylamide gel electrophoresis
SOCCs Store-operated cation channels
SR Sarcoplasmic reticulum
STIC Spontaneous transient inward current
STIM1 Stromal interaction molecule 1
STOC Spontaneous transient outward current
TRP Transient receptor potential
TIC Transient inward current
TMEM16A Ca2+-activated Cl- conductance
VGCC Voltage-gated Ca2+ channel
Vm Membrane potential
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Chapter One: INTRODUCTION
1.1 Overview.
The cardiovascular system consists of the heart, blood, and blood vessels. The principal
function of blood vessels is to transport blood throughout the body. There are three main
types of blood vessels: the arteries, which carry blood away from the heart; the capillaries,
where exchange of water and chemicals between the blood and the tissues occurs; and the
veins that transmit blood from the capillaries back to the heart. Arterial blood vessels can be
further divided into: 1) conduit arteries (e.g. aorta) that conduct blood to peripheral tissues;
and 2) resistance vessels (e.g. cerebral and feed arteries) that regulate blood flow to various
organs depending upon their physiological needs (Segal & Jacobs, 2001). Conduit vessels
are thought to change their diameter passively in response to the volume of blood entering or
leaving them. Resistance arteries, on the other hand, actively change diameter in response to
a variety of stimuli including: 1) shear stress and intraluminal pressure (Davies, 2009; Knot
& Nelson, 1995; Segal, 2000); 2) sympathetic neuronal outputs (Brayden & Bevan, 1985;
Lee, 1982; Si & Lee, 2002); and 3) hormonal and metabolic factors (Filosa et al., 2006;
Zhang et al., 2005). Arteries are comprised of three layers including: 1) the tunica adventitia,
an outer layer of connective tissue; 2) the tunica media, a layer primarily containing smooth
muscle cells; and 3) the tunica intima, an inner layer of endothelial cells lining the blood
vessel wall (Figure 1; Faraci & Heistad, 1990).
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Figure 1: Structure of arteries. The arterial wall is composed of three layers: an inner layer, the
tunica intima, composed mainly of endothelial cells; a middle layer, the tunica media, with
smooth muscle cells as the main component; and an outer layer, the tunica adventitia, composed
mainly of collagenous tissue (Davis et al., 2011).
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1.2 Role of Ca2+ in smooth muscle (Ca2+ homeostasis).
Ca2+ is an essential second messenger involved in controlling and regulating numerous
cellular functions and reactions including muscle contraction, fertilization, cell locomotion, gene
expression, and apoptosis. Because high levels of Ca2+ are toxic to cells and can lead to cell
death, intracellular Ca2+ is carefully regulated (Wray et al., 2005). Ca2+ signaling in vascular
smooth muscle is complex, however, in recent years, advanced technologies have provided tools
to better understand the roles and regulation of Ca2+ in different cells and tissues (Carafoli et al.,
2001; Wray et al., 2005). Improved cell culture methods, the availability of high-affinity
fluorescent Ca2+ indicators (including genetically-encoded indicators based on green fluorescent
proteins and membrane-permeant small molecules, e.g. Fura-2, and Fluo-4) combined with high-
resolution fluorescent microscopy have allowed significant insights into intracellular Ca2+
signaling (Knot et al., 2005). The intracellular free Ca2+ concentration [Ca2+]i ranges from resting
levels of 100-200 nM up to levels of ~700 nM during contraction while extracellular [Ca2+]free is
1-2 mM. Thus Ca2+ will be favoured to enter the cell and [Ca2+]i must be regulated by various
ion channels, exchangers and pumps found in the plasmalemmal and sarcoplasmic reticulum
(SR) membranes of smooth muscle cells (Berridge et al., 2000; Carafoli et al., 2001; Wray et al.,
2005). Smooth muscle cells are responsible for the contractility of all hollow organs, including
blood vessels, the gastrointestinal tract, the bladder and the uterus. In resistance arteries, smooth
muscle cells actively relax and contract to a variety of stimuli that effectively change vessel
diameter and consequently regulate tissue blood flow. Increases in [Ca2+]i elicit smooth muscle
contraction initiated by the binding of Ca2+ to calmodulin (CaM) and the activation of myosin
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light chain kinase (MLCK). In general, the smooth muscle literature recognizes two principal
mechanisms responsible for the rise in [Ca2+]i. The first involves the activation of a G protein-
coupled receptor (GPCR) by either a neurotransmitter or hormone, where the stimulation of the
receptor activates phospholipase C (PLC) that in turn evokes the production of inositol 1,4,5-
trisphosphate (IP3) and diacylglycerol (DAG). The binding of IP3 to IP3 receptors (IP3R) on the
SR leads to Ca2+ release from the SR through the receptors. The second mechanism involves an
increase in resting membrane potential and the opening of voltage-gated Ca2+ channels
(VGCCs). This induces an influx of extracellular Ca2+ (Figure 2; Knot & Wilson, 1998). This
mechanism has long been proposed to play an essential role in evoking contraction of vascular
smooth muscle (Knot & Nelson, 1998; Welsh et al., 2002; Welsh et al., 2000). There are several
types of VGCC including L-type (e.g. CaV1.2) and T-type channels (e.g. CaV3.1 and CaV3.2) that
are believed to be expressed in vascular smooth muscle cells (Abd El-Rahman et al., 2013; Kuo
et al., 2010; Nikitina et al., 2007).
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Figure 2: Regulation of smooth muscle contraction through the phosphorylation and
dephosphorylation of the myosin II regulatory light chain (LC20) by MLCK and MLCP,
respectively. The influx of Ca2+ through VGCC elevates global cytosolic [Ca2+] and can also
induce Ca2+ release from the SR. Ca2+ binding to calmodulin activates MLCK and
phosphorylates MLC20 to induce contraction by cross-bridge cycling between actin and myosin.
Meanwhile, dephosphorylation of MLC20 by MLCP elicits relaxation. Signaling pathways via G
protein-coupled receptors (GPCR): 1) Gq/11 activates phospholipase Cβ (PLCβ), which converts
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phosphatidylinositol 4,5-bisphosphate (PIP2) into inositol 1,4,5-trisphosphate (IP3) and
diacylglycerol (DAG). IP3 activates Ca2+ release from the SR via inositol 1,4,5-trisphosphate
receptors (IP3R). DAG that activates protein kinase C (PKC); 2) G12/13 is coupled to RhoGEF,
which activates RhoA, which in turn stimulates Rho-associated kinase (ROK). PKC and ROK
phosphorylate CP1-17, and the myosin phosphatase targeting subunit 1 (MYPT1) of MLCP,
respectively, to inhibit MLCP and elicit Ca2+ sensitization of contraction seen as a leftward shift
in the force vs. [Ca2+]i curve in the lower left in the figure (Cole & Welsh, 2011).
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1.3 Ca2+ channels.
Ca2+ ion entry into the cytosol mediates a wide range of cellular responses, including the
activation of muscle contraction, regulation of Ca2+-dependent enzymes, regulation of gene
transcription, and apoptosis (Dolmetsch et al., 2001; Kisilevsky & Zamponi, 2008; Martin-
Moutot et al., 1996; Sutton et al., 1999). Ca2+ may enter cells by a number of different
mechanisms, including non-voltage gated Ca2+ channels and voltage-gated Ca2+ channels
(VGCCs).
1.3.1 Non-voltage-gated Ca2+ channels.
Non-voltage-gated Ca2+ channels include receptor-operated channels (ROCs), store-
operated Ca2+ channels (SOCCs), and mechano-sensitive or stretch-activated channels (SACs)
(Burnstock, 1972; Benham et al, 1985; Loirand et al, 1991; Fleischmann et al, 1997). ROCs are
regulated by agonist–receptor interaction and they are activated by many different agonists (e.g.
acetylcholine, angiotensin II, endothelin-1, norepinephrine, serotonin and vasopressin) acting
through plasma membrane receptors. They also include the ligand-gated channel P2X that is
activated by ATP (Sanders, 2001; Benham et al, 1985). ROC current is a non-selective cation
current (Icat) that is carried by both mono- and divalent cations with differing degrees of Ca2+
selectivity (Sanders, 2001). SOCCs are activated by the emptying of intracellular Ca2+ stores
(Berridge, 1995; Hirota et al., 2007; Jones & Braun, 2009; Lee et al., 2005). Growing evidence
suggests that proteins, such as Orai1 and STIM1, are components of SOCCs. Orai1 has been
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identified as the main pore-forming subunit of the SOCCs (Guibert et al., 2008; Muik et al.,
2011).
Stretch-activated channels (SACs) are activated by mechanical stimulation of smooth
muscle cells such as compression, stretch and shear stress. SACs are mechano-transducers that
convert physical forces into biological signals and cell responses. Cation influx through SACs
induces membrane depolarization that subsequently activates VGCCs to increase [Ca2+]i (Yin &
Kuebler, 2010). Several types of transient receptor potential (TRP) channels are expressed in
smooth muscle cells and have been linked to SAC activity (Gonzales et al., 2010b; Gonzalez-
Cobos & Trebak, 2010; Ramsey et al., 2006; Yang et al., 2006). TRP channels facilitate Ca2+
and Na+ influx into the cells, raise [Ca2+]i and [Na+]i respectively, and depolarize the plasma
membrane. The superfamily of TRP channels can be divided into several families and
subfamilies: three main subfamilies, TRPC (canonical), TRPV (vanilloid) and TRPM
(melastatin), and several subfamilies including TRPP (polycystin), TRPML (mucolipin), TRPA
(ankyrin) and TRPN. TRPC, TRPM, and TRPV display mechanosensitive properties while
TRPN channels do not (Earley, 2010; Gonzalez-Cobos & Trebak, 2010; Muraki et al., 2003;
Ramsey et al., 2006; Yang et al., 2006; Yin & Kuebler, 2010). These channels exhibit different
relative permeabilities to cations (Ca2+, Na+, K+, Mg2+, Mn2+, Cs+, Li+). While all TRP channels
are permeable to cations, only two TRP channels are impermeable to Ca2+ (TRPM4 and
TRPM5), and two others are highly Ca2+ permeable (TRPV5 and TRPV6; Owsianik et al.,
2006). TRP channel pharmacology is complex, and is different in different tissues and species
(Earley et al., 2004; Ramsey et al., 2006; Welsh et al., 2002).
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1.3.2 Voltage-gated Ca2+ channels (VGCCs).
Voltage-gated Ca2+ channels are expressed in many tissues and cells, e.g. the central and
peripheral nervous system and smooth muscle cells, where they demonstrate a wide range of
physiological, pharmacological and electrophysiological properties. Many different factors have
been identified that control VGCCs activity. These include alternate splicing, auxiliary subunit
associations, peptide and small organic blockers, GPCRs, protein kinases and Ca2+-binding
proteins (Braunstein et al., 2009; Catterall, 2000; Hofmann et al., 1999). There are several
different kinds of VGCCs that have different physiological functions and electrophysiological
and pharmacological properties. VGCCs include: 1) the CaV1 group (L-type), which has four
members (CaV1.1-1.4); 2) the CaV2 group, which has three members: CaV2.1 (P/Q-type), CaV2.2
(N-type), CaV2.3 (R-type); and 3) the CaV3 group (T type), which has three members (CaV3.1-
3.3). The CaV1 and CaV2 channels are classified as high-voltage-activated (HVA) channels, due
to their high voltage thresholds of activation, while the CaV3 channels are classified as low-
voltage-activated (LVA) channels (Catterall, 2000; Cheng et al., 2009).
1.3.3 General structure of VGCCs.
VGCCs are hetero-oligomeric protein complexes that have a main pore-forming α1-
subunit of 190–250 kDa that is in association with auxiliary β-, α2δ- and γ-subunits. The α1-
subunit has four membrane-spanning domains (I–IV) that are linked together in a single
polypeptide chain. Each domain contains six transmembrane segments (S1–S6); S4 is the voltage
sensor; a pore-forming loop (P-loop) is formed between S5 and S6. The C- and N-termini are
both intracellular (Figure 3; Catterall, 2000; Altier & Zamponi, 2008). The selectivity filter
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consensus sequence for the Ca2+ pore-forming loop has four glutamic acid repeats (EEEE). The
cytoplasmic loops connecting the four domains are structurally essential for interaction with the
β-subunits and membrane-binding proteins. They are also essential for channel gating and
interaction with second messengers. An auxiliary β-subunit and a transmembrane disulphide-
linked α2δ subunit complex are important components of most types of VGCCs. They are
required for optimal surface expression, channel regulation and gating kinetics (Catterall, 2000;
Hofmann et al., 1999; Marcantoni et al., 2008).
1.3.4 L-type channel structure.
In addition to the pore-forming α1-subunit of CaV1.2 in vascular smooth muscle cells
(VSMCs), other auxiliary subunits have been identified. There are four β subunit isoforms (β1b,
β2, β3, β4), and four α2δ subunit isoforms. The β and α2δ subunits directly interact with the α1-
subunit, the β-subunit modulates activation and inactivation of the channel, while the α2δ-subunit
is necessary for plasma membrane expression of CaV1.2 (Bielefeldt, 1999). There are a number
of splice variants of the α1-subunit of CaV1.2 channels that have been extensively investigated,
and a numbers of diseases have been linked to aberrant splicing. Previous work indicated that the
gene encoding the α1 subunit of CaV1.2 consists of 55 exons and that at least 19 exons undergo
alternative splicing (Liao et al., 2005; Tang et al., 2004).
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Figure 3: The topology of the high voltage-gated calcium channels. The pore-forming α1-
subunit is composed of four domains (I-IV), each consisting of six transmembrane segments (S1-
S6). Domain S4 contains the voltage sensor (labeled +). S5 and S6 segments form the pore of the
channel. β, α2-δ, and γ auxiliary subunits are also shown. The N- and C-termini of the channel
are intracellular (Altier & Zamponi, 2008).
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With the recent discovery of exon 1c at the N-terminus (Cheng et al., 2009), the CaV1.2
gene now is known to contain at least 56 exons of which 20 exons are subject to alternative
splicing. Based on splicing, CaV1.2 channels can be generally segregated into cardiac or smooth
muscle isoforms (Liao et al., 2005; Liao et al., 2007; Liao et al., 2004).
L-type channels are high-voltage-activated (HVA) Ca2+ channels, that activate at voltages
much more positive than the resting membrane potential. Their activation is fast and voltage
dependent, while inactivation is slow (> 1s), but it is more rapid in the presence of Ca2+ and
comparatively slow in the presence of Ba2+ (Catterall, 2000). Deactivation is also quick, ensuring
that the channels close rapidly when the membrane is repolarized to resting levels (˂ 1 ms;
Catterall, 2000). CaV1.2 is two to three times more permeable to Ba2+ than Ca2+, and lacks Ca2+-
dependent inactivation when Ba2+ is used as the charge carrier in electrophysiological recording
solutions (Nikitina et al., 2007). In most studies of vascular smooth muscle, the high Ba2+
concentrations that were used (10-20 mM, compared to the use of a physiological Ca2+
concentration of ~1.8 mM) induced a hyperpolarized shift in the current-voltage (IV) relationship
of CaV1.2 of ~10 mV. This occurs because Ba2+ obstructs the influx of ions. This is referred to
as surface charge screening and is due to the binding of positive ions to the membrane, on a
selective binding site on the channel protein (Kuo et al., 2010; Salemme et al., 2007; Li et al.,
2010). L-type channels are distinct from other CaV channels in their high sensitivity to 1, 4-
dihydropyridines (DHPs). Their blockage by DHPs occurs in a state-dependent manner as a
result of the binding of DHPs to inactivated channels and increases with membrane
depolarization (Dolphin, 2006). Previous studies also showed that the IIIS5, IIIS6 and IVS6
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domains determine the binding of DHPs to the α1-subunit of L-type Ca2+ channels (Liao et al.,
2007).
1.3.5 T-type channel structure.
The structure of the α1-subunit of T-type channels is related to the structure of the α1-
subunit of CaV1.2 channels. Unlike other VGCCs, T-type channels (CaV3.x) do not require
auxiliary subunits for functional expression (Lambert et al., 1997; Leuranguer et al., 1998), and,
to date, the existence of auxiliary subunits (β, α2δ, γ) with CaV3 α1-subunits in vascular smooth
muscle cells has not been reported. The α1-subunit of CaV3.1 consists of 38 exons (Mittman et
al., 1999), and a recent study showed that at least 15 sites undergo alternative splicing in human
brain. The α1-subunit of the CaV3.2 gene consists of 36 exons, and a more recent study revealed
over 12 alternative splicing sites (Zhong et al., 2006). Almost all CaV3 splice variants have been
found in non-smooth muscle cells (Shcheglovitov et al., 2008; Zhuang et al., 2000). As noted
above, the selectivity filter consensus sequence for the pore loops of CaV1.x and CaV2.x has four
repeats of glutamic acid (EEEE), whereas, in the CaV3.x pore loop, two glutamates are replaced
by aspartic acid residues (EEDD). The difference in the amino acid composition of the repeat
segments in the pore loops of L- and T-type channels is thought to account for the differences in
Ca2+ permeability of the channels (Jan & Jan, 1990; Talavera et al., 2001). L-and T-type Ca2+
channels have similar overall sizes, with long loops connecting membrane spanning domains I
and II as well as II and III, and short loops connecting membrane spanning domains III and IV.
In T-type Ca2+ channels, the functions of the loops are unknown, but may contain essential
regulatory sites (e.g. for protein kinases); this is in contrast to HVA channels in which these
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loops are essential for binding auxiliary subunits (Marcantoni et al., 2008; Perez-Reyes, 2003).
Short amino acid sequences connect to the N-terminus, while long amino acid sequences connect
to the C-terminus of T-type channels. T-type Ca2+ channels differ from other voltage-gated Ca2+
channels in their low threshold for activation and inactivation. These channels are characterized
by their activation at relatively hyperpolarized potentials, their fast activation and inactivation
and their slow deactivation. Unlike CaV1.2 channels, they have Ca2+-independent inactivation
kinetics (Perez-Reyes, 2003). T-type Ca2+ channels are almost equally permeable to Ca2+ and
Ba2+, thus, the use of Ba2+ as a charge carrier helps to differentiate between L- and T-type Ca2+
channels when both are expressed in the same cell type (Kuo et al., 2010; Nikitina et al., 2007).
High Ba2+ concentrations (10-20 mM) result in a hyperpolarized shift in voltage-dependence of
T-type channels (Perez-Reyes, 2003). Generally, T-type Ca2+ channels are characterized by their
greater sensitivity to agents such mibefradil, kurtoxin, R(-)efonidipine and Ni2+ (Lee et al., 1999;
Marcantoni et al., 2008; Perez-Reyes, 2003).
1.3.6 General functions of VGCCs.
VGCCs play a key role in mediating many cellular functions. Like other ion channels,
they contribute not only to shaping the action potential and to oscillations of membrane voltage,
but they also control Ca2+ influx through the plasma membrane and are involved in excitation–
secretion, excitation–contraction and excitation-transcription coupling (Dolphin, 2006;
Marcantoni et al., 2008).
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1.3.7 L-type channel (CaV1.2) function.
In cardiac myocytes, Ca2+ influx through L-type Ca2+ channels shapes the plateau phase
of the cardiac action potential. Ca2+ influx triggers Ca2+ release from SR stores via ryanodine
receptors (RyR), a process known as Ca2+-induced Ca2+-release, CICR. L-type Ca2+ channels can
mediate global increases in [Ca2+]i in smooth muscle cells (Lambert et al., 1997; Moosmang et
al., 2003). In vascular smooth muscle, L-type Ca2+ channels (CaV1.2) play an important role in
the development of myogenic tone. Pressurization of cerebral arteries leads to graded membrane
depolarization and opening of CaV1.2 channels. This causes further depolarization and steady
state Ca2+ entry due to the slow inactivation of the channels (Lambert et al., 1997; Knot &
Nelson, 1998; Moosmang et al., 2003). The high selectivity of DHPs for CaV1.2 L-type channels
(Table1) over CaV3 channels has allowed the functional role of CaV1.2 in vascular smooth
muscle cells to be studied. DHPs cause arterial dilation in a state-dependent manner (Dolphin,
2006; Perez-Reyes, 2003) supporting the conclusion that CaV1.2 channels are the major source of
Ca2+ entry in response to membrane potential depolarization. Thus they are key players in the
development of myogenic tone, which is essential for proper smooth muscle contraction and
function (Knot & Nelson, 1998; Moosmang et al., 2003).
1.3.8 T-type channel (CaV3.1 and CaV3.2) function.
The T-type Ca2+ channels produce the pacemaker potential in the SA node of the heart.
Moreover, in the central nervous system (CNS), T-type Ca2+ channels contribute to tonic bursting
activity in the thalamus (Dolphin, 2006; Perez-Reyes, 2003). Their role in vascular smooth
muscle has not yet been completely identified. Recent studies have provided evidence for the
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presence of T-type Ca2+ channels (CaV3.1 and CaV3.2) in resistance arteries (e.g. mesenteric
arteries and renal microcirculation; Abd El-Rahman et al., 2013; Jensen et al., 2004; Kuo et al.,
2010; Navarro-Gonzalez et al., 2009) and suggest that Ca2+ influx through T-type Ca2+ channels
could contribute to a global rise in [Ca2+]i and the development of myogenic tone, especially
when vessels are hyperpolarized. However, experimental support for this is limited and
incomplete (Kuo et al., 2010; Nikitina et al., 2007). Meanwhile the presence of two subtypes of
T-type Ca2+ channels in vascular smooth muscle (CaV3.1, and CaV3.2) raises the question: in
smooth muscle cells do both subtypes function in the same way? Published research regarding T-
type channel function suggests multiple roles for T-type channels. Some results suggest that Ca2+
influx through T-type channels facilitates arterial constriction and other researchers propose that
T-type channels may mediate dilation (Braunstein et al., 2009; Chen et al., 2003; Hansen et al.,
2011; Kuo et al., 2010; Poulsen et al., 2011). The latter possibility is important to investigate
since CaV3.2 knockout (α13.2-/-) mice showed a constricted arterial phenotype (Chen et al.,
2003). This study suggested that Ca2+ influx through CaV3.2 channels activates a Ca2+-dependent
K+ conductance (BKCa) that induces hyperpolarization and, therefore, vasodilation. This
suggested a possible mechanism in which VGCCs induce K+ channel activity and cause
vasodilation in smooth muscle cells. Such a mechanism needs to be extensively examined. At the
present time Ni2+ is the best pharmacological tool available to distinguish between Cav3.1 and
CaV3.2. Ni2+ blocks both subtypes of T-type channels, however, the IC50 of Ni2+ for CaV3.1 is
250 µM and is 12 µM for CaV3.2 (Table1; Lee et al., 1999; Perez-Reyes, 2003); thus a low
concentration of Ni2+ can be used to selectively inhibit the CaV3.2 channel subtype.
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VGCCs
Inhibitor
Channel
IC50
Cell type
Reference
Nifedipine
CaV1.2
CaV3.1
CaV3.2
10-15 nM
3 µM
1.2 µM
CHO
HEK293
HEK
Morel et al. 1998
Akaike et al. 1989
Lee et al. 2006
Mibefradil
CaV1.2
CaV3.1
CaV3.2
3 µM
0.012-0.014 µM
0.069 µM
HEK
HEK293
HEK293
Jimenez et al. 2000
Morita et al. 2002
Martin et al. 2000
R(-)efonidipine
CaV1.2
CaV3.1
CaV3.2
1000 µM
0.1 µM
0.1 µM
BHK
BHK
BHK
Furukawa et al. 2004
Furukawa et al. 2004
Furukawa et al. 2004
Kurtoxin
CaV1.2
CaV3.1
CaV3.2
>10 µM
0.07 µM
0.07 µM
Xenopus oocytes
Cardiomyocytes
Cardiomyocytes
Chuang et al. 1998
Horiba et al. 2008
Horiba et al. 2008
Ni2+
CaV1.2
CaV3.1
CaV3.2
324 µM
250 µM
12 µM
rbU-SMC
HEK293
HEK293
Bradley et al. 2004
Lee et al. 1999
Lee et al. 1999
Table 1. IC50 values for inhibition of VGCCs by various blockers.
Note: VGCCs, voltage-gated Ca2+ channels; CHO, Chinese hamster ovary; HEK, human
embryonic kidney; BHK, baby hamster kidney; rbU, rabbit urethra.
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1.4 Potassium channels in smooth muscle.
In general, potassium (K+) channels are comprised of a pore-forming α-subunit with
cytoplasmic N- and C-termini and contain pore-forming six transmembrane domains (S1–S6).
The S4 domain contains the voltage sensor (Korovkina & England, 2002). Each α-subunit is
associated with auxiliary β-subunits, which influence the biophysical characteristics of the
channel (Bahring et al., 2001). In BKCa channels the α-subunits contain an additional seventh
transmembrane region (S0) at the exoplasmic N-terminus (Tanaka et al., 2004). Potassium
channels are the main channels involved in mediating hyperpolarizing responses and the
induction of relaxation in smooth muscle cells (Ko et al., 2008). In vascular smooth muscle there
are four general types of K+ channels: large conductance Ca2+-activated (BKCa), voltage-gated
(KV), inward rectifier (KIR) and ATP-sensitive (KATP) K+ channels. Although hyperpolarization
is induced by all K+ channels, each K+ channel is regulated differently (Nelson and Quayle,
1995). BKCa channels are both voltage and Ca2+ sensitive (Cox & Petrou, 1999; Nelson &
Quayle, 1995). These channels are involved in inducing vasodilation through a negative
feedback mechanism (Jaggar et al., 1998b; Knot & Nelson, 1995). KV channels open in response
to membrane depolarization and allow K+ efflux, repolarization and a return to the resting
membrane potential. The function of KV channels, therefore, is to limit membrane depolarization
and maintain resting vascular tone (Korovkina & England, 2002). KIR channel activity increases
with membrane hyperpolarization, in contrast to KV and BKCa channels, and it is thought to play
a role in determining the resting membrane potential of vascular smooth muscle (Ko et al.,
2008). KATP channel current is linear in nature, and channel gating is voltage independent; it
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links cell metabolism to electrical activity of the plasma membrane by controlling membrane
potential (Thorneloe & Nelson, 2005; Teramoto, 2006; Ashcroft, 2005).
1.5 Sarcoplasmic reticulum (SR).
The SR in smooth muscle is the major intracellular Ca2+ store and plays a key role in
regulating Ca2+ homeostasis in smooth muscle cells. When the SR is close to the plasma
membrane it is defined as superficial or peripheral and when it is positioned away from the
plasma membrane it is described as deep or central. Ca2+ homeostasis, local Ca2+ release, and
interactions with plasma membrane ion channels, and hence cell excitability, have been
associated with the peripheral SR (Wray & Burdyga, 2010). It has been proposed that the central
SR is involved in providing Ca2+ to the myofilaments to induce contraction (Wray & Burdyga,
2010). The SR elicits relaxation when Ca2+ is sequestered into the SR by Ca2+-ATPase (SERCA)
activity, or constriction through the release of Ca2+. Two major Ca2+ channels are found in the
SR membrane, IP3R and RyR, which contribute to Ca2+ homeostasis, and are involved in
generation of a variety of Ca2+ signals (e.g. Ca2+ waves, sparks and puffs; Wray & Burdyga,
2010) .
1.5.1 Inositol 1, 4, 5-trisphosphate receptors (IP3R).
In addition to Ca2+, IP3Rs require IP3 for activation and SR Ca2+ release. IP3 is produced
by the activation of Gq/11- coupled GPCRs in the plasma membrane. IP3R regulation is complex
as both Ca2+ and IP3 can activate and/or inhibit IP3Rs (Bootman et al., 1995; Chalmers et al.,
2007; Sanders, 2001). There are three isoforms of the IP3R (IP3R1, IP3R2 and IP3R3; Furuichi et
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al., 1989). The IP3R protein has a molecular weight of ~310 kDa and displays three distinctive
regions: 1) the N-terminal ligand-binding region, with a suppressor domain and IP3-binding core;
2) the C-terminal channel-forming region with six transmembrane helices and a pore loop
between TM5 and TM6; and 3) the regulatory region connecting the N- and C-termini (Furuichi
et al., 1989). In vascular smooth muscle, IP3Rs are the main SR Ca2+ release channels. When
GPCRs are activated, IP3 and DAG are generated from PLC-mediated cleavage of PIP2; IP3
opens IP3R channels (Jaggar et al., 2000). Furthermore, Ca2+ release events have been observed
from single IP3Rs and named Ca2+ “blips” (Amberg et al., 2007; Lee et al., 2002; Taylor et al.,
2008). Similar to Ca2+ sparks, produced from RyRs, Ca2+ puffs are generated by the concerted
opening of a cluster of IP3Rs (Furuichi et al., 1989; Mironneau et al., 1996). Ca2+ puffs may
combine with Ca2+ sparks to generate oscillations which propagate as Ca2+ waves through the
cell (Mironneau et al., 1996).
1.5.2 Ryanodine receptors (RyR).
Molecular and biochemical analyses have identified three different RyR isoforms (RyR1,
RyR2 and RyR3), encoded by three different genes on different chromosomes (Coronado et al.,
1994; Fill & Copello, 2002). Expression of RyR1, RyR2 and RyR3 isoforms are predominant in
skeletal muscle, cardiac muscle and brain, respectively (Coronado et al., 1994; Fill & Copello,
2002). All three isoforms are expressed in vascular smooth muscle cells, where RyR2 is the
predominant isoform (Chalmers et al., 2007; Nelson et al., 1995; Wellman & Nelson, 2003).
RyRs are the largest known ion channel proteins (~560 kDa/subunit) and form homotetramers
(Chalmers et al., 2007; Fill & Copello, 2002). The most widely accepted model of the RyR
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describes the RyR channel as an assembly of 6 transmembrane (TM) domains, in which TM5
and TM6 form the pore region with a pore loop having a conserved sequence motif,
GXRXGGGXGD, that contains a ryanodine-binding site and a selectivity filter (Takeshima et
al., 1989; Zhao et al., 1999). About 80% of the mass of the RyR is in the cytosolic domain,
which contains a number of protein-binding sites including sites for calmodulin (CaM) and
protein kinase A (PKA) binding (Bers, 2004). [Ca2+]i has a biphasic effect on RyR activity, it
activates RyRs at micromolar [Ca2+]i (~10 μM) and inhibits RyRs at higher [Ca2+]i (>100 μM;
Chalmers et al., 2007). It has been proposed that upon membrane depolarization in vascular
smooth muscle, Ca2+ enters the cytosol via VGCCs and activates RyR channels by a Ca2+-
induced Ca2+ release (CICR) process, in a manner similar to that described in cardiac muscle
(Chalmers et al., 2007; Sanders, 2001). Ca2+ efflux from activated IP3Rs may also diffuse and
trigger Ca2+ release from neighboring RyRs through CICR (Chalmers et al., 2007; Sanders,
2001). SR Ca2+ through CICR has been proposed as a mechanism for eliciting contraction in
vascular smooth muscle cells (Jaggar et al., 1998b; Nelson et al., 1995). The discovery of local
Ca2+ signaling revealed another mechanism through which RyRs can affect vascular
contractility. Ca2+ sparks constitute one of the main local events that involve Ca2+ release
through RyRs (Nelson et al., 1995; Perez et al., 2001). Ca2+ sparks have been suggested to
activate BKCa channels to elicit hyperpolarization and prevent the vessel from becoming over-
constricted. Thus RyRs may be part of a negative feedback mechanism to control vessel diameter
(Nelson et al., 1995; Perez et al., 2001; Wellman & Nelson, 2003).
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1.5.3 Ca2+ sparks.
Ca2+ sparks are local Ca2+ events caused by openings of RyRs. They were initially
identified in cardiac muscle and subsequently in skeletal and smooth muscle (Jaggar et al., 2000;
Nelson et al., 1995). Improved detection of Ca2+ sparks has allowed these events to be
characterized as having a rise time of ~20 ms, a half-time of decay of 50-60 ms, a spatial spread
of 2.4 μm and a frequency of 1 Hz (Nelson et al., 1995; Perez et al., 2001; Perez et al., 1999).
Modulation of Ca2+ sparks by ryanodine and caffeine indicates that RyR openings are involved
in these events (Knot et al., 1998; Mironneau et al., 1996; Nelson et al., 1995; Perez et al., 2001;
Perez et al., 1999). It is clear in cardiac muscle that the influx of Ca2+ via VGCCs plays a key
role in the generation of Ca2+ sparks via CICR. Evidence for a similar mechanism in vascular
smooth muscle is less clear (Chalmers et al., 2007). In vascular smooth muscle, several studies
indicate that Ca2+ sparks can occur for a period of time in the presence of L-type Ca2+ channel
blockers or in the absence of extracellular Ca2+ (Nelson et al., 1995; Zhao et al., 1999).
However, Ca2+ spark activity can be modulated by L-type channel activity. In intact cerebral
arterial segments, Ca2+ spark frequency and amplitude were seen to be augmented by high
extracellular [K+], suggesting that Ca2+ influx via VGCCs can directly or indirectly regulate Ca2+
sparks (Jaggar et al., 1998a). Ca2+ influx through VGCC could also be important for refilling the
SR store without having a direct effect on the RyR channels. Ca2+ sparks can be generated at a
low frequency in un-stimulated smooth muscle cells, and can activate BKCa channels to produce
a spontaneous transient outward current (STOC; Knot et al., 1998; Nelson et al., 1995; Perez et
al., 1999). In pressurized rat cerebral arteries at physiological membrane potential of -40 mV,
simultaneous measurements of Ca2+ sparks and STOCs indicated that most Ca2+ sparks are
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associated with the initiation of STOCs (Perez et al., 1999). This association occurs despite the
fact that both Ca2+ spark and STOC amplitudes vary (Perez et al., 1999). Although a rise in
[Ca2+] is normally associated with constriction of vascular smooth muscle Ca2+ sparks and
STOCs cause relaxation (Knot et al., 1998). Ca2+ sparks can, therefore, regulate myogenic tone
by activating BKCa channels, which in turn regulates membrane potential, leading to the
inhibition of L-type Ca2+ channels. This limits Ca2+ entry and reduces contraction. Thus, RyR-
mediated Ca2+ sparks control vascular tone through a feedback mechanism that promotes
vasodilation.
1.6 Summary.
L-type Ca2+ channels were the first VGCCs identified in vascular smooth muscle cells.
Subsequently other voltage gated Ca2+ channel types (especially T-type Ca2+ channels) were also
identified. Different functional roles of the T-type Ca2+ channel subtypes have been proposed in
smooth muscle cells, either augmenting the myogenic response by increasing global [Ca2+] or
reducing it through a negative feedback mechanism. In the latter case, it has been suggested that
T-type Ca2+ channels regulate BKCa activity to elicit vessel dilation, however, this has not been
well established. The work described below was designed to identify the T-type Ca2+ channels
present in cerebral arterial smooth muscle cells and to understand their functional role in the cells
especially with regard to tone development. Identification of the different types of VGCCs
expressed and their electrophysiological characteristics and functional roles in cerebral arterial
smooth muscles, will improve our understanding of Ca2+ homoeostasis and of tone in these
arteries.
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1.7 Hypothesis.
Specific L- and T-type Ca2+ channels are expressed in cerebral arterial smooth muscle
and contribute to myogenic tone development and control of localized Ca2+ signaling.
A variety of techniques were employed to test the hypothesis, including molecular, biochemical,
electrophysiological, functional, structural, and computational modeling approaches. The
following two principal objectives were used to guide experiments designed to test this
hypothesis:
1) To identify the different subtypes of voltage-gated Ca2+ channels that are expressed
in cerebral vascular smooth muscle and determine their roles in myogenic tone
development.
2) To determine the functional role of T-type Ca2+ channels, specifically CaV3.2, in
cerebral vascular smooth muscle.
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Chapter Two: Identification of L- and T-type Ca2+ Channels in Rat Cerebral Arteries:
Role in Myogenic Tone Development
Rasha R. Abd El-Rahman, Osama F. Harraz, Suzanne E. Brett, Yana Anfinogenova, Rania E.
Mufti, Daniel Goldman, and Donald G. Welsh
This work was published in Am J Physiol Heart Circ Physiol 304: 58–71, 2013.
I was responsible for the biochemical and functional data collection and analysis, and I wrote
the drafts of the manuscript. Yana Anfinogenova and Osama Harraz carried out the patch clamp
experiments. Daniel Goldman performed the blood flow modeling.
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2.1 INTRODUCTION
Arterial smooth muscle cells actively respond to vasoactive stimuli, altering arterial
diameter and tissue blood flow. Changes in vessel tone are the result of myosin light chain
(MLC20) phosphorylation, a process dynamically controlled by Ca2+ influx from the extracellular
space (Deng et al., 2002). Voltage-gated Ca2+ channels are the principal conductances that
regulate extracellular Ca2+ influx. These membrane channels are hetero-oligomeric complexes
comprised of a pore-forming α1-subunit and accessory proteins that influence gating
characteristics and protein trafficking (Kisilevsky & Zamponi, 2008). The α1-subunit is
composed of four domains, each of which contains six transmembrane segments, an S4 voltage
sensor and a P loop that confers ion selectivity (Jan & Jan, 1990; Talavera et al., 2001).
Molecular studies have identified three classes of α1-subunits (CaV1-3), and within each category
there are several subtypes. CaV1/CaV2 subunits display electrical properties characteristic of high
voltage-activated Ca2+ channels (i.e. L-, P/Q-, N- & R- types; Catterall, 1995). In contrast, CaV3
subunits encode for Ca2+ channels activated by lower voltages (i.e. T-type; Iftinca & Zamponi,
2009; Marcantoni et al., 2008).
CaV1.2 is a key pore-forming subunit of L-type Ca2+ channels in cerebral arterial smooth
muscle. There are at least 3 splice variants and, upon depolarization, its graded activation plays a
central role in setting cytosolic [Ca2+] (Cheng et al., 2009; Liao et al., 2007). L-type Ca2+
channel activation is important for tone development; channel blockade does not eliminate all
arterial responsiveness. Case-in-point is the work of Mufti et al. (Mufti et al., 2010) who
showed that ~20% of the myogenic response was insensitive to diltiazem, an L-type Ca2+
channel inhibitor. Such observations have fostered the view that there are additional Ca2+ influx
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pathways that help set cytosolic [Ca2+] and thus the level of myogenic tone (Dimopoulos et al.,
2007; Walsh et al., 1995). In this context, recent studies have noted that CaV3.1 and CaV3.2, α1-
subunits of T-type Ca2+ channels, are present in resistance arteries (Jensen et al., 2004; Nikitina
et al., 2007). T-type Ca2+ channels activate ~10-15 mV negative to L-type Ca2+ channels and
thus are positioned to regulate Ca2+ influx at more hyperpolarized potentials (Perez-Reyes,
2003). While Ca2+ influx through T-type Ca2+ channels could facilitate myogenic tone
development, experimental findings are limited and incomplete (Navarro-Gonzalez et al., 2009).
The present study determined which voltage-gated Ca2+ channels are expressed in rat
cerebral arteries and what role they play in myogenic tone development. To meet this objective,
we combined the experimental strengths of molecular biology, patch clamp electrophysiology,
vessel myography and computational modelling. Our RT-PCR and western blot analysis revealed
that the α1-subunits of CaV1.2 (L-type) and CaV3.1/3.2 (T-type) were expressed at the mRNA
and protein level in isolated smooth muscle cells and whole cerebral arteries, respectively.
Consistent with these observations, patch clamp electrophysiology revealed the presence of two
distinct Ba2+ currents, with electrical and pharmacological fingerprints characteristic of L- or T-
type Ca2+ channels. Functionally, L-type Ca2+ channel inhibition attenuated myogenic tone
development, particularly at higher pressures where arteries are depolarized. In comparison, the
contribution of T-type Ca2+ channels was more limited and best observed at lower pressure in
hyperpolarized vessels. While their contribution to tone development is limited, computational
modeling indicates that T-type Ca2+ channel activity could possibly alter resting blood flow by
20-50%. In closing, our findings demonstrate that α1-subunits of L- and T-type Ca2+ channels
are both expressed in arteries and that they encode for a distinct voltage-dependent conductance.
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They further reveal that each current contributes, albeit unequally, to the maintenance of
myogenic tone, a key physiological response in the cerebral circulation.
2.2 MATERIALS AND METHODS
2.2.1 Animal procedures.
Animal procedures were approved by the Animal Care and Use Committee at the
University of Calgary. Briefly, female Sprague-Dawley rats (10-12 weeks of age) were
euthanized via carbon dioxide asphyxiation. The brain was carefully removed and placed in cold
phosphate-buffered (pH 7.4) saline solution containing (in mM): 138 NaCl, 3 KCl, 10 Na2HPO4,
2 NaH2PO4, 5 glucose, 0.1 CaCl2, and 0.1 MgSO4. Middle and posterior cerebral arteries were
carefully dissected out of surrounding tissue and cut into 2-3 mm segments.
2.2.2 Vessel myography.
Arterial segments were mounted in a customized arteriograph and superfused with warm
(37 °C) physiological salt solution (PSS; pH 7.4; 21% O2, 5% CO2, balance N2) containing (in
mM): 119 NaCl, 4.7 KCl, 20 NaHCO3, 1.1 KH2PO4, 1.2 MgSO4, 1.6 CaCl2, and 10 glucose (Kuo
et al., 1991; Welsh et al., 2000). To limit the tonic dilatory influence of the endothelium on
myogenic tone development (Knot et al., 1999; Kuo et al., 1991), these cells were removed by
passing air bubbles through the vessel lumen (1 to 2 min); successful removal was confirmed by
the loss of bradykinin-induced dilation. Arteries were equilibrated for 30 min at 15 mmHg and
contractile responsiveness assessed by briefly exposing (∼10 s) arteries to 60 mM KCl. After
equilibration, intravascular pressure was increased incrementally from 15 to 20, or 80 mmHg.
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Vessels were then subjected to one of three experimental protocols: 1) nifedipine (50 and 300
nM) + mibefradil (1 and 5 µM), 2) nifedipine + kurtoxin (1 µM), or 3) nifedipine + efonidipine
(3 µM). Each protocol was followed by the addition of a Ca2+-free physiological salt solution (0
externally added Ca2+ + 2 mM EGTA). In one additional set of experiments, arterial tone was
sequentially measured at 20, 40, 60, 80, and 100 mmHg under control conditions and in the
presence of nifedipine (300 nM) ± mibefradil (5 µM). Arterial diameter was monitored using an
automated edge detection system (IonOptix).
2.2.3 Isolation of arterial smooth muscle cells.
Smooth muscle cells from middle and posterior cerebral arteries were enzymatically
isolated as described previously (Luykenaar et al., 2004; Talavera et al., 2001). Briefly, arterial
segments were placed in an isolation medium (37 °C, 10 min) containing (in mM): 60 NaCl, 80
Na+ glutamate, 5 KCl, 2 MgCl2, 10 glucose, and 10 HEPES with 1 mg/ml albumin (pH 7.4).
Vessels were then exposed to a two-step digestion process that involved: 1) a 15-min incubation
in isolation medium (37 °C) containing 0.6 mg/ml papain and 1.8 mg/ml dithioerythritol and 2) a
15-min incubation in isolation medium containing 100 µM Ca2+, 0.7 mg/ml type F collagenase
and 0.4 mg/ml type H collagenase. After treatment, tissues were washed repeatedly with ice-cold
isolation medium and triturated with a fire-polished pipette. Liberated cells were stored in ice-
cold isolation medium for use the same day.
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2.2.4 PCR analysis.
Smooth muscle cells (~200) were isolated from middle and posterior cerebral arteries.
Individual cells were collected manually with a suction pipette and pooled together in RNase-
and DNase-free collection tubes to extract total RNA (RNeasy mini kit with DNAase treatment;
Qiagen, Valencia, CA). Total RNA was extracted (RNeasy mini kit with DNAase treatment;
Qiagen, Valencia, CA) and first-strand cDNA synthesized using the Sensi-script RT kit (Qiagen)
with oligo d(T) primer. Subsequently, a sample of total volume of 50 µl was prepared as follows:
2 µl of each first-strand cDNA reaction was used as templates in a PCR reaction containing 0.25
µM forward and reverse primers (see below), 0.2 mM deoxynucleoside triphosphates, and 2.5
µM recombinant Taq DNA polymerase. The PCR reaction protocol includes a denaturation step
at 92 °C for 5 min; 35 cycles of 92 °C for 45 s, 55° C for 60 s, and 72 °C for 2 min; and a final
extension step at 72 °C for 5 min. Forward and reverse specific primers (made by the University
of Calgary Core DNA Services) were as follows: CaV1.1 (400 bp), (forward) 5´-
TGCTGATCGTCATGCTCTTC-3´, (reverse) 5´-ATGGCCTTGAACTCATCCAG-3´; CaV1.2
(400 bp), (forward) 5´-GCCTCTTCACGGTGGAG-3´, (reverse) 5´-
TCCCAATCACTGCATAGATAA-3´; CaV1.3 (399 bp), (forward) 5´-
TCCGAAGAGCCTGCATTAGT-3´, (reverse) 5´-GCGGTCTTAACACTCGGAAG-3´;
CaV1.4 (500 bp), (forward) 5´-GCAAAGTGGCTCTTCAGGAC-3´, (reverse)
5´-GTTGAGGGGAACATTCATGG-3´; CaV2.1 (501 bp), (forward)
5´-CTGCTTTGAAGAGGGGACAG-3´, (reverse) 5´-CGAAGAGCTCCATCAAAAGG-
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3´; CaV2.2 (504 bp), (forward) 5´-TGTTCAGCAAGTGGCTTTTG-3´, (reverse) 5´-
TTTCAGGGGGAACAGACAAC-3´; CaV2.3 (502 bp), (forward) 5´-
TCCTGCATGACAACCAACAT-3´, (reverse) 5´-GGTCTCGGAAGTACGATCCA-3´;
CaV3.1 (501 bp), (forward) 5´-TCTTCCAGGACAGGTGGAAC-3´, (reverse) 5´-
TCAGCACGAAGGACACAAAG-3´; CaV3.2 (402 bp), (forward) 5´-
AGTTTCCTCTTTGGGGGCTA-3´, (reverse) 5´-CAGGAAAACCCAAACCTGAA-3´; CaV3.3
(501 bp), (forward) 5´-CCCTGGAGATGATCCTGAAA-3´, (reverse) 5´-
AGTTGCCAAAGGTCATGAGG-3´; β-actin (130 bp), (forward) 5´-
TATGAGGGTTACGCGCTCCC-3´, (reverse) 5´-ACGCTCGGTCAGGATCTTCA-3´.
After electrophoresis on a 1.5% agarose gel, the PCR products were excised, extracted, and
purified with a kit purchased from Qiagen (Valencia, CA). Sequencing was performed at the
University of Calgary Core DNA Services. Smooth muscle cell samples were screened for
template and endothelial cell contamination, as previously described (Wu et al., 2007).
2.2.5 Western blotting.
Cortex or cerebral arterial segments from two rat brains, or tissues from heart, retina, or
kidney arterioles were placed in 100 µl of lysis buffer (pH 7.4) containing (in mM): 150 NaCl, 1
CaCl2, 1 MgCl2, 10 HEPES, 0.5% Tween, and 10% mammalian protease inhibitor cocktail
(Sigma-Aldrich, St. Louis, MO). Samples were mechanically disrupted, exposed to three freeze-
thaw cycles, and then centrifuged (10 min; 13,000 rpm). Supernatant was placed in a clean tube,
assayed for total protein, and stored at -20 °C for up to 1 week. Samples were prepared for
electrophoresis by placing 15 µl of supernatant in 5 µl of 4X sample buffer plus 2 µl of DTT.
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After heating (10 min; 90 °C) was completed, 20 µg of protein were loaded on a 5.6%
polyacrylamide gel. Proteins were transferred to a polyvinylidene difluoride (PVDF) membrane
at 4 °C for 1 h; blotted membranes were initially washed (PBS, 5 min) and then exposed to a
0.5% glutaraldehyde PBS solution (45 min) to cross link and fix proteins. After a second set of
washes in Tris-buffered saline (TBS), the membrane was blocked with 1.0% ECL blocking agent
(GE Healthcare, Buckinghamshire, UK) in TBS containing 0.02% Tween-20 (TBST) for 1 h and
subsequently incubated overnight (4 °C) in a TBST (0.1%) solution containing a primary rabbit
antibody [CaV1.2 (1:200), CaV3.1 (1:200), CaV3.2 (1:200); Alomone Laboratories]. The next
morning, the membrane was washed (TBST, 0.02%), and incubated in a TBST (0.1%) solution
containing horseradish peroxidase-conjugated secondary antibody (HRP anti-rabbit antibody)
(1:10,000 dilution, Jackson Laboratories). The membrane was washed for a final time in TBST
(0.02%; 1 h, 20-22 °C) and HRP detected with the Amersham ECL Advance Western Blotting
Detection Kit (GE HealthCare). Emitted light was detected with a chemiluminescence image
analyzer and analyzed with Multi Gauge v3.0 software (Fujifilm, ON, CA).
2.2.6 Electrophysiology.
Conventional patch-clamp electrophysiology was used to measure whole-cell currents in
isolated smooth muscle cells. Briefly, recording electrodes (pipette resistance, 4–7 MΩ) were
fashioned from borosilicate glass and back-filled with pipette solution containing (in mM): 135
CsCl, 2.5 Mg-ATP, 10 HEPES, and 10 EGTA (pH 7.2). This pipette was then gently lowered
onto the cell and negative pressure applied to rupture the membrane. With series resistance ˂10
MΩ and input resistance >10 GΩ, cells were then voltage clamped (-60 mV) and equilibrated for
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5 min in a bath solution containing (in mM): 110 NaCl, 135 CsCl, 4 TEA-Cl, 1.2 MgCl2, 10
BaCl2, 10 HEPES, and 10 glucose (pH 7.4). The voltage protocol consisted of a prepulse to -90
or -60 mV (200 ms), followed by a series of voltage steps from -80 mV to +40 mV (200–300 ms,
10-mV increments). This protocol was run under control conditions and in the presence of
nifedipine (300 nM) ± mibefradil (5 µM), kurtoxin (5 µM), or efonidipine (3 µM). Whole-cell
currents were recorded on an Axopatch 200B amplifier (Axon Instruments), filtered at 1 kHz,
digitized at 5 kHz, and stored on a computer for subsequent analysis with Clampfit 10.2
software. Cell capacitance ranged between 14 and 18 pF and was measured with the cancellation
circuitry in the voltage-clamp amplifier. A 1M NaCl-agar salt bridge between the reference
electrode and the bath solution was used to minimize offset potentials (< 2 mV). All experiments
were performed at room temperature (20 –22 °C).
2.2.7 Computational blood flow modeling.
A previously developed theoretical model (Goldman & Popel, 2000; Pries et al., 1990;
Pries et al., 1994) was used to calculate two-phase [red blood cell (RBC) and plasma] steady-
state flow in the arteriolar networks considered. This model implements conservation of blood
and RBC volume flow at each node joining three arterial branches and includes known blood
rheology, specifically the Fahraeus and Fahraeus-Lindqvist effects in unbranched segments and
phase separation at diverging bifurcations. Besides geometric information, the model requires
specification of boundary conditions in the form of hematocrit at the inlet segment (upstream end
of the 1st-order posterior cerebral artery) and pressure changes between the inlet node and outlet
nodes (downstream ends of the 3rd-order posterior arteries). All flow simulations used an inlet
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hematocrit of 0.4, and inlet and outlet pressures of 75 mmHg and 50 mmHg, respectively.
Arterial diameter/length values were set according to functional measurements from posterior
cerebral vessels pressurized to 60 mmHg. Values were as follows: 1st order: 164 µm diameter,
3.6 mm length; 2nd order: 150 µm diameter, 3.5 mm length; and 3rd order: 125 µm diameter, 1.1
mm length.
2.2.8 Statistical analysis.
Data are expressed as means ± SE, and n indicates the number of vessels or cells. No
more than two experiments were performed on vessels from a given animal. Where appropriate,
paired, unpaired t-tests, and one way ANOVA were performed to compare the effects of a given
condition/treatment on arterial diameter, or whole cell current (see figure legends for specific
details). P values ≤ 0.05 were considered statistically significant.
2.2.9 Solutions and chemicals.
All buffers, chemicals, and reagents were purchased from Sigma-Aldrich unless
otherwise stated.
2.3 RESULTS
2.3.1 Ca2+ channel α1-subunit expression in middle and posterior cerebral arteries.
This study began with an mRNA characterization of CaV α1-subunit expression in whole
cerebral arteries and isolated smooth muscle cells. Primers were designed against rat sequences
and efficacy confirmed by screening a range of appropriate control tissues. Skeletal muscle was
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used as the control for CaV1.1 and brain tissue was used for CaV1.2 and CaV1.3. Retina was the
control tissue for CaV1.4. Brain tissue was used as control for all CaV2.x channels. Afferent
arterioles from kidney were used as control for CaV3.1 and CaV3.2. Cardiac tissue was the
control for CaV3.3 channel. All α1-subunit products were sequenced to confirm their identities.
With the use of whole cerebral arteries, this study confirmed that α1-subunits for high (CaV1.1,
CaV1.2, CaV1.3, and CaV2.2) and low (CaV3.1, CaV3.2, and CaV3.3) voltage-activated Ca2+
channels were observable at the mRNA level (Fig. 4B). This expression profile was reduced to
CaV1.2, CaV3.1, and CaV3.2 in isolated smooth muscle cells (Fig. 4C). Note that isolated smooth
muscle cells were pre-screened for endothelial contamination by using primers for ET-1; no
preps that screened positive for ET-1 were used. With the use of the mRNA observations as a
guide, western blot analysis was subsequently performed to assess protein expression. With the
use of brain as the positive control, this study confirmed that the α1-subunits of L-type (CaV1.2)
and T-type (CaV3.1 and CaV3.2) Ca2+ channels were present at the protein level in whole cerebral
arteries (Fig. 5).
2.3.2 Whole-cell Ba2+ currents in cerebral arterial smooth muscle cells.
Building on the preceding observations, we next ascertained whether whole-cell patch
clamp electrophysiology could delineate L- and T-type Ca2+ channels from one another.
Cerebral arterial smooth muscle cells were enzymatically isolated, and whole-cell currents were
monitored in the presence of 10 mM Ba2+ to accentuate charge flow through L-type Ca2+
channels. Stepping from a prepulse of -90 mV, voltage protocols first revealed an inward Ba2+
current that activated at ~ -30 mV and peaked at +20 mV (Fig. 6), however, when the prepulse
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was changed to -60 mV to facilitate T-type Ca2+ channel inactivation, no statistically significant
change in current was detected at any voltage. Further experiments showed that nifedipine,
applied at concentrations approximately five- to 30-fold higher than the IC50 value for the
smooth muscle variant of CaV1.2 (Jensen & Holstein-Rathlou, 2009), blocked ~75% of the
whole-cell Ba2+ current (Fig. 7, A and B). Plots in Fig. 7C indicate that the current-voltage
relationship of the nifedipine-insensitive current is leftward shifted in relation to the nifedipine-
sensitive component when 50 nM or 300 nM nifedipine was used. Note that a nifedipine
concentration of 300 nM is predicted, based on previous work, to block >98% of all L-type Ca2+
channels in this vascular preparation (Morel et al., 1998). Higher micromolar concentrations
were avoided since they interfere with CaV3.1 and CaV3.2 channels (Akaike et al., 1989; Lee et
al., 2006). Further work was performed to determine whether the nifedipine-insensitive current
reflected T-type Ca2+ channel activation. The experimental approach consisted of two elements.
The first centered on whether the nifedipine-insensitive component was blocked by T-type Ca2+
channel inhibitors including mibefradil, efonidipine, and kurtoxin. These agents were applied at
concentrations 10- to 20-fold above the IC50 value for CaV3.1 and CaV3.2 and should block >
95% of all T-type Ca2+ channels (Furukawa et al., 2004; Horiba et al., 2008; Martin et al., 2000;
Morita et al., 2002). The second involved an electrical analysis in which the time constants of
activation/inactivation (τactivation, τinactivation) were calculated for the nifedipine- and the T-type
Ca2+ channel inhibitor-sensitive components. Whole cell currents were measured using a
prepulse of -90 mV at a series of voltages from -40 mV to +40 mV. Currents were measured for
control and in the presence of 300 nM nifedipine. Currents were then measured after 5 µM
mibefradil was added to the bath. The nifedipine-sensitive current at each voltage was
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determined by subtracting the current in the presence of nifedipine from that measured for
control. The mibefradil-sensitive component at each voltage was determined by subtraction of
the current recorded in the presence of both nifedipine and mibefradil from that recorded in the
presence of nifedipine. Figure 8, A and B, indicate that the nifedipine-insensitive component of
the whole-cell Ba2+ current was largely blocked by 5 µM mibefradil, a T-type Ca2+ channel
blocker. The time constants for activation/inactivation, calculated at +10 to +20 mV, were lower
for the mibefradil-sensitive component than the nifedipine-sensitive component, indicating faster
activation and inactivation. This finding is consistent with mibefradil blocking activated T-type
Ca2+ channels (Fig. 8C).
Similar experiments were carried out using two other T-type channel blockers: kurtoxin
and efonidipine. Whole cell currents were measured using a prepulse of -90 mV at a series of
voltages from -40 mV to +40 mV. Currents were measured for control and in the presence of 300
nM nifedipine. Currents were then measured after 5 µM kurtoxin (Fig. 9, A and B) or 3 µM
efonidipine (Fig. 10, A and B) was added to the bath. The nifedipine-sensitive current at each
voltage was determined by subtracting the current in the presence of nifedipine from that
measured for control. The kurtoxin/efonidipine-sensitive components at each voltage were
determined by subtraction of the current recorded in the presence of both nifedipine and kurtoxin
or efonidipine from that recorded in the presence of nifedipine. The activation/inactivation time
constants were also smaller for the kurtoxin/efonidipine-sensitive than the nifedipine-sensitive
component of the whole-cell Ba2+ current (Figures 9C and 10C). Reverse-order experiments (i.e.
addition of the T-type channel blockers before nifedipine) were not performed since these T-type
Ca2+ channel inhibitors will block L-type Ca2+ channels when applied at micromolar
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concentrations (Jensen & Holstein-Rathlou, 2009). Note, in these longer recordings, a small,
linear current characteristic of leak was at times evident after the addition of T-type Ca2+ channel
blockers.
2.3.3 Ca2+ channel blockers and myogenic tone.
Having determined that L- and T-type Ca2+ channels are electrically present in cerebral
arterial smooth muscle cells, we next determined their role in myogenic tone development. The
approach consisted of the sequential application of L- and then T-type Ca2+ channel blockers to
middle/posterior cerebral arteries pressurized to 20 or 80 mmHg. At 20 mmHg, arterial
membrane potential will rest near -55 mV, whereas at 80 mmHg, tissues will depolarize to -35
mV (Lacinova, 2005; Smirnov & Aaronson, 1992). At these different voltages, Ca2+ influx
through L- and T-type Ca2+ channels should vary along with their relative contributions to the
myogenic response. At 20 mmHg, cerebral arteries displayed 43 ± 4 µm of tone (Fig. 11, A and
C). Change in diameter was determined by subtraction of the diameter measured for control (no
blockers presents) from the diameter measured in Ca2+ free solution. The addition of nifedipine,
50 nM and 300 nM, to the superfusate modestly attenuated tone development, as did 1 and 5 µM
mibefradil, a T-type Ca2+ channel blocker (see Fig. 11A). Pressurizing middle or posterior
cerebral arteries to 80 mmHg augmented resting tone to 104 ± 6 µm (Fig. 11, B and D). In this
depolarized and constricted state, nifedipine application had a disproportionately greater effect
on myogenic tone development than did mibefradil. Nifedipine-sensitive change in diameter (Δ
Diameter in Fig. 11E) was calculated as the vessel diameter measured in the presence of 300 nM
nifedipine minus the value measured for control (absence of any channel blockers). The
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mibefradil-sensitive change in diameter was calculated as the diameter recorded in the presence
of both 300 nM nifedipine and 5 µM mibefradil minus the value recorded in the presence of 300
nM nifedipine. These values were determined from measurements made with arterial pressures
of 20 mmHg and 80 mmHg, see Fig. 11E. The nifedipine-sensitive and mibefradil-sensitive
changes in diameter were also expressed as percentages of the maximal change in diameter for
the artery. Maximal change was calculated as the difference between artery diameters recorded
for control (no blockers present) and in Ca2+ free buffer. These results are shown in Fig. 11E,
which indicates, by using both absolute and relative representations, that L-type Ca2+ channel
activation plays an increasing role in tone development with pressurization. In contrast, the T-
type Ca2+ channel component did not substantively change in absolute terms with vessel
pressurization; it did, however, significantly decrease when expressed relative to the maximal
response (see Fig. 11E). A similar pattern was observed when the preceding experiment was
repeated with kurtoxin (Fig. 12) and efonidipine (Fig. 13), two structurally dissimilar T-type
Ca2+ channel blockers (Furukawa et al., 2004; Horiba et al., 2008). In these experiments
nifedipine-sensitive, kurtoxin-sensitive and efonidipine-sensitive changes in diameter were also
calculated as described above for the experiments in which mibefradil was used. These values
were also expressed as percentages of the maximum vessel response as described above.
The relative importance of L- and T-type Ca2+ channels was better revealed in Figure 14
where myogenic tone was monitored over a more complete pressure range (20–100 mmHg)
before and after the sequential addition of 300 nM nifedipine and 5 µM mibefradil. As above, the
functional contribution of L-type Ca2+ channels to arterial tone increased with pressurization,
whereas that of T-type Ca2+ channels decreased. Although T-type Ca2+ channel blockade elicits
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only limited vasomotor responses, these alterations are likely sufficient to influence blood flow,
as evident from blood flow modeling (Fig. 15). This computational work was used to predict
how changes in vessel diameter could affect blood flow. This work revealed that a 5–15%
change in resting diameter (comparable to that induced by T-type Ca2+ channel blockade at 60
mmHg) could elicited a 20–50% change in network perfusion. With these simulations, we
assume that T-type Ca2+ channel expression is constant among the three vessel orders.
2.4 DISCUSSION
In this study of the rat cerebral circulation, we determined which voltage-gated Ca2+
channels are expressed and whether they are important to myogenic tone development. In this
regard, we used 1) molecular/biochemical approaches to determine which α1-subunits were
present, 2) patch clamp electrophysiology to characterize Ca2+ channel properties, and 3) vessel
myography to ascertain function. An mRNA assessment revealed that the α1-subunits of L-type
(CaV1.2) and T-type (CaV3.1 and CaV3.2) Ca2+ channels were present in isolated rat cerebral
arterial smooth muscle cells. Western analysis subsequently confirmed protein expression in
intact cerebral arteries. Whole-cell electrophysiology demonstrated that an inward Ba2+ current
was readily observable and divisible into a nifedipine-sensitive (L-type) and a nifedipine-
insensitive component. The latter was inhibited by mibefradil, kurtoxin, and efonidipine and
displayed activation/inactivation properties consistent with T-type Ca2+ channels. Functional
experiments further revealed that L-type Ca2+ channels are principally responsible for driving
myogenic tone, although a limited role for T-type Ca2+ channels is suggested in hyperpolarized
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vessels. Blood flow modeling indicates that T-type Ca2+ channel modulation could alter blood
flow through the posterior cerebral circulation by ~ 20–50%.
2.4.1 Background.
Voltage-gated Ca2+ channels are expressed in a diverse range of excitable and
nonexcitable tissues (Catterall et al., 2005). Structurally, the pore-forming α1-subunit comprises
four membrane-spanning domains (I-IV) linked together in a single polypeptide chain. Each
domain contains six transmembrane segments plus a P loop that dips incompletely into the pore
to confer selectivity (Dolphin, 2006; Hofmann et al., 1999). Molecularly, voltage-gated Ca2+
channels are categorized according to α1-subunit expression, and there are, at present, three main
classifications including CaV1.x (L-type), CaV2.x (P/Q-, N-, and R types), and CaV3.x (T-type).
CaV1.x and CaV2.x encode for Ca2+ channels that are more active at depolarized voltages
compared with the low voltage-activated CaV3.x subunits (Ball et al., 2009; Hansen et al., 2011).
In vascular smooth muscle, CaV1.2 principally controls steady-state Ca2+ entry and, therefore,
myosin light chain phosphorylation and tone development (Harder, 1984; Knot & Nelson, 1998).
Although important to tone development, recent studies have suggested that this α1-subunit is not
singularly expressed in arterial smooth muscle (Braunstein et al., 2009; Kuo et al., 2010;
Nikitina et al., 2007; Pluteanu & Cribbs, 2011). For example, immunohistochemical
observations have suggested that α1-subunits of T-type Ca2+ channels are present in the renal,
mesenteric, and cerebral circulation (Braunstein et al., 2009; Feng et al., 2004; Navarro-
Gonzalez et al., 2009). Supporting electrophysiological findings are sparse, although selected
studies have shown that a nifedipine-insensitive Ba2+ current with properties reminiscent of T-
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type Ca2+ channels can be isolated in arterial smooth muscle (rat and dog basilar arteries)
(Navarro-Gonzalez et al., 2009; Nikitina et al., 2007). At present, it is not clear what role T-type
Ca2+ channels play in arterial tone development. Past functional findings have been decidedly
mixed with only a small number indicating a limited to prominent contribution, depending on the
vascular bed and stimulus (Braunstein et al., 2009; Hansen et al., 2011; Kuo et al., 2010; Poulsen
et al., 2011). Others, however, have argued that T-type Ca2+ channels have no role in the
contractile process and that their principal function centers on cell cycle regulation and
proliferation (Oguri et al., 2010; Perez-Reyes, 2003; Rodman et al., 2005).
2.4.2 Molecular and electrical characterization.
This study began by examining which voltage-gated Ca2+ channels are present in cerebral
arterial smooth muscle. Work was initiated at a molecular level with a PCR analysis of whole
arteries and isolated smooth muscle cells from rat middle/posterior cerebral arteries. Consistent
with past findings (Braunstein et al., 2009), multiple α1-subunits were observed in whole arteries
including CaV1.1-1.3, CaV2.2, and CaV3.1–3.3 (Fig. 4). The breadth of the mRNA profile was
expected since whole arteries comprise three major cell types (smooth muscle, endothelium, and
perivascular nerves), all of which can express CaV α1-subunits (Cribbs, 2001). Limiting mRNA
analysis to isolated smooth muscle cells reduced the expression pattern to CaV1.2 (L-type) and
CaV3.1/3.2 (T-type). Western blot analysis subsequently confirmed that each α1-subunit was
expressed at the protein level in cerebral arteries (Fig. 5). Although this study did not pursue an
immunohistochemical analysis, Kuo et al. have shown that all three α1-subunits are indeed
present in rat cerebral arterial smooth muscle (Kuo et al., 2010). With molecular/biochemical
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evidence of L- and T-type Ca2+ channels expressed in cerebral arterial smooth muscle we next
used patch clamp electrophysiology to functionally isolate and identify the two general
conductances. Like past studies, experimentation began by monitoring Ba2+ currents in cells held
at -90 or -60 mV, the concept being that the hyperpolarized holding potential would relieve
voltage-dependent inactivation of T-type Ca2+ channels and thus augment whole-cell current
(Navarro-Gonzalez et al., 2009; Nikitina et al., 2007). We did not observe a significant change in
current. Therefore, we next monitored inward Ba2+ currents in the presence and absence of
nifedipine to determine whether a dihydropyridine-insensitive current with T-type Ca2+ channel
properties could be functionally isolated (Fig. 7). Nifedipine blocks L-type Ca2+ channels in a
state-dependent manner by binding to inactivated pores (Dolphin, 2006; Liao et al., 2007).
Concentrations were set 5- to 30-fold above the IC50 for the smooth muscle splice variants of
CaV1.2 (Jensen & Holstein-Rathlou, 2009) and below levels that interfere with T-type Ca2+
channels (Akaike et al., 1989; Lee et al., 2006). At levels that should block >98% of all L-type
Ca2+ channels (300 nM), whole-cell inward current decreased by ~75%, a finding in agreement
with Nikitina et al. (Nikitina et al., 2007) in dog basilar artery and Kuo et al. (Kuo et al., 2010) in
the rat cerebral circulation. The residual nifedipine-insensitive current displayed a leftward
shifted current-voltage relationship characteristic of T-type Ca2+ channels (Iftinca & Zamponi,
2009; Perez-Reyes, 2003). A more detailed examination of the nifedipine-insensitive current
followed with experiments monitoring the effects of T-type Ca2+ channel blockers and
characterizing key electrical properties. In regards to the former (the effects of the blockers),
Figs. 8–10 show that mibefradil, efonidipine, and kurtoxin effectively blocked much of the
nifedipine-insensitive Ba2+ current. The analysis of mibefradil/efonidipine/kurtoxin-sensitive
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currents revealed that the activation/inactivation time constants were smaller than those
determined for the nifedipine-sensitive current and corresponded well with published values for
CaV3.1/3.2 and CaV1.2, respectively (Kuo et al., 2010; Perez-Reyes, 2003). Overall, the
preceding patch clamp observations indicate L- and T-type Ca2+ channels are not only
electrically present in cerebral arterial smooth muscle but distinguishable from one another.
2.4.3 T-type Ca2+ channels and myogenic tone.
When current through T-type Ca2+ channels is recorded in solutions containing
physiological concentrations of Ca2+, the channels display a V0.5 for steady-state
activation/inactivation that is 10-15 mV negative to CaV1.2 (Nikitina et al., 2007; Perez-Reyes,
2003). As such, there will be a subsequent leftward shift in the window current (Kuo et al., 2010;
Navarro-Gonzalez et al., 2009; Nikitina et al., 2007; Perez-Reyes, 2003). From this knowledge,
it is logical to predict that the contribution of T-type Ca2+ channels to tone development should
be more prominent at hyperpolarized potentials. To test this hypothesis, this study examined
myogenic tone development and the effect of sequential addition of L- and then T-type Ca2+
channel blockers on cerebral arteries pressurized to 20 or 80 mmHg. It was theorized that at 20
mmHg, vessels would reside in a hyperpolarized state (~-55 mV) and that T-type Ca2+ channel
activity would be greater given their activation/inactivation properties (Perez-Reyes, 2003). In
contrast, at 80 mmHg, cerebral arteries would be more depolarized (~-35 mV) and L-type Ca2+
channel activity would be predicted to dominate to a greater extent over T-type (Bannister et al.,
2009; Knot & Nelson, 1995). Findings in Figs. 11–13 clearly illustrate that T-type Ca2+ channel
blockers, such as mibefradil, kurtoxin and efonidipine, do impair the nifedipine-insensitive
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component of the myogenic response. When expressed on a relative scale, the contribution of T-
type Ca2+ channels was greatest at low intravascular pressures where the vessel is hyperpolarized
(-55 mV) (Kuo et al., 2010; Nikitina et al., 2007; Perez-Reyes, 2003). As expected, the
functional significance of L-type Ca2+ channels increased with pressurization, a finding
consistent with the literature and the electrophysiological properties of high voltage-activated
Ca2+ channels (Bannister et al., 2009; Knot & Nelson, 1998). The unequal contribution of L- and
T-type Ca2+ channels to myogenic tone development is better observed in Fig. 14 when cerebral
arteries were stepped sequentially from 20 to 100 mmHg. Although previous studies have
attempted similar measurements (Hansen et al., 2011; Kuo et al., 2010; Nikitina et al., 2007),
this study is the first to show clear evidence that T-type Ca2+ channels contribute to myogenic
tone development in a manner consistent with their voltage-dependent properties (Perez-Reyes,
2003).
In regard to the preceding findings and their interpretation, this study implicitly assumed
that T-type Ca2+ channels operate as a general influx pathway, providing a portion of the
extracellular Ca2+ required for contractile activation. Although reasoned, it is plausible that T-
type Ca2+ channels might also facilitate arterial tone through alternative mechanisms. For
example, Ca2+ flux via CaV3.1 or CaV3.2 channels might be sufficient to activate a Ca2+-
dependent inward current such as TRPM4, a transient receptor potential channel that contributes
to pressure-induced depolarization (Earley, 2010). They might also initiate and/or maintain Ca2+
waves, asynchronous events dependent on Ca2+ release from the SR (Braunstein et al., 2009;
Martin-Cano et al., 2009). Like T-type Ca2+ channels, these events are more important in setting
myogenic tone at hyperpolarized rather than depolarized voltages (Mufti et al., 2010).
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2.4.4 Implications to blood flow control.
The observations in Figures 11–14 suggest that T-type Ca2+channels contribute to
myogenic tone development in the cerebral circulation. Although their overall role is limited, it
would be imprudent to overlook their impact on tissue perfusion since flow is proportionally
related to blood pressure and vessel radius raised to the 4th power. To better illustrate the impact
of T-type Ca2+ channels on tissue perfusion, blood flow simulations were performed on a
network structure that mimicked the proximal aspects of the posterior cerebral circulation (Fig.
15). This network structure consisted of seven segments whose diameter and length were set
according to measurements on 1st- to 3rd-order posterior cerebral arteries pressurized to 60
mmHg. Blood and RBC volume flow were conserved, whereas hematocrits along with inlet and
outlet pressure were set to 0.4, 75, and 50 mmHg, respectively. With this model, we observed
that 5–15% changes in resting arterial diameter, generally akin to dilatory responses induced by
T-type Ca2+ channel inhibition at 60 mmHg, augmented network perfusion by ~20–50%. From
such findings, we deduce that T-type Ca2+ channel modulation could be functionally important
and have a measurable impact on tissue perfusion.
2.4.5 Summary.
This study demonstrated that multiple voltage-gated Ca2+ channels are present in the rat
cerebral circulation. More specifically, molecular and electrophysiological approaches indicated
that both L-type (CaV1.2) and T-type (CaV3.1 and CaV3.2) Ca2+ channels are expressed and
electrically discernible in smooth muscle cells from middle/posterior cerebral arteries. Functional
analysis indicated that T-type Ca2+ channels contribute to the genesis of the myogenic response.
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Their functional significance would be more evident at low intravascular pressure where the
hyperpolarized state of the vessel would optimize steady-state Ca2+ influx, given the voltage-
dependent properties of the channel (Perez-Reyes, 2003). Although the overall contribution of T-
type Ca2+ channels to tone control is modest, blood flow modeling indicates that T-type Ca2+
channel modulation could alter resting blood flow by as much as 50%.
2.4.6 Limitations and future directions.
This study provided evidence for the presence of CaV1.2 (L-type) and CaV3.1 and CaV3.2
(T-type) voltage-gated Ca2+ channels in rat cerebral arteries and the results presented also
suggest functional roles for CaV3.1 and CaV3.2 T-type VGCCs in myogenic tone development.
However, there are some experimental limitations and additional experiments that should be
considered.
(i) Molecular and biochemical approach.
The semi-quantitative reverse transcription PCR technique was performed on isolated
smooth muscle cells to scan for mRNA message of VGCCs. In these experiments posterior and
middle cerebral arteries were isolated as described previously (Luykenaar et al., 2004; Talavera
et al., 2001). The tissues were treated enzymatically, then washed repeatedly with ice-cold
isolation medium and triturated with a fire-polished pipette to liberate the cells. Individual cells
were collected manually with a suction pipette and pooled together in RNase- and DNase-free
collection tubes to extract total RNA (RNeasy mini kit with DNAase treatment; Qiagen,
Valencia, CA). First-strand cDNA was synthesized using the Sensi-script RT kit (Qiagen). The
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results showed the presence of the α1–subunit of L-type (CaV1.2) and T-type (CaV3.1 and
CaV3.2) channels in isolated cerebral arterial smooth muscle cells.
Although the preparations were pre-screened for endothelial contamination (ET-1), a
screening for neuronal contamination in the preparations was not done. As the arteries are
comprised of three cell types (smooth muscle, endothelial, perivascular nerves), any
contamination from perivascular nerves could cause misleading results regarding the expression
of VGCCs in smooth muscle; this is especially true for T-type VGCCs, which are abundantly
expressed in the neuronal tissues.
One way to screen the preparations for neuronal cell contamination would be to use semi-
quantitative reverse transcription PCR with primers that will allow for the detection of DNA that
codes for neuronal cell specific marker protein (e.g. neurofilament (NF), subtypes of NF, NF-L,
NF-M, NF-H, or α-internexin). A second way to screen the preparations for a neuronal cell
contamination would be to use real-time reverse transcription PCR, which quantitates differences
in mRNA expression, to run two parallel reactions that would amplify a smooth muscle specific
marker (e.g. smooth muscle myosin) and a neuronal marker (e.g. neurofilament) by using SYBR
green core reagents (Applied Biosystems, Carlsbad, CA, USA). SYBR green is a dye that binds
to double stranded DNA but not to single-stranded DNA and it fluoresces very brightly, and can
be used to detect the amplified DNA. A third method to check for neuronal contaminations
would also use reverse transcription PCR. Forward and reverse primers for a smooth muscle
specific marker (e.g. smooth muscle myosin) and a for a neurofilament marker (e.g. NF or any
subtype of NF) would be added to a single sample and used to amplify DNA. SYBR green would
be used to label all amplified DNA. A melt curve would then be generated to indicate the
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presence of one or two components. If only one component was found it would presumably be
from smooth muscle and would indicate the sample had no neuronal contamination. If two
components were found in the melt curve that would indicate the sample is contaminated by
neurons. If the results of the screening show there is no neuronal contamination, then our results
indicate the presence of CaV3.1 and CaV3.2 RNA in smooth muscle cells, which would be in
agreement with the results of Kuo and Navarro-Gonzales et al. who applied real-time PCR, and
detected the presence of the two subtypes (CaV3.1 and CaV3.2) of T-type VGCCs in rat cerebral
arteries (Kuo et al., 2010; Navarro-Gonzalez et al., 2009).
A test could also be done to check if the primers used in PCR reactions for CaV3.1 and
CaV3.2 resulted in amplification of any other types of DNA (e.g. splice variants or DNA from
another type of channel). In this test PCR reaction could be run using the primers for one of the
channels and SYBR green could be used to label all of the amplified DNA. A melt curve could
be generated and would indicate whether one or more components were amplified; additional
components could be splice variants for example for the target channels.
Western blot analysis was performed with whole cerebral arteries. However, western blot
analysis could also be carried out on samples prepared from isolated smooth muscle cells. This
would involve preparing a suspension of isolated cells from cerebral artery segments, collecting
individual smooth muscle cells from the cell suspension using a suction pipette and then pooling
the collected cells from multiple preparations as was done for the PCR analysis.
The previous approaches (biochemical, molecular) have indicated the presence of
different types and subtypes of VGCCs in rat cerebral arteries. Immunofluorescence microscopy
involving labelling of isolated smooth muscle cells with antibodies to the channels would be
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another approach that could be used to further investigate the presence of both L-and T-type
VGCCs in these cells. Experiments of this type are described in Chapter 3, section 3.4.4.
Primary antibodies against selected α1-subunits of L- and T-type VGCCs (CaV1.2,
CaV3.1, and CaV3.2) were used and the results indicated the presence of these channels in rat
cerebral arteries; however, the specificity of the primary antibodies could be better tested to
make sure there is not any non-specific binding for the antibodies used. There are a number of
ways that could be used to look at specificity of the primary antibodies. 1) The labelling of the
whole blot can be examined to look for extraneous bands (at molecular weights different than
that expected for the channels) that might indicate non-specific binding. 2) Competition assays
using the peptide antigens for the primary antibodies can also be used to examine specificity of
the antibodies. 3) Primary antibodies from different sources could be compared and used to
determine which has the highest specificity. 4) A small interference RNA (siRNA) approach
could be used to selectively reduce the expression of the protein that is targeted by a particular
antibody. siRNA interference is a methodology whereby small segments of double-stranded
RNA are introduced into isolated cerebral arteries using reversible permeabilization in organ
culture media, to degrade the mRNA required for protein translation. The degradation is
controlled by an endogenous RNA-induced silencing complex, DICER and RISC (Hamilton and
Baulcombe, 1999). Western blot analysis of protein from tissue treated with siRNA should show
less labeling of the target protein band(s) if the primary antibody is specific for the protein.
Small interfering RNA (siRNA) is a small segment of double stranded RNA that binds to
and degrades the mRNA essential for protein translation. Therefore, it greatly reduces or
eliminates the expression of the targeted protein, and induces a type of protein knockout.
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However, the ability to deliver siRNA to cells is limited as many mammalian cell types are
resistant to the transfection methods needed to introduce the synthetic siRNA into them. The
technique requires organ culture for approximately about 3 days. This could expose the vessels
to contamination or tissue damage. Furthermore, the siRNA technique can also be non-specific;
off-target effects of siRNA can also occur if it targets mRNA that is not the intended one.
Application of the siRNA technique is also expensive for large experiments and it mediates
short-term knockdown, so it is not useful for long term studies.
As part of applying the siRNA technique, control tissues have to be tested to confirm the
validity of the technique itself, as both the controls and the treated siRNA tissues are subject to
organ culture for about 3 days. One control tissue would be non-treated (i.e. have no siRNA) and
the second control tissue would be treated with a scrambled RNA sequence that would not affect
the expression of the target protein. If western blot analysis of samples from the control and
siRNA treated tissues do not show changes in the bands labeled by the primary antibody being
tested then that would indicate that those bands are due to non-specific labelling by the antibody.
(ii) Patch-clamp electrophysiological approach.
- Patch-clamp measurements using Ba2+
Conventional patch clamp electrophysiology results demonstrated the presence of two
inward VGCC channel currents (nifedipine-sensitive and nifedipine-insensitive). In cerebral
arterial smooth muscle cells, Ba2+ (10 mM) was used as the charge carrier because the current
carried by Ba2+ is larger than the current carried by Ca2+ therefore it is easier to measure and
differentiate between L- and T-type channels (Perez-Reyes, 2003, Harraz & Welsh, 2013).
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Although using Ba2+ makes it easier to measure currents through VGCCs, Ba2+ also causes the L-
type channels to lose their Ca2+-dependent inactivation properties (Nikitina et al., 2007) and the
high concentration of Ba2+ used leads to a shift in the current-voltage (IV) curve toward more
hyperpolarized values by ~10 mV. This shift has to be considered when comparing channel types
and their properties, as it could give a false indication about the identity of the channels being
investigated (Abd El-Rahman et al., 2013; Kuo et al., 2010). For example it would make it hard
to distinguish between different types of VGCCs and splice variants of VGCCs (Abd El-Rahman
et al., 2013; Kuo et al., 2010). One way to address these possible problems would be to use Ca2+
as the charge carrier (1.8 mM, as higher concentration will cause the cells to contract). Although
the current amplitude will be approximately three times less than the current amplitude when
Ba2+ is used (Kuo et al., 2010) this method has been used to record L- and T-type currents
(Harraz and Welsh, 2013) but it was not possible to differentiate between different types of
channels because of the small current.
- Protocol with different holding potentials.
Figure 6, shows results of experiments in which Ba2+ currents were measured at different
voltages from starting prepulses of -90 mV or -60 mV. The prepulse of -60 mV was used to
maximize inactivation of T-type channels. In these experiments we did not find statistically
significant differences in the currents recorded at any voltage when the two prepulses were
compared. Therefore these results could not be used as evidence of the presence of two different
inward currents. These results could be further investigated by repeating the experiments to
increase the n numbers for the two different prepulses. We did notice a possible small (but not
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significant) decrease in currents when -60 mV was used as a prepulse. These experiments could
be redone by repeating the two protocols several times in a row and comparing results sets (i.e.
of 1st set of traces to 2nd ; 2nd set of traces to 3rd etc…) to eliminate the possibility that the
currents change over time (due to run-down for example). Control experiments in which a series
of traces are compared all at the same prepulse (i.e. at -90 mV prepulse or -60 mV prepulse)
could also be done and analyzed the same way to determine if the rundown of the currents with
time affected the results.
- Use of T-type channel blockers.
One difficulty with the use of pharmacological blockers in the present study is that the
available blockers are not as specific as one would like. For example mibefradil shows high
affinity for blocking T-type Ca2+ channels if Ca2+ is used as the charge carrier but competes with
Ba2+ more than Ca2+ for permeation through the pore, thus both concentration and divalent cation
type affect the efficacy of the drug (Morita et al., 2002; Martin et al., 2000; Lee et al., 1999;
Perez-Reyes, 2003). These results should be taken into consideration when investigating the
presence of different VGCCs in arterial smooth muscle, because it could affect the interpretation
of the data. For example it might be difficult to differentiate between splice variants of a
particular type of a channel and the different types of channels. Another way to test for blockers
specificity would be to measure the currents that remain in the presence of one T-type channel
blocker and then measure it again after the addition of a 2nd T-type blocker along with the first
one (different combinations of the blockers could be used e.g. mibefradil then kurtoxin,
mibefradil then efonidipine, kurtoxin then efonidipine, or vice versa,…etc). If the second blocker
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causes additional effects on the channel that might be an indication that additional channels types
are affected; if the second blocker addition causes no further change in the remaining current that
would strengthen the conclusion that the same channels are being affected by the channel
blockers.
Also, the siRNA approach discussed above could be employed to address the problem of
using non-specific blockers. For these experiments Ca2+ currents from cells isolated from
untreated or tissue treated with scrambled siRNA could be compared with currents from cells
treated with siRNA to knock down CaV3.1 and CaV3.2 T-type channels. The effects of L- and T-
type channel blockers could also be compared in these cells; siRNA treatment should eliminate
effects of the T-type channel blockers if they are specific.
(iii) Vessel myography approach.
The experimental scheme for the vessel myography relied mainly on the use of different
pharmacological blockers (mibefradil, kurtoxin, efonidipine) that block T-type VGCCs (CaV3.1
and CaV3.2) when used at a specific range of concentrations (see Table 1; Lee et al., 1999;
Perez-reyes, 2003). However, these pharmacological blockers also affect other channels, at
higher concentrations (K+, Na+, L and N-type Ca2+ channels, and Cl- channels) which could
mislead the vessel myography results. Furthermore, the blockers block T-type Ca2+ channels in
different ways from one another. Mibefradil inhibits the channels in both dose-dependent and
use-dependent manner, as the channels become more active, inhibition increases because it
blocks the channels through the open pore (Morita et al., 2002; Jimenez et al, 2000; Lee et al,
1999; Perez-Reyes, 2003). Block by kurtoxin, a 63 residue peptide toxin, is voltage-dependent so
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a minor depolarization causes inhibition as kurtoxin binds to the channel’s voltage sensor, and
produces a complex modification of gating and accelerated deactivation (Horiba et al., 2008; Lee
et al., 1999; Perez-Reyes, 2003). R (-) efonidipine acts the same way as the DHPs act on L-type
channels. It stabilizes the inactivated states of T-type VGCCs (Furukawa et al., 2004; Lee et al.,
1999; Perez-Reyes, 2003).
Those properties suggest that the results we achieved could be complex because the
pressure-dependent effects we observe could be influenced by the ways in which the drugs bind
to different states of the channels and the parts of the channel that they interact with. The
experiments in Figures 11, 12, and 13 examining diameter changes could be repeated by addition
of nifedipine, and then the addition of one of the T-type blockers followed by the addition of a
second T-type blocker in combination with nifedipine and the first T-type blocker. This would
allow us to determine if effects of the blockers are additive or not. Additive effects would
suggest that other types of channels might also be affected by the blockers.
To overcome the non-specificity of the blockers the siRNA approach outlined above
could be introduced to rat cerebral arteries to investigate VGCCs in control and siRNA treated
arteries.
(iv) Computational approach.
The mathematical model that was constructed to predict blood flow in a cerebral arterial
network demonstrated, on a theoretical level, that T-type VGCCs are capable of significantly
affecting blood flow. This mathematical model permits many different questions to be
investigated on a theoretical level and could open the door for more wet lab experiments and
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investigations. Two questions that could be investigated and addressed with the model are: 1)
how might different expression levels of the channels at different vessel orders affect blood flow
and 2) At what order would a given dilation have the greatest effect. At the same time,
experiments could be done to screen for the quantitative expression of the VGCCs at different
vessel orders using reverse transcriptase PCR. Such experiments could indicate a relatively
greater role for T-type channels, compared to L-type channels, at different vessel orders. This
could also suggest that vessel myography could be used to investigate the role of T-type channels
in different vessel orders.
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Figure 4: mRNA expression of the α1-subunits of voltage-gated Ca2+ channels. Voltage-
gated Ca2+ channel expression in control tissues (skeletal muscle [CaV1.1], brain [CaV1.2,
CaV1.3, CaV2.1, CaV2.2, and CaV2.3], heart [CaV3.3], afferent arteriole [CaV3.1 and CaV3.2], and
retina [CaV1.4]) (A). Whole arteries or isolated smooth muscle cells from middle/posterior
cerebral arteries were collected and processed for RT-PCR. CaV α1-subunit expression in whole
cerebral arteries (B) and in enzymatically isolated smooth muscle cells (C), which indicate the
presence of L-type (CaV1.2), and T-type channels (CaV3.1 and CaV3.2). “M” indicates the size
marker lane (Performed by Rasha Abd El-Rahman).
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Figure 5: Protein expression of the α1-subunits of voltage-gated Ca2+ channels. Rat cortex
along with vessel segments from middle/posterior cerebral arteries were collected and separately
prepared for western blot analysis. Protein extracts were electrophoresed on 5.6%
polyacrylamide gel, transferred to PVDF and then labeled with primary antibodies against
CaV1.2, CaV3.1, and CaV3.2 (Performed by Rasha Abd El-Rahman).
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Figure 6: Inward Ba2+ currents in cerebral arterial smooth muscle cells. Whole cell patch
clamp electrophysiology was used to measure Ba2+ currents in smooth muscle cells isolated from
middle/posterior cerebral arteries. The voltage protocol consisted of a prepulse at -60 or -90 mV
(200 ms) followed by a series of steps (-80 to +40 mV, 200 ms, 10 mV increments). A)
Representative Ba2+ currents from a smooth muscle cell exposed to a prepulse at -90 mV (left)
or -60 mV (right). B) IV plot of peak inward Ba2+ current (n = 6) recorded with a prepulse at -90
mV (ο) or -60 mV (•). (Performed by Osama Harraz; cells were prepared by Rasha Abd El-
Rahman).
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Figure 7: Nifedipine-sensitive and -insensitive Ba2+ currents in cerebral arterial smooth
muscle cells. Whole cell patch clamp electrophysiology was used to measure Ba2+ currents in
smooth muscle cells isolated from middle/posterior cerebral arteries. The voltage protocol
consisted of a pre-pulse from -60 to -90 mV (200 ms) followed by a series of steps (-80 to +40
mV (300 ms, 10 mV increments). A) Representative Ba2+ current under control conditions and in
the presence of 50 nM nifedipine (left two panels), and representative Ba2+ current under control
conditions and in the presence of 300 nM nifedipine (right two panels). B) IV plot of peak
inward Ba2+ current along with the nifedipine-sensitive and –insensitive components (n = 6). C)
IV plot of the nifedipine-sensitive and -insensitive Ba2+ current expressed relative to maximal
current at +10 mV for 50 nM nifedipine (left) and 300 nM nifedipine (right). (Performed by
Osama Harraz; cells were prepared by Rasha Abd El-Rahman).
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Figure 8: Effects of mibefradil on the nifedipine-insensitive component of the inward Ba2+
current. Whole cell patch clamp electrophysiology was used to measure Ba2+ currents in
smooth muscle cells isolated from middle/posterior cerebral arteries. The voltage protocol
consisted of a pre-pulse from -60 to -90 mV (200 ms) followed by a series of steps (-80 to +40
mV, 200 ms, 10 mV increments). This protocol was run under control conditions and in the
presence of nifedipine (300 nM) + mibefradil (5 µM). A) Representative traces of Ba2+ current
under control conditions and in the presence of nifedipine and nifedipine plus mibefradil. The
nifedipine-sensitive component and the mibefradil-sensitive component were determined by
subtraction of measured currents as described in the Methods section. B) IV plot of nifedipine-
and mibefradil-sensitive currents (n = 9). C) Time constant (τ) for inactivation (left & center
panels) and activation (right panel) for the nifedipine-sensitive and mibefradil-sensitive
components at 10 mV (left) and 20 mV (center). The time constant for activation at 10 mV for
these current components is shown in the right panel (n = 9). * denotes significant difference
from the nifedipine-sensitive current (paired t-test, and one way ANOVA, P < 0.05) (Performed
by Yana Anfinogenova).
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Figure 9: Effects of kurtoxin on the nifedipine-insensitive component of the inward Ba2+
current. Whole cell patch clamp electrophysiology was used to measure Ba2+ currents in smooth
muscle cells isolated from middle/posterior cerebral arteries. The voltage protocol consisted of a
pre-pulse from -60 to -90 mV (200 ms) followed by a series of steps (-80 to +40 mV, 200 ms, 10
mV increments). This protocol was run under control conditions and in the presence of
nifedipine (300 nM) + kurtoxin (5 µM). A) Representative traces of Ba2+ current under control
conditions and in the presence of nifedipine and nifedipine plus kurtoxin. The nifedipine-
sensitive component and the kurtoxin-sensitive component were determined by subtraction of
measured currents as described in the Methods section. B) IV plot of nifedipine- and kurtoxin-
sensitive currents (n = 6). C) Time constant (τ) for inactivation (left & center panels) and
activation (right panel) for the nifedipine-sensitive and kurtoxin-sensitive components at 10 mV
(left) and 20 mV (center). The time constant for activation at 10 mV for these current
components is shown in the right panel (n = 6). * denotes significant difference from the
nifedipine-sensitive current (paired t-test, and one way ANOVA, P < 0.05) (Performed by Yana
Anfinogenova).
.
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Figure 10: Effects of efonidipine on the nifedipine-insensitive component of the inward Ba2+
current. Whole cell patch clamp electrophysiology was used to measure Ba2+ currents in smooth
muscle cells isolated from middle/posterior cerebral arteries. The voltage protocol consisted of a
pre-pulse from -60 to -90 mV (200 ms) followed by a series of steps (-80 to +40 mV, 200 ms, 10
mV increments). This protocol was run under control conditions and in the presence of
nifedipine (300 nM) + efonidipine (3 µM). A) Representative traces of Ba2+ current under
control conditions and in the presence of nifedipine and nifedipine plus efonidipine. The
nifedipine-sensitive component and the efonidipine-sensitive component were determined by
subtraction of measured currents as described in the Methods section. B) IV plot of nifedipine-
and efonidipine-sensitive currents (n = 6). C) Time constants (τ) for inactivation (left & center
panels) and activation (right panel) for the nifedipine-sensitive and efonidipine-sensitive
components at 10 mV (left) and 20 mV (center). The time constant for activation at 10 mV for
these current components is shown in the right panel (n = 6). * denotes significant difference
from the nifedipine-sensitive current (paired t-test, and one way ANOVA, P < 0.05) (Performed
by Yana Anfinogenova).
.
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Figure 11: The effects of nifedipine and mibefradil on myogenic tone. Cerebral arteries were
pressurized to 20 or 80 mmHg while diameter was monitored in the absence and presence of
nifedipine (50 and 300 nM) and nifedipine plus mibefradil (1 and 5 μM). A & B) Representative
traces revealing the effect of nifedipine and mibefradil on cerebral arteries pressurized to 20 or
80 mmHg. C & D) Summary data denoting the influence of nifedipine and mibefradil on
cerebral arteries pressurized to 20 (n = 7) or 80 (n = 6) mmHg. * and ** denote significant
increases from control and nifedipine (300 nM), respectively (paired t-test and one way
ANOVA, P < 0.05). E) The nifedipine- and mibefradil-sensitive vasomotor responses are plotted
as an absolute change or as a percent of maximal response. * denotes significant difference
from 20 mmHg (unpaired t-test, and one way ANOVA, P < 0.05) (Performed by Rasha Abd El-
Rahman).
.
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Figure 12: The effects of nifedipine and kurtoxin on myogenic tone. Cerebral arteries were
pressurized to 20 or 80 mmHg while diameter was monitored in the absence and presence of
nifedipine (50 and 300 nM) and nifedipine plus kurtoxin (1 μM). A & B) Representative traces
revealing the effect of nifedipine and kurtoxin on cerebral arteries pressurized to 20 or 80
mmHg. C & D) Summary data denoting the influence of nifedipine and kurtoxin on cerebral
arteries pressurized to 20 (n = 5) or 80 (n = 6) mmHg. * and ** denote significant increases
from control and nifedipine (300 nM), respectively (paired t-test and one way ANOVA, P <
0.05). E) The nifedipine- and kurtoxin-sensitive vasomotor responses are plotted as an absolute
change or as a percent of maximal response. * denotes significant difference from 20 mmHg
(unpaired t-test, and one way ANOVA, P < 0.05) (Performed by Rasha Abd El-Rahman).
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Figure 13: The effects of nifedipine and efonidipine on myogenic tone. Cerebral arteries
were pressurized to 20 or 80 mmHg while diameter was monitored in the absence and presence
of nifedipine (50 and 300 nM) and nifedipine plus efonidipine (3 μM). A & B) Representative
traces revealing the effect of nifedipine and efonidipine on cerebral arteries pressurized to 20 or
80 mmHg. C & D) Summary data denoting the influence of nifedipine and efonidipine on
cerebral arteries pressurized to 20 (n = 5) or 80 (n = 6) mmHg. * and ** denote significant
increases from control and nifedipine (300 nM), respectively (paired t-test, P < 0.05). E) The
nifedipine- and efonidipine-sensitive vasomotor responses are plotted as an absolute change or as
a percent of maximal response. * denotes significant difference from 20 mmHg (unpaired t-test,
and one way ANOVA, P < 0.05) (Performed by Rasha Abd El-Rahman).
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Figure 14: The effects of nifedipine and mibefradil on myogenic tone through a full range
of intravascular pressures. Cerebral arteries were sequentially pressurized from 20 to 100
mmHg while arterial diameter was monitored in the absence and presence of nifedipine (300
nM) and nifedipine plus mibefradil (5 µM). A) Representative traces revealing the effect of
nifedipine and mibefradil on cerebral arteries sequentially pressurized between 20 and 100
mmHg. B & C) The nifedipine- and mibefradil-sensitive vasomotor responses are plotted as an
absolute change or as a percent of maximal response. Absolute diameter (in μm) for control and
in Ca2+-free solution were as follows (n = 6): 20 mmHg, 175 ± 4, 188 ± 3; 40 mmHg, 171 ± 2,
211 ± 4; 60 mmHg, 153 ± 3, 220 ± 4; 80 mmHg,142 ± 4, 227 ± 4; and 100 mmHg, 135 ± 4 , 234
± 4. * denotes significant difference from the preceding pressure step (paired t-test and one way
ANOVA, P < 0.05) (Performed by Rasha Abd El-Rahman).
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Figure 15: Computational modeling predicts that T-type Ca2+ channel modulation alters
tissue blood flow. A) A computational model (Goldman & Popel, 2000; Pries et al., 1990;
Pries et al., 1994) was used to calculate two-phase steady-state flow through an arterial network
consisting of 7 arterial branches whose diameter/ length approximates the posterior cerebral
arterial network diverging off the Circle of Willis. This model implements conservation of blood
and RBC volume flow at each node joining vessel segments and includes known blood rheology.
In addition to geometric information, this model required specification of boundary conditions in
the form of hematocrit (0.4) and pressure changes between the inlet (75 mmHg) and outlet (50
mmHg) segments. B) The impact of different dilation (5-15 %) on blood flow for the different
vessel orders. C) Represents the percentage change in blood flow at different percentages of
dilation (5, 10, and 15 %). (Computer programming by Daniel Goldman and simulations
performed by Daniel Goldman and Rasha Abd El-Rahman).
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Chapter Three: CaV3.2 Channels and the Induction of a Negative Feedback Response
In Cerebral Arterial Smooth Muscle
Rasha R. Abd El-Rahman, Osama F. Harraz, Kamran Bigdely-Shamloo, Sean M. Wilson,
Monica Rubalcava, Rania E. Mufti, Albert L. Gonzales, Scott Earley, Edward J. Vigmond
and Donald G. Welsh
I was responsible for structural and functional data collection and analysis and I wrote drafts of
the manuscript to be submitted for publication. Osama Harraz and I carried out the perforated
patch clamp experiments. Suzanne B. Welsh performed membrane potential measurements.
Computational modeling was performed by Kamran Bigdely-Shamloo and Edward J. Vigmond,
and Sean M. Wilson and Monica Rubalcava were responsible for the Ca2+ spark experiments.
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3.1 INTRODUCTION
Cerebral resistance arteries control tissue blood flow through alterations in smooth
muscle contractility initiated by neuronal stimulation, hormonal and metabolic state, and
intraluminal pressure (Wray et al., 2005). These factors initiate change in vessel diameter in part
by increasing cytosolic [Ca2+], a response that augments myosin light chain (MLC20)
phosphorylation via the regulation of myosin light chain kinase (MLCK) and myosin light chain
phosphatase (MLCP). The elevation in cytosolic [Ca2+] is sustained by the influx of extracellular
Ca2+ and by Ca2+ release from the sarcoplasmic reticulum (SR) (Berridge et al., 2000; Carafoli et
al., 2001). Extracellular Ca2+ influx is in turn driven by resting membrane potential and the
activity of voltage-gated Ca2+ channels (VGCCs).
VGCCs are heteromultimeric protein complexes which consist of a pore-forming α1-
subunit containing four domains (I-IV) of six transmembrane segments. In vascular smooth
muscle, CaV1.2 (L-type) is the key channel regulating extracellular Ca2+ influx, ultimately
leading to MLC20 phosphorylation and consequently tone development (Knot & Nelson, 1998;
Lambert et al., 1997). While CaV1.2 channels dominate from a functional perspective, recent
studies have additionally noted the presence of T-type Ca2+ channels including CaV3.1 and
CaV3.2. Arguably Ca2+ influx through T-type channels could, like their L-type counterpart,
facilitate the global rise in cytosolic [Ca2+] needed to drive smooth muscle cell contraction and
arterial constriction. Oddly, however, studies centered on CaV3.2 have noted that blockade of
these channels results in arterial constriction rather than dilation (Chen et al., 2003). To explain
these paradoxical observations, the investigators suggested that Ca2+ influx through CaV3.2 may
act in a localized manner to regulate defined Ca2+-dependent targets. It was speculated that these
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targets could range from nitric oxide synthase in endothelial cells to BKCa channels in vascular
smooth muscle (Chen et al., 2003).
In this study, we investigated whether Ca2+ influx through CaV3.2 channels can indeed
operate in a localized manner to activate a Ca2+-dependent target to influence cerebral arterial
tone. In this regard, we show through the use of vessel myography, confocal and electron
microscopy, computational modelling, and electrophysiology that flux through these channels
within a discrete domain activates the cytosolic gate of RyRs. This in turn leads to the repetitive
generation of Ca2+ sparks, finite SR Ca2+ release events that activate BKCa channels and provide
negative feedback to depolarized and constricted arteries, resulting in dilation of the arteries.
Overall, this is the first study to indicate that T-type Ca2+ channels can drive a Ca2+-induced Ca2+
release (CICR) event that is key to setting arterial tone and tissue perfusion in the cerebral
circulation.
3.2 MATERIALS AND METHODS
3.2.1 Animal procedures.
Animal procedures were approved by the Animal Care and Use Committee at the
University of Calgary. Briefly, female Sprague–Dawley rats (10–12 weeks of age) were killed
via carbon dioxide asphyxiation. The brain was carefully removed and placed in cold phosphate-
buffered (pH 7.4) saline solution containing (in mM): 138 NaCl, 3 KCl, 10 Na2HPO4, 2
NaH2PO4, 5 glucose, 0.1 CaCl2 and 0.1 MgSO4. Middle and posterior cerebral arteries were
carefully dissected out of surrounding tissue and cut into 2–3 mm segments.
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3.2.2 Vessel myography and measurements of membrane potentials.
Arterial segments were mounted in a customized arteriograph and superfused with warm
(37 oC) physiological salt solution (PSS; pH 7.4; 21% O2, 5% CO2, balance N2) containing (in
mM): 119 NaCl, 4.7 KCl, 20 NaHCO3, 1.1 KH2PO4, 1.2 MgSO4, 1.6 CaCl2 and 10 glucose (Kuo
et al., 1991; Welsh et al., 2000). To limit the endothelium’s tonic dilatory influence on myogenic
tone development (Knot et al., 1999; Kuo et al., 1991), air bubbles were passed through the
vessel lumen (1–2 min); successful removal of the endothelium was confirmed by the loss of
bradykinin-induced dilation. Arteries were equilibrated for 30 min at 15 mmHg and contractile
responsiveness assessed by briefly exposing (~10 s) the tissue to 60 mM KCl. Following
equilibration, intravascular pressure was increased incrementally from 20 to 100 mmHg and
arterial tone was monitored under control conditions and in the presence of paxilline (1 µM,
BKCa inhibitor) and/or Ni2+ (50 µM, CaV3.2 blocker). Maximal arterial diameters were
subsequently ascertained in Ca2+-free PSS (zero externally added Ca2+ + 2 mM EGTA). Smooth
muscle membrane potential (Vm) was assessed by inserting a glass microelectrode backfilled
with 1 M KCl (tip resistance =120–150 MΩ) into the vessel wall. Independent measurements of
Vm were first made at 60 mmHg and then in the presence of Ni2+ (50 µM), a CaV3.2 blocker. The
criteria for successful cell impalement included: (1) a sharp negative Vm deflection upon entry;
(2) a stable recording for at least 1 min following entry; and (3) a sharp return to baseline upon
electrode removal.
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3.2.3 Immunohistochemistry.
Rats were anaesthetized with sodium pentobarbital and perfused intra-cardially with 250
ml of 0.1 M phosphate-buffered saline solution (PBS, pH 7.4), followed by 100 ml of 4%
paraformaldehyde (pH 7.4) in PBS at room temperature. Brains were then removed and post-
fixed in 4% paraformaldehyde for 1 h at 22 °C. Posterior and middle cerebral arteries (MCAs)
were subsequently excised and placed in a conical vial with a standard working PBS solution
containing 3% goat serum, 0.1% Tween, and 1% dimethylsulphoxide (DMSO). Primary
antibodies against smooth muscle actin (monoclonal antibody, 1:25 dilution), CaV1.2 (1:25
dilution), CaV3.1 (1:100 dilution), CaV3.2 (1:100 dilution), BKCa (1:100 dilution), IP3R (1:100
dilution), neurofilament (1:100 dilution) and RyR2 (1:100 dilution) were added to the working
solution and incubated for 48 h at 4 °C. The Ca2+ channel and RyR antibodies were polyclonal.
Next, the tissues were washed 3 times with PBS (15 min each, 22 °C) and then incubated with
working PBS solution containing secondary antibodies for 4 h at 22 °C. Following a final set of
washes with PBS (3X), sections were mounted on gel-coated slides and covered with antifade
medium (90% glycerol/ PBS/ 0.1% p-phenylenediamine; pH 10). Coverslips were sealed with
nail polish and slides were stored at -20 °C. Controls consisted of omitting the primary
antibodies or preabsorbing them with an excess of purified antigen peptide. All reactions
involved the use of fluorophore-conjugated secondary antibodies (Alexa Fluor 488-goat anti-
mouse IgG (1:1000 dilution), Alexa Fluor 555-goat anti-rabbit IgG (1:1000 dilution), Alexa
Fluor 405-goat anti-chicken IgG (1:1000 dilution)). Immunolabelling was assessed using an
Olympus FV300 BX50 confocal microscope equipped with (405 λ, blue), (555 λ, red) and (488
λ, green) filter sets.
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3.2.4 Electron tomography
Rats were anaesthetized with sodium pentobarbital and perfused intra-cardially with 250
ml of 0.1 M phosphate-buffered saline solution (PBS, pH 7.4), followed by 100 ml of 4%
paraformaldehyde (pH 7.4) in PBS at room temperature. Brains were then removed and post-
fixed in 4% paraformaldehyde (22 °C) for 1 h. Cerebral arteries were dissected free from
surrounding tissues. Samples were immersed in 1.6% paraformaldehyde and 2.5%
glutaraldehyde in 0.1 M sodium cacodylate buffer (Sabatini et al.,1962) at pH 7.4 at 4 °C
overnight. After washing three times with the buffer (0.1 M sodium cacodylate), the samples
were post-fixed in 1% osmium tetroxide buffered with sodium cacodylate for 1 h at room
temperature, dehydrated through a graded series of acetone concentrations (30 – 100%) and
embedded in Epon 812 mixture resin. Thick sections (300-400 nm) were cut on a Reichert-Jung
Ultracut E microtome using a diamond knife and collected on single hole grids with Formvar
supporting film. The sections were first stained with 20 % aqueous uranyl acetate/Reynolds’s
lead citrate (Reynolds, 1963) and then placed on one side of a transmission electron microscopy
slot grid (1 × 2 mm slot) covered with a continuous Formvar film (∼40 nm) to dry (10 min).
Colloidal gold particles (10 nm diameter) were then placed on both sides of the grid to serve as
fiducial markers, and a thin carbon coating was applied for mechanical stabilization and to
reduce electric charging. Once prepared, sections were viewed on a Tecnai F20 transmission
electron microscope (200 keV), regions of interest were defined, and images were captured with
a 1,024 × 1,024 charge-coupled device camera (GIF 794, Gatan, Pleasanton, CA). To perform
dual-axis transmission electron-microscopic tomography, Serial EM software (Mastronarde,
2005) was employed to capture one image per degree of sample rotation (136 degrees in total).
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Tomographic reconstruction was performed by weighted back projection using the IMOD
software package (Kremer et al., 1996; Mastronarde, 1997). This yielded a contiguous stack of
two-dimensional photomicrographs with ∼4-nm resolution. The same software was used to trace
subcellular structures on each section of the contiguous stack. We then compiled the traces to
produce a 3-D rendition of the microdomain structure.
3.2.5 Immunogold-labeling.
Rats were anaesthetized with sodium pentobarbital and perfused intra-cardially with 250
ml of 0.1 M phosphate-buffered saline solution (PBS, pH 7.4), followed by 100 ml of 4%
paraformaldehyde (pH 7.4) in PBS at room temperature. Brains were then removed and post-
fixed in 4% paraformaldehyde (22 °C) for 1 h and cerebral arteries were dissected free from
surrounding tissues. In accordance with directions provided with the Aurion immunogold reagent
kit, fixed arteries were exposed to 0.1% sodium borohydride in PBS (15 min), 0.05% Triton X-
100 (30 min), Aurion blocking solution 1 h at 4 oC and then PBS (2 x 10 min). Primary
antibodies (1:100 dilution) were subsequently added to the buffer and incubated for 48 h at 4 °C.
Tissues were subsequently washed with buffer (6 x 10 min), exposed to incubation medium
containing ultra-small gold labelled secondary antibody particles (0.8 nm labeled goat anti-
rabbit, 1:100 dilution, 48 h at 4 °C) and then washed with incubation buffer (6 x 10 min) and
PBS (2 x 10 min). Prepared arteries were fixed again in PBS containing 2.5% glutaraldehyde (2
h) and then washed in PBS (2 x 10 min). Arteries were sequentially processed as follows: 1)
Enhancement conditioning solution (ECS, 4 x 10 min); 2) Silver enhancement solution (2 h); 3)
0.03 M sodium thiosulphate in ECS (10 min); 4) ECS wash (4 x 10 min); and 5) PBS wash (2 x
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10 min). Tissues were then postfixed for 1 h in a 1% osmium tetroxide-PBS solution, dehydrated
in ethanol, and embedded in Epon resin. Ultrathin sections of about ~70 nm were cut, lightly
stained with 2% aqueous uranyl acetate and Reynolds lead citrate (15 min) (Reynolds, 1963),
and viewed/photographed using a Hitachi H7650 transmission electron microscope (80 keV) and
an AMT 16000 digital camera.
3.2.6 Isolation of arterial smooth muscle cells.
Smooth muscle cells from middle and posterior cerebral arteries were enzymatically
isolated as previously described (Luykenaar et al., 2004; Talavera et al., 2001). Briefly, arterial
segments were placed in an isolation medium (37 °C, 10 min) containing (in mM): 60 NaCl, 80
Na-glutamate, 5 KCl, 2 MgCl2, 10 glucose and 10 HEPES with 1 mg/ml albumin (pH 7.4).
Vessels were then exposed to a two-step digestion process that involved: 1) 15 min incubation in
isolation medium (37 °C) containing 0.6 mg/ml papain and 1.8 mg/ml dithioerythritol; and 2) a
15 min incubation in isolation medium containing 100 μM Ca2+, 0.7 mg/ml type F collagenase
and 0.4 mg/ml type H collagenase. Following treatment, tissues were washed repeatedly with
ice-cold isolation medium and triturated with a fire-polished pipette. Liberated cells were stored
in ice-cold isolation medium for use the same day.
3.2.7 Proximity ligation assay (PLA).
Freshly isolated smooth muscle cells were studied using the Duolink in situ PLA
detection kit. Briefly, cells were fixed in phosphate buffered saline (PBS) containing 4%
paraformaldehyde for 15 min, permeabilized in PBS containing 0.1% Tween for 15 min and
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quenched in PBS containing 100 mM glycine for 5 min. Cells were then washed with PBS,
blocked for 30 min at 37 °C in Duolink blocking solution, and incubated overnight at 4 °C with
pairs of primary antibodies in Duolink antibody diluent solution (rabbit anti-CaV3.2 and mouse
RyR2). Control experiments were done with no primary antibody or only one primary antibody.
Cells were labelled with Duolink PLA PLUS and MINUS probes for 2 h at 37 °C. The secondary
antibodies of the PLA PLUS and MINUS probes are attached to synthetic oligonucleotides that
hybridize when in close proximity (i.e. ≤ 40 nm separation). The hybridized oligonucleotides are
then ligated prior to rolling circle amplification. The concatemeric amplification products
extending from the oligonucleotide arm of the PLA probes were detected using red fluorescent
fluorophore-tagged, complementary oligonucleotide sequences and a Zeiss Apotome
epifluorescence microscope. Thus the red fluorescence product is localized to sites where the two
different primary antibodies bind with separation ≤ 40 nm.
3.2.8 Computational modeling.
We have constructed a mathematical model, which incorporates important ultrastructure
and Ca2+ handling features of the vascular smooth muscle cell in cerebral arteries. The smooth
muscle cell is described as a cylinder, 80 µm in length and 5 µm in diameter, as shown in Fig.
21A. For simulation purposes, the cell is subdivided into segments 8.5 µm in length. Based on
microscopic data, in each segment there is a circumferential band, which is interrupted in
approximately two locations, thus forming two discrete sections circumferentially. In the model,
each segment is therefore further sub-divided into two semi-cylindrical “slices”. Simulations
presented in this paper are based on the behaviour of one of these slices. The frequencies of Ca2+
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spark-like events have been scaled to represent the whole cell, based on the assumption that
individual slices act independently. The model includes mechanisms responsible for Ca2+
dynamics in the cerebral arterial smooth muscle cell. This includes the Na+-Ca2+ exchanger
(NCX), plasma membrane Ca2+ ATPase (PMCA), sarco/endoplasmic reticulum Ca2+ ATPase
(SERCA), ryanodine receptor (RyR), calmodulin, calsequestrin, and the Ca2+ channels CaV1.2,
CaV3.1 and CaV3.2. In its present form, the model is thus limited to investigations of Ca2+
dynamics in response to voltage-clamp stimuli. As illustrated in Fig. 21, the model explicitly
includes the interaction between CaV3.2 and RyR in a restricted microdomain (“subspace”),
representing the 10-15 nm space between caveolae in the plasma membrane and the SR.
Mathematically, the model comprises 12 ordinary differential equations, which were solved
using the “ode15s” ODE solver in MATLAB (The MathWorks, Natick, MA, USA).
3.2.9 Electrophysiology.
Perforated patch-clamp electrophysiology was used to measure whole-cell K+ currents in
isolated smooth muscle cells. The bathing solution contained (in mM): 134 NaCl, 4 KCl, 2
MgCl2, 2 CaCl2, 10 glucose, and 10 HEPES (pH 7.4). The pipette solution contained (in mM):
110 K aspartate, 30 KCl, 10 NaCl, 2 MgCl2, 10 HEPES, and 0.05 EGTA (pH 7.2) with 200
mg/ml amphotericin B. Membrane currents were recorded while the cells were held at a steady
membrane potential of -40 or -20 mV. Whole-cell currents were recorded on an Axopatch 200B
amplifier (Axon Instruments, Inc), filtered at 1 kHz, digitized at 5 kHz and stored on a computer
for subsequent analysis with Clampfit 10.2 software. Cell capacitance ranged between 14-18 pF
and was measured with the cancellation circuitry in the voltage-clamp amplifier. A 1 M NaCl–
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agar salt bridge between the reference electrode and the bath solution was used to minimize
offset potentials (< 2 mV). All experiments were performed at room temperature (20 –22 °C).
3.2.10 Ca2+ sparks.
Ca2+ sparks were measured in rat myocytes in situ using the en-face preparation
technique (Hadley et al., 2012) with the Ca2+-sensitive dye Fluo-4 AM using a Zeiss LSM 710
NLO laser scanning confocal imaging workstation on an inverted microscope platform (Zeiss
Axio Observer Z1). Fluo-4 AM was dissolved in DMSO and added from a 1 mM stock to the
arterial suspension at a final concentration of 10 µM, along with 0.1% pluronic F127 for 1 - 1.5 h
at room temperature in the dark in balanced salt solution. Arterial segments were then washed for
30 min to allow for dye de-esterification and then cut into linear strips. The arterial segments
were pinned to Sylgard blocks and placed in an open bath imaging chamber mounted on the
confocal imaging stage. Cells were illuminated at 488 nm with a krypton argon laser and the
emitted light was collected using a photomultiplier tube (PMT). Line scans were imaged at 529
frames generated every 1 s. The acquisition period for Ca2+ spark recordings was 18.9 s. The
resultant pixel size ranged from 0.11 to 0.068 µm per pixel. To ensure that sparks within the cell
were imaged, the pinhole was adjusted to provide an imaging depth of ~2.5 µm. This depth is
roughly equivalent to the width of 50% of the cell based on morphological examination of live
preparations, and was equivalent to that used in previous studies (Hadley et al., 2012; Janiak et
al., 2001). Ca2+ sparks were analyzed by visual inspection for the frequency of firing. The
number of sparks per 100 micrometer per second was computed from these observed sparks. The
Spark Master plug-in for Image J (Picht et al., 2007) was used to determine the fractional
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fluorescence intensity for the Ca2+ spark events. In these studies, the threshold for Ca2+ spark
event detection was 3.2 times the standard deviation of the background noise above the mean
background level. Prior to analysis the background fluorescence was subtracted from each image
assuming homogeneous background levels in each cell (Hadley et al., 2012).
3.2.11 Statistical analysis.
Data are expressed as means ± S.E. and n indicates the number of vessels or cells. No
more than two experiments were performed on vessels from a given animal. Where appropriate,
paired, unpaired t-tests, and one way ANOVA were performed to compare the effects of a given
condition/treatment on arterial diameter, or whole-cell current (see figure legends for specific
details). P values ≤ 0.05 were considered statistically significant.
3.2.12 Solutions and chemicals
All buffers, chemicals and reagents were purchased from Sigma-Aldrich (St. Louis, MO)
unless otherwise stated. Donkey and goat sera were purchased from Jackson Immuno-Research
(West Grove, PA). The primary antibodies against smooth muscle actin and neurofilament were
obtained from Abcam Inc. (Cambridge, MA) whereas those primary antibodies directed against
CaV1.2, 3.1, 3.2, and RyR were purchased from Alomone (Jerusalem, Israel). Secondary
antibodies, which included Alexa Fluor 488-goat anti-mouse IgG, Alexa Fluor 405-goat anti-
chicken and Alexa Fluor 555-goat anti-rabbit IgG, were obtained from Invitrogen Life
Technologies (Eugene, OR). Aurion immunogold reagents and the PLA Duolink detection kit
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were purchased from Electron Microscopy Sciences (Hatfield, PA) and Olink (Uppsala,
Sweden), respectively.
3.3 RESULTS
3.3.1 Vasomotor and electrical effects of CaV3.2 blockade.
We started our investigation by assessing the vasomotor effect of Ni2+, a selective CaV3.2
inhibitor at micromolar concentrations (Lee et al., 1999; Perez-Reyes, 2003), on pressurized
cerebral arteries. Under control conditions, arterial tone progressively increased as vessels were
sequentially pressurized from 20 to 100 mmHg (Figure 16A & 16B). Interestingly, Ni2+
blockade of CaV3.2 channels enhanced myogenic tone at pressures < 80 mmHg. This Ni2+-
induced rise in tone coincided with an arterial depolarization of 5 ± 0.9 mV (Figure 16C, 16D &
1E). This latter result suggests that Ca2+ influx through CaV3.2 channels indirectly moderates
constrictor responses by enhancing the activity of a K+ conductance involved in establishing
negative membrane potentials to oppose constrictor responses. This could include BKCa channels
whose activity is gated by the transient release of Ca2+ from the SR via ryanodine receptors
(RyR).
3.3.2. Ca2+ channel localization.
To explore the potential spatial relationship between CaV channels and RyR, we
performed immunohistochemistry on whole fixed cerebral arteries. Tissues were labeled with
antibodies directed against smooth muscle actin and a single channel of interest (CaV3.2, RyR2,
or CaV1.2). In Figure 17A, left panel, the upper composite panel indicates that smooth muscle
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actin runs lengthwise in cells and periodically disappears as this structural protein leaves the
focal plane. Intriguingly, CaV3.2 labeling ran perpendicular to the long axis of the cells and was
observed in the regions devoid of smooth muscle actin. This CaV3.2 labeling pattern is
particularly evident in the gallery subpanels. A similar labelling pattern was observed for RyR
(Figure 17A), an observation suggestive of RyR and CaV3.2 lying in close apposition to one
another. In striking contrast, CaV1.2 labeling ran lengthwise in a ribbon like fashion along
smooth muscle cells (Figure 17B, center and magnified right panel). Note that a portion of the
topical labeling observed in the latter preparations was attributed to CaV1.2 protein expression in
perivascular nerves (Figure 18D). Secondary antibody and peptide control experiments were
performed for all experiments and were negative (Figure 18). The preceding results suggest a
possible close association between CaV3.2 and RyR; we therefore performed a Proximity
Ligation Assay (PLA). Isolated smooth muscle cells were probed with primary antibodies
against CaV3.2 and RyR and then exposed to secondary antibodies tagged with synthetic
oligonucleotide strands. If the two proteins are within 40 nm of one another, the strands
hybridize and a multiplication reaction produces a visible red fluorescent product. Figure 19A-
19C indicates that a red fluorescent product was found at discrete sites in rat cerebral arterial
smooth muscle cells. This fluorescent product was observed both near and distal to nuclei. In
striking contrast, PLA product was absent from cells exposed to both secondary antibodies alone
and in experiments where one of the primary antibodies (CaV3.2 or RyR) was omitted (Figure
19D-19F; Figure 19G-19I; & Figure 19J-19L). With evidence of CaV3.2 and RyR channels
located close to one another, we performed electron tomography to better ascertain the structural
relationship between the SR and the plasma membrane. Image analysis and model
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reconstructions of tomograms revealed a repetitive microstructure along the length of the smooth
muscle cell, comprised of caveolae and SR-like structures (Figure 20A-20C). Immuno-gold
labeling was subsequently performed on whole mounted tissues to further delineate CaV3.2 and
RyR localization. As indicated in Figure 20D-20F, CaV3.2 labeling was observable in caveolae.
In comparison, a portion of RyRs was localized to internal membranous structures that lay in
close apposition to caveolae. In this study, all the figures presented have the same magnification
values of 12000X or 30000X. For each antibody 5 figures were chosen. The images were
selected for the analysis if they showed a complete smooth muscle cell. Table 2 shows results of
analysis for each image. For RyR labelling 27± 2 % of the labeling occurred near the caveolae;
for CaV3.2 32 ± 4 % of the labelling was in the PM in caveolae. Cumulatively, the preceding
results indicate the presence of microdomains in cerebral arterial smooth muscle comprised of
caveolae and SR and which contain T-type Ca2+ (CaV3.2) and RyR channels, respectively.
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Image #
RyR CaV3.2
Number of
labelled
points
Number of
points near
caveolae
% of points
near
caveolae
Number of
labelled
points
Number of
points in
caveolae
% of points
in caveolae
1 45 13 29 124 29 23
2 70 19 27 114 30 26
3 30 9 30 107 35 33
4 43 9 21 148 49 33
5 37 10 21 155 70 45
Average 27 32
Standard
Error
2 4
Table 2: Immunogold-labeling for RyR and CaV3.2
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3.3.3 Computational modeling.
To predict whether Ca2+ flux through CaV3.2 channels could possibly initiate a CICR-like
response by activating RyR, we built a microdomain model (Figure 21A) using our structural
data, existing electrophysiological measurements of CaV channels (Aslanidi et al., 2010; Liao et
al., 2005) and published mathematical representations of RyR (Keizer & Levine, 1996). Figure
21B indicates that when voltage is stepped from -60 to -40 mV, repetitive Ca2+ spark-like events,
with an amplitude of 30 µM and a frequency of 0.12 Hz, were predicted to occur in the plasma
membrane subspace. These Ca2+ spark-like events were eliminated or reduced in frequency by ~
50-60 % in simulations in which RyR and CaV3.2 were respectively blocked (Figure 21C &
21D). Additional simulations in Figure 21E show that the frequency of Ca2+ spark-like events is
graded with depolarization. Our computational work supports the view that CaV3.2 channels
could drive a CICR response. In theory, these discrete events could activate BKCa channels and
moderate cerebral arterial tone.
3.3.4 BKCa currents and Ca2+ sparks.
To explore the relationship between CaV3.2, RyR and BKCa channels, perforated patch
clamp electrophysiology was used to monitor spontaneous transient outward current (STOCs),
events that arise when SR-driven Ca2+ sparks activate a discrete pool of BKCa channels. Under
control conditions (in the absence of any blockers), repetitive STOC generation was observed in
rat cerebral arterial smooth muscle cells with the frequency increasing as the holding potential
was depolarized from -40 to -20 mV. In keeping with CaV3.2 channels facilitating STOC
generation, 50 µM Ni2+ (a selective CaV3.2 inhibitor) reduced the frequency of these events by
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~60% (Figure 22A). In comparison, STOCs were eliminated by 1 µM paxilline, a BKCa channel
inhibitor (Figure 22B). Fluo-4 confocal imaging was subsequently used to monitor Ca2+ sparks,
with line scan analysis revealing that these events were present in rat cerebral arteries (Figure
22C). In regions where Ca2+ sparks were detectable, Ni2+ application was found to reduce event
frequency by 50% (Figure 22D). Thus 50 µM Ni2+ reduces the frequency of both Ca2+ sparks and
STOCs. Cumulatively, these results indicate that Ca2+ influx through CaV3.2 channels can indeed
enhance BKCa channel activity via a CICR-driven process that involves RyR and Ca2+ spark
induction.
3.3.5 Functional implications of CaV3.2 blockade.
If CaV3.2 can drive STOC activity, the functional consequences of this event would be
the generation of a modest hyperpolarization that feeds back negatively upon constriction as
indicated by the data in Figure 16A. To further test this possibility additional experiments were
performed and provided evidence to support this concept. Like the Ni2+-induced constriction
observed in Figure 16A and 16B and reprinted for clarity in Figure 23A and 23B, paxilline (1
µM, BKCa channel blocker) enhanced myogenic constriction at intravascular pressures < 80
mmHg (Figure 23C and 23D). When Ni2+ and paxilline were sequentially added on top of one
another, the first agent induced constriction while the second had no additive effect (Figure 23E-
23H). These latter results are consistent with CaV3.2 and BKCa channels working in series with
one another within a single signaling pathway to limit constriction.
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3.4 DISCUSSION
T-type Ca2+ channels are present in the cerebral circulation but their role in tone
development is controversial. Functional observations vary widely with some results indicating
they play no significant role (Oguri et al., 2010; Perez-Reyes, 2003; Rodman et al., 2005) while
others have linked Ca2+ influx to vasoconstriction (Akaike et al., 1989; Hansen et al., 2011;
Poulsen et al., 2011) and to vasodilation (Chen et al., 2003). Past work has noted that the role of
CaV3.2 in mediating either vasoconstriction or vasodilation is not clear. In this study, we
employed functional, structural, computational and electrical approaches to show that CaV3.2
facilitates cerebral arterial dilation. Our work suggests CaV3.2 channels are part of a restricted
microdomain along with RyRs. When CaV3.2 opens, the influx of Ca2+ into the subspace
between the plasma membrane and the SR would lead to RyR gating and the generation of Ca2+
sparks. These discrete Ca2+ events initiate STOCs, BKCa channel events that induce
hyperpolarization and vasodilation. Our results indicate that Ca2+ flux through specific T-type
channels (CaV3.2) is not directed to the bulk cytosol but rather to microdomains where they
evoke a CICR and subsequently K+ channel activity to limit arterial constriction.
3.4.1 Background.
Increases in [Ca2+]i elicit smooth muscle constriction by enhancing MLC20
phosphorylation, a process controlled by the balanced activation of myosin light chain kinase
(MLCK) and myosin light chain phosphatase (MLCP) (Deng et al., 2002). Ca2+ influx through
voltage-gated Ca2+ channels is the principal regulator of [Ca2+]i and the principal voltage-gated
Ca2+ channel α1-subunits expressed in cerebral arterial smooth muscle cells are CaV1.2 (L-type),
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CaV3.1 and CaV3.2 (T-type), as shown in Chapter 2. It is well recognized that CaV1.2 plays a key
role in controlling steady-state Ca2+ entry and consequently tone development. This can be
readily observed by applying dihydropyridines and monitoring the change in [Ca2+]i and arterial
diameter (Harder, 1984; Knot & Nelson, 1998). The function of T-type Ca2+ channels in arterial
smooth muscle has yet to be convincingly determined and it is a topic of active debate (Perez-
Reyes, 2003; Rodman et al., 2005). In this regard, several studies have argued that T-type
channels are unimportant in tone generation and that their primary role centers on the regulation
of cell proliferation (Oguri et al., 2010; Perez-Reyes, 2003; Rodman et al., 2005). Others,
however, have presented alternative findings, suggesting that Ca2+ influx through T-type
channels in some cases facilitates arterial constriction but also can interestingly facilitate dilation
(Braunstein et al., 2009; Chen et al., 2003; Hansen et al., 2011; Kuo et al., 2010; Poulsen et al.,
2011). The latter result is particularly intriguing, and the authors (Chen et al., 2003)
hypothesised that the channels involved in facilitating vasodilation might be targeted to a smaller
cellular domain, where they are able to potentially control the activity of BKCa channels.
3.4.2 Mechanism of CaV3.2-induced vasodilation.
It is in this context that we began to mechanistically examine CaV3.2 and its unexpected
role in arterial tone development. Using a Ni2+ concentration that selectively targets CaV3.2, we
first observed, analogous to the findings of Chen et al. (Chen et al., 2003), the ability of this
inhibitor to constrict pressurized cerebral arteries. This Ni2+-induced constriction was
particularly prominent when vessels were pressurized between 20-60 mmHg, and was
diminished at higher pressure. With this constrictor effect being inconsistent with Ca2+ influx
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causing a bulk rise in [Ca2+]i, we hypothesized that flux might be more localized and directed
towards the regulation of a Ca2+-sensitive conductance. As such, immunohistochemistry was
performed on whole fixed arteries, and we observed CaV3.2 labeling in regions devoid of smooth
muscle actin. A similar labelling pattern was observed for RyR, an SR Ca2+ release channel that
can generate Ca2+ sparks. This labeling pattern contrasted with that of CaV1.2, which showed a
labelling pattern distinct from CaV3.2 with labelling running parallel to the long axis of the
smooth muscle cell in thin ribbon-like threads. This pattern is in keeping with the role of CaV1.2
in elevating global [Ca2+]i. A similar labelling pattern was observed for CaV3.1, BKCa, and IP3R
(see appendix). With immunohistochemical patterns indicating a potentially close association
between CaV3.2 and RyR, a proximity ligation assay (PLA) was performed. This assay entails
the application of two primary antibodies to isolated smooth muscle cells followed by the
addition of secondary antibodies conjugated to oligonucleotides. When in close proximity, the
oligonucleotide tails interact forming a circular construct that can be replicated and fluorescently
visualized as a discrete red product. Consistent with CaV3.2 and RyR residing within 40 nm of
one another, distinctive red fluorescent labeling could be observed particularly in regions of the
cell surrounding the nucleus. Some cells showed labelling closer to the surface, but they tended
to be contracted. Similar labeling was absent from cells exposed to secondary antibodies alone or
where one of the primary antibodies had been omitted from the ligation assay. Further
experiments would need to be carried out to investigate whether the methods for cell fixation
might affect the labelling and, if so, to find an optimal method. As CaV3.2 resides on the plasma
membrane and RyR resides on the SR, we subsequently employed electron microscopy to better
define regions of close association between the SR and the plasma membrane. Electron
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tomography enables the construction and visualization of 3-D model structures from a
contiguous stack of 2-D photomicrographs (~3.5 nm resolution) rendered from a thick tissue
section. In regions peripheral to the nucleus, the SR came into close proximity with the plasma
membrane and, in particular, in close proximity to the caveolae. Immunogold labelling suggested
that some CaV3.2 resides in caveolae and some RyRs are found underneath caveolae.
We built a computational model to first address on a theoretical level whether Ca2+ influx
through T-type Ca2+ channels could be predicted to cause spontaneous repetitive opening of RyR
to produce Ca2+ spark-like events. The computational model was built based upon the literature,
structural measurements, localization, and measurements of channel activity (Aslanidi et al.,
2010; Bondarenko et al., 2004; Cannell & Soeller, 1997; Keizer & Levine, 1996; Liao et al.,
2005). It consists of an 8.5 µm slice of an arterial smooth muscle cell: a microdomain 15 nm in
length is comprised of SR and a subspace of appropriate volume, and membrane proteins have
been distributed accordingly. Our model shows that depolarization could indeed elicit repetitive
spark-like events, which can be graded as a function of voltage, and these events are completely
blocked by RyR blockade, and attenuated by about 50-60 % by CaV3.2 blockade. The
computational model provides theoretical support for the idea that the CaV3.2 can gate the
opening of RyR channels. This provided motivation for further investigations to determine if we
could find biological evidence to support this conclusion.
To determine whether there is an experimentally observable, functional connection
between the influx of Ca2+ via T-type channels and 1) BKCa channel activation and 2) Ca2+ spark
production, we first measured spontaneous transient outward currents (STOCs) using the
perforated patch technique in the absence and presence of CaV3.2 and BKCa channel blockers.
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Our results showed a reduction or elimination of STOCs in the presence of Ni2+ or paxilline at -
40 and -20 mV, suggesting that CaV3.2 channels might regulate BKCa channel activity. Ca2+
spark events have been detected in pressurized arteries and found to be triggered from RyR
channels in the SR (Brayden & Nelson, 1992; Knot & Nelson, 1998; Perez et al., 2001; Wellman
et al., 2002). Ni2+ (50 µM) a selective CaV3.2 blocker, reduced Ca2+ spark event frequency in
pressurized vessels by 50%, consistent with regulation of RyR activity by CaV3.2 channels. We
conclude that CaV3.2 channels could indirectly drive BKCa channel activity via SR Ca2+ release
through RyR. To investigate how modulation of signalling between CaV3.2 and BKCa channels
could affect arterial function we performed additional myography experiments with Ni2+, a
CaV3.2 blocker, and paxilline, a BKCa channel blocker. Paxilline induced a similar augmentation
of tone as did Ni2+ blockade of CaV3.2, particularly at pressures below 80 mmHg. When Ni2+ was
first applied to the preparation, paxilline did not induce further tone, and conversely tissues pre-
treated with paxilline, were unaffected by Ni2+. These results are consistent with the idea that
these two channels are linked sequentially.
3.4.3 Summary.
In summary, we provide eveidence that Ca2+ influx through one subtype of T-type Ca2+
channels operates to facilitate vasodilation. More specifically, our results are consistent with the
interpretation that localized Ca2+ flux through CaV3.2 is sufficient, within a restricted spatial
domain, to activate RyR and sequentially elicit Ca2+ sparks, leading to BKCa channel activation
and arterial hyperpolarization and vasodilation. Our results suggest that this mechanism
attenuates myogenic tone in rat vascular cerebral arteries, the key constrictor response that sets
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basal tone and flow to the brain. This negative feedback response that counters constrictor
responses possibly extends to other critical organs such as the kidney where efferent arterioles
not only display myogenic tone but also express CaV3.2. This mechanism could also moderate
tone development in vascular beds under the control of the sympathetic nervous system, such as
the mesenteric circulation. We conclude that voltage-gated Ca2+ channels should not be viewed
as a simple influx pathway that directly elevates [Ca2+]i. Selected subtypes of VGCCs could
work in a localized manner to discretely regulate signaling pathways essential to setting the
contractile state of vascular smooth muscle.
3.4.4 Limitations and future directions.
The current research suggests a unique role for a VGCC in a negative feedback
mechanism. There are several concerns with the experiments presented in this chapter that need
to be considered however.
(i) Vessel myography approach.
Ni2+ was the only CaV3.2 blocker used in the studies of the specific role of CaV3.2 in
mediating BKCa channel activity due to the lack of other specific blockers that can differentiate
between the two subtypes of T-type Ca2+ channels that we found to be present in cerebral arteries
(Lee et al., 1999; Perez-Reyes, 2003). Ni2+ blocks CaV3.2 channels not only by binding within
the permeation pathway of the pore, as with all T-type Ca2+ channels, but also by binding to an
extracellular histidine amino acid near S4 (voltage sensor), which allosterically affects the
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channel and hence the current. This explains the high binding specificity of Ni2+ to CaV3.2
channels.
Thus the results presented in this chapter could not be confirmed by using additional
CaV3.2 blockers as there are no better specific blockers of CaV3.2 available. The availability of
more selective blockers would facilitate further differentiation and understanding of the
functional roles of CaV3.2 in vascular smooth muscle. As an alternative to the use of more non-
specific blockers, the use of siRNA or/and knockout (KO) mice (see Chapter 4) for CaV3.2 could
be applied. An siRNA approach as discussed in Chapter 2 could be used to selectively block the
expression of CaV3.2 in artery segments in culture. Arteries treated with siRNA and from control
arteries (treated with no siRNA or treated with scrambled siRNA) could be compared in vessel
myography experiments in which vessel diameter is measured as a function of increasing
intravascular pressure. If CaV3.2 functions as described above and has a vasodilatory influence,
then in the siRNA treated vessels, increasing pressure should lead to greater myogenic tone than
in vessels not treated with siRNA or in those treated with a scrambled siRNA. Similarly vessels
from cerebral arteries of CaV3.2 KO mice would be expected to develop greater myogenic tone
than those from control mice if CaV3.2 has the same function in mice as that proposed here for
rat cerebral arteries. The KO approach is discussed further in Chapter 4.
(ii) Structural approach.
In order to investigate the functional role of CaV3.2 VGCCs in the plasmalemma and
their relationship with RyRs in the SR, several different structural approaches were used.
Immunohistochemistry using whole mounted vessels showed that both CaV3.2 and RyR have
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overlapping distributions that indicate a possible relationship between them. However, several
kinds of experiments could be done to strengthen the immunohistochemistry data. For example
immunohistochemistry experiments could be carried out using isolated smooth muscle cells and
with a higher power microscope objective to better visualize the labelling in individual cells.
Primary antibodies to CaV1.2, CaV3.1 (see supplemental Figure 27), CaV3.2, and RyR could be
used to study the presence and distributions of these channels, it would also be of interest to
examine the distribution of other relevant proteins such as caveolin, BKCa (see supplemental
Figure 28) and IP3R (see supplemental Figure 29). The distributions of antibody labelling could
confirm our previous results but could also reveal distributions of the channels that were
different or more extensive than was seen when we labelled whole mounted vessels.
Experiments using isolated cells could help to rule out the possibility that the labeling seen in the
whole arteries was due to other cell types present in the arteries (e.g. endothelial cells or
perivascular nerve cells).
Dual labeling experiments (both standard and immunofluorescence and PLA
experiments) could be carried out using isolated single cells and higher magnification.
Antibodies to CaV3.2 and caveolin could be used to double label cells to further test for the
presence of CaV3.2 channels in caveolae. Moreover, a dual labelling experiment and
conventional immunofluorescence could be carried out using antibodies to CaV3.2 and RyR to
also examine possible co-localization of the two channels. Co-localization of CaV3.2 and RyR in
these experiments would strengthen our evidence of a possible relationship between the
channels. Other dual labelling experiments could be done using antibodies to CaV3.1 and CaV3.2,
CaV1.2 and RyR, or CaV1.2 and caveolin. Results of dual labeling experiments that showed co-
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localization of CaV3.2 with caveolin combined with results showing co-localization of CaV3.2
with RyR would support the hypothesis that Ca2+ influx through CaV3.2 could trigger Ca2+
release through RyR. If we found co-localization of CaV1.2 and caveolin it might suggest that
CaV3.2 is not the only VGCC that could influence RyR function. From our hypothesis we might
expect not to see co-localization of CaV3.1 with CaV3.2 or of CaV3.1 with caveolin. However, if
we do see some co-localization, this might indicate additional functions for the two channel
types. Furthermore, triple labelling for different channels could also be done (e.g. CaV3.2, RyR,
caveolin; CaV3.1, RyR and caveolin; CaV3.1, CaV3.2 and caveolin; CaV1.2, RyR, and caveolin)
and could provide more information about the possible function of these channels in arterial
smooth muscle.
A PLA assay was used to determine if CaV3.2 channels and ryanodine receptors existed
in cells in close proximity to each other. Results of the assay indicate that there are populations
of both channels in close proximity to each other (≤ 40 nm). PLA results did not show discrete
punctate signals as it might have been expected to however. The manufacturers of the assay
(Olink Bioscience) suggest that this may be because of high expression levels of the studied
proteins. This could cause a high amount of labelling and lead to blurred images (Olink
Bioscience).
Electron tomography, using a series of images generated with a transmission electron
microscope, was used to provide a picture of a possible arrangement of the cell plasma
membrane and near membrane intracellular structures. Our results suggest the presence of a
microdomain structure in vascular smooth muscle cells. In these experiments cerebral arteries
were isolated, fixed in paraformaldehyde/glutaraldehyde, dehydrated and then embedded in Epon
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resin prior to being cut into thick sections for analysis. It is possible that the preparation of the
sample introduces artefacts in the structures actually present in the cells of the arteries. Therefore
the electron tomography experiments could possibly be enhanced by using frozen-hydrated
samples (Cryo-EM), where samples are rapidly frozen at a very low temperature (near -196 οC)
so that water becomes vitrified and not crystallized inside the tissue and cells; this could give
better preservation of structures within the cells, and the cells will be less prone to damage by the
electron beam of the electron microscope.
Electron microscopy using immunogold labelled secondary antibodies and primary
antibodies to CaV3.2 and to RyR was used to look at the distribution of the two proteins in thin
sections of cerebral arteries. My results indicated that some CaV3.2 channels are localized in the
plasma membrane in caveolae and in close proximity to RyR in the SR and that about one third
of the labelling of CaV3.2 and RyR was associated with caveolae-like or SR-like structures,
respectively.
. The specificity of the primary antibodies was tested in competition experiments in
which the antibodies were pre-incubated with peptide antigens. However, further work could be
done to investigate the distributions of CaV3.2 and RyR.
To investigate the possibility that the distributions of CaV3.2 and RyR labelling that we
saw could be due to random labelling by the two antibodies, an EM image could be artificially
(via computer) labelled with random distributions for the two antibodies using the same amounts
of labelling as measured experimentally. Different size points could be used to represent the
immunogold labelling by the two antibodies and the degree of co-localized points and the
association of points with caveolae and SR structures could be determined. These experiments
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would provide an estimate of the apparent co-localization or association with caveolae and SR in
the same areas that would be expected to occur due to random labelling by the antibodies. If the
average of the results of repeated random distribution experiments showed the same degree of
co-localization that was observed in the electron microscopy experiments this would argue
against the idea that CaV3.2 and RyR are co-localized. Similarly if the random distribution
experiments showed the same degree of association of points with caveolae- or SR-like structure
as were observed in EM experiments this would argue against the idea that the proteins are
localized in areas where the SR and caveolae are in close proximity.
Co-immunoprecipitaion (Co-IP) is a technique used to investigate protein-protein
interactions, and could be used to investigate the presence of CaV3.2 and proteins associated with
CaV3.2. The presence of caveolin in the Co-IP samples would be a strong indication that CaV3.2
could be found in caveolae. Secondly, the co-localization of Cav3.2 and RyR could be examined
by fluorescence resonance energy transfer (FRET), where the two proteins of interest are labeled
with different fluorochromes, such that the emission spectrum of one overlaps the absorption
spectrum of the other. If the two fluorochromes are within a certain distance of each other (∼ 10
nm) a FRET signal would be observed.
(iii) STOCs and Ca2+ sparks.
STOCS were measured by using perforated patch clamp at both - 40 and -20 mV for
control experiments, and in the presence of Ni2+ and paxilline. The limitation of using Ni2+ as the
only blocker available has been highlighted before. Meanwhile, paxilline was used as BKCa
channel blocker in the study to compare its effect to the effect of Ni2+. The BKCa channel
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blocker, iberiotoxin, a very highly selective inhibitor of BK channels (more selective than
paxilline; Galvez et al., 1990) could be used and compared to the results of paxilline to see if
there is any difference in the blocking between the two blockers. Furthermore, STOCS could be
measured at different voltage ranges (e.g. from -50 mV to +10 mV) to monitor their production
in control cells and in the presence of CaV3.2 blockers (Ni2+) and BKCa blockers (paxilline and/
or iberiotoxin) to test if there is any contribution by other channels in these voltage ranges.
Ca2+ spark production was measured using the fluorescent Ca2+ dye fluo-4 and confocal
microscopy. The dissociation constant (Kd) for Ca2+ binding to fluo-4 is 0.345 nM (Life
Technologies, 2010) and the fluorescence of the dye increases when the dye binds Ca2+. One
concern with the use of fluo-4 to measure Ca2+ sparks is that the dynamic range of the dye might
not be great enough to accurately measure the magnitude of some Ca2+ sparks. Recently,
Figueroa et al., 2013 adapted this method to include the simultaneous use of two fluorescent
dyes (fluo-4 and fluo-4FF) to measure Ca2+ sparks. Fluo-4FF has the same spectral properties as
fluo-4 but the Kd for Ca2+ binding to the dye is 9.7 μM (Life Technologies, 2010). The use of
the two dyes together increases the range of magnitudes of Ca2+ sparks that can be measured
(Figueroa et al., 2013).
Moreover, the limitations of using Ni2+ as CaV3.2 channel blockers could be overcome by
using a siRNA approach or by using CaV3.2 channel knockout mice to measure STOCs and Ca2+
sparks in control and treated vessels or knockout mice. The use of knockout mice is further
discussed in Chapter 4.
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(iv) Computational approach.
The mathematical model that has been constructed for Ca2+ handling in vascular smooth
muscle cells of cerebral arteries was used to test the possibility, on a theoretical level, that T-type
VGCC subtype CaV3.2 and RyR Ca2+ channels are able to induce CICR, hence Ca2+ spark
activity. The model could be further developed with information about channel distribution and
channel density, once this information is available from the results of future experiments.
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3.5 Figures.
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Figure 16: Effect of Ni2+ on pressure-induced tone and membrane potential (Vm) in rat
cerebral arteries. A&B) Middle and posterior cerebral arteries were pressurized from 20-100
mmHg while diameter was monitored under control conditions, in the presence of the CaV3.2
blocker Ni2+ (50 µM), and in Ca2+-free medium. Representative traces (A) and summary data (B,
n = 7 arteries from 6 rats) reveal the constricting effects of Ni2+. Arterial Vm in pressurized
cerebral arteries (60 mmHg) in the absence (C) and presence of 50 µM Ni2+ (D). Representative
traces are shown in (C) and (D). Summary data is shown in (E), where n = 6 arteries from 6 rats.
* denotes significant difference from control; P value ≤0.05 (paired t-test, and one way ANOVA
(panel (A) and panel (B) performed by Rasha Abd El-Rahman and panels (C-D) by Suzanne
Welsh).
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Figure 17: Ca2+ channel localization in rat cerebral arteries. Cerebral arteries were labeled
with antibodies against smooth muscle actin (green) and CaV3.2/RyR/CaV1.2 (red). In (A, top),
CaV3.2 and RyR labeling (arrowheads) run perpendicular to the long axis of smooth muscle cells
(long white arrow) in regions devoid of actin. In (17A, bottom, the same as 17A upper) smooth
muscle actin (green) and CaV3.2/RyR (red) are presented in individual gallery panels. In (B),
CaV1.2 labeling (arrowheads) runs parallel to the long axis of smooth muscle cells (long white
arrow). Photomicrographs are representative of 3 individual experiments (Performed by Rasha
Abd El-Rahman).
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Figure 18: Controls of smooth muscle actin, CaV3.2, RyR, and CaV1.2. Cerebral arteries were
labeled with antibodies against smooth muscle actin and CaV3.2/RyR/CaV1.2. Secondary
antibody and peptide control experiments were negative for labeling of smooth muscle actin and
CaV3.2/RyR/CaV1.2 in panels A, B, and C. In (D) cerebral arteries were labeled with antibodies
against neurofilament protein (purple) in perivascular nerves that are embedded in the vascular
smooth muscle cell layer. Secondary antibody control experiments are where both primary
antibodies are absent. Peptide control experiments utilized a peptide that blocks the specific
epitope of the primary antibody. Peptides were added to primary antibodies overnight at a
dilution 1: 1, then added to tissue samples. Photomicrographs are representative of 3 individual
experiments (Performed by Rasha Abd El-Rahman).
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Figure 19: Proximity ligation assay and the colocalization of CaV3.2 and RyR in rat
cerebral arterial smooth muscle cells. Isolated smooth muscle cells labeled with primary
antibodies against CaV3.2 and RyR were subjected to a proximity ligation assay. A-C) A gallery
representation, where (A) is differential interference contrast (DIC), (B) nuclei labeled with
Hoechst 33342 blue stain, where (C) reveals the presence of red fluorescent product consistent
with CaV3.2 and RyR localizing within 40 nm of one another. For orientation purposes, cell
nuclei were labeled with blue stain Hoechst 33342 blue. D-F) Proximity ligation assay in the
absence of both primary antibodies but in the presence of both secondary antibodies, where (D)
is DIC, (E) is the stained nuclei, and (F) shows no red fluorescent product. G-I) Proximity
ligation assay in the absence of one primary antibody (no CaV3.2 antibody only RyR antibody)
but in the presence of both secondary antibodies, where (G) is DIC, (H) is the stained nuclei, and
(I) shows no red fluorescent product. J-L) Proximity ligation assay in the absence of one primary
antibody (no RyR antibody only CaV3.2 antibody) but in the presence of both secondary
antibodies, where (J) is DIC, (K) is the stained nuclei, and (L) shows no red fluorescent product.
Scale bars are 10 μm and the optical section depth in each image is 0.3 – 0.5 μm.
Photomicrographs are representative of ~20 smooth muscle cells in 2-3 cell preparations
(Performed by Rasha Abd El-Rahman).
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Figure 20: Electron microscopic imaging of rat cerebral arterial smooth muscle cells. A-C)
3-D tomographic imaging was performed on posterior cerebral arteries to ascertain the presence
of microdomains. In (A), tomography performed on a 300 nm thick section of tissue to generate
a contiguous stack of 2-D photomicrographs with ~3.5 nm resolution. Subcellular structures
were traced on each section. In (B) and (C), these tracings were compiled to produce 3-D
renditions. Note that discrete membranous regions can be observed where caveolae and SR
come in close apposition to one another. Photomicrographs are representative of 3 different
arterial preparations. D-F) Immunogold labeling of RyR and CaV3.2 channels in rat cerebral
arteries. In (D), RyR labeling (red arrows) can be observed underneath the plasma membrane
near caveolae. Boxed area in top panel was magnified and placed underneath. In (E), CaV3.2
labeling (red arrows) is observed near the plasma membrane in association with caveolae (the
boxed area was magnified in the panel beneath it). In (F) control experiment (no primary
antibody) reveals an absence of electron dense particles (the boxed area was magnified in the
panel beneath it). This photomicrograph is representative of 3 different arterial preparations
(Performed by Rasha Abd El-Rahman).
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Figure 21: A microdomain model of smooth muscle Ca2+ dynamics. Using tomographic,
immunolabeling and electrophysiological data, along with mathematical models of channel
kinetics, a microdomain model of Ca2+ dynamics has been created. In (A), the model consists of
an 8.5 µm slice of an arterial smooth muscle cell. The microdomain is 15 nm in length and is
comprised of SR and a subspace of appropriate volume. Membrane proteins have been
distributed accordingly and the level of expression set by mathematical optimization procedures.
Key proteins included were CaV1.2, CaV3.1, CaV3.2, RyR, Na+-Ca2+ exchanger (NCX), SERCA
& PMCA pumps, calmodulin and calsequestrin. With this model, Ca2+ concentrations can be
calculated in the SR, the subspace and the cytosol. In (B), simulations were performed whereby
voltage was stepped from -60 to -40 mV to induce repetitive Ca2+ spark-like events. The
frequency of these Ca2+ spark-like events was eliminated or reduced upon blockade of RyR (C)
or CaV3.2 channels (D), respectively, and increased with depolarization (E) (simulation
programming by Kamran Bigdely-Shamloo and figures by Rasha Abd El-Rahman).
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Figure 22: Spontaneous transient outward currents (STOCs) and Ca2+ spark production in
cerebral arterial smooth muscle cells. A & B) Perforated patch clamp electrophysiology was
used to measure STOCs under control conditions and in the presence of Ni2+ (50 µM) (panel A)
and paxilline (1 µM) (panel B). Representative traces and summary data (n = 8 cells) can be
found in (A) and (B). * denotes significant difference from control; P < 0.05 (paired t-test, and
one way ANOVA). C & D) Posterior and middle rat cerebral arteries were loaded with Fluo-4
and line scan imaging performed to investigate the generation of Ca2+ sparks under control
conditions and in the presence of Ni2+ (50 µM). Representative line scan (arrows denote Ca2+
sparks) and summary data can be found in (C) and (D), respectively. * denotes significant
difference from control; P < 0.05 (paired t-test, and one way ANOVA). Experiments shown in
panels (A) and (B) were performed by Osama Harraz and Rasha Abd El-Rahman. Cell isolation
and preparation was performed by Rasha R. Abd El-Rahman, experiments shown in panels (C)
and (D) were performed by Sean Wilson and Monica Rubalcava.
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Figure 23: Effect of Ni2+ and paxilline on myogenic tone in rat cerebral arteries. A-D) Rat
cerebral arteries were pressurized from 20-100 mmHg while diameter was monitored under
control conditions and in the presence of Ni2+ (50 µM, n = 6 arteries), paxilline (1 µM, n = 6
arteries) and Ca2+-free medium. Representative traces (A & C) and summary data (B & D) reveal
the constricting effects of Ni2+ and paxilline, where the red bars represent the difference between
the control and the addition of either Ni2+ or paxilline. * denotes significant difference from
control. E) Rat cerebral arteries were pressurized to 60 mmHg and then sequentially exposed to
Ni2+ (50 µM, n = 7) and then paxilline (1 µM, n = 6) or in the other order (G). F & H) bar graphs
demonstrate the change in diameters at different intravascular pressures (20 - 60 mmHg), where
at 60 mmHg no further change of diameter was observed when paxilline was added on top of
Ni2+ (F), or vice versa (H). * denotes significant difference from 60 mmHg; P value ≤ 0.05
(paired T-test, and one way ANOVA) (Performed by Rasha Abd El-Rahman).
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Chapter Four: DISCUSSION
4.1 Overview.
As discussed above, in vascular smooth muscle L-type voltage-gated Ca2+ channels
(CaV1.2) are believed to be the major channel regulating Ca2+ entry, subsequent contraction and
development of arterial tone (Bannister et al., 2009; Davis & Hill, 1999; Harder, 1984; Knot &
Nelson, 1998; Moosmang et al., 2003). Other studies have demonstrated, however, that other
VGCCs are present in smooth muscle cells of different vascular tissues (Braunstein et al., 2009;
Kuo et al., 2010; Nikitina et al., 2007; Oguri et al., 2010; Abd El-Rahman et al., 2013; 2009;
Feng et al., 2004; Navarro-Gonzalez et al., 2009). Although LVA Ca2+ channel currents has been
measured in resistance arteries with properties consistent with T-type channels (i.e. fast
activation/inactivation, and slow deactivation), additional information is necessary to determine
their role. It has been suggested that T-type channels function in cell proliferation and cell cycle
regulation (Oguri et al., 2010; Perez-Reyes, 2003; Rodman et al., 2005). Other investigators
have suggested that T-type Ca2+ channels play a role in myogenic tone development (Perez-
Reyes, 2003; Rodman et al., 2005). This exact role is controversial and varies between the
investigators according to the type of tissues they studied.
Evidence has been presented suggesting that Ca2+ influx through some T-type channels
facilitates arterial constriction (CaV3.1), while influx through other T-type channels (CaV3.2)
tends to dilate arteries (Braunstein et al., 2009; Chen et al., 2003; Hansen et al., 2011; Kuo et al.,
2010; Poulsen et al., 2011). With regard to the potential involvement of CaV3.2 channels in
vasodilation, some investigators have theorized that these channels might be targeted to small
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cellular domains, where they might be involved in the regulation of the activity of BKCa channels
(Chen et al., 2003).
The overall goal of this thesis was to develop a mechanistic understanding of the function
of L- and T-type VGCCs in the generation and regulation of myogenic tone in vascular smooth
muscle cells. The two main objectives were: 1) identification of the type(s) of VGCCs expressed
in vascular smooth muscle cells and their roles in the generation of myogenic tone; and 2)
determination of the specific function of one of the T-type Ca2+ channels, CaV3.2, in regulating
myogenic tone in cerebral arteries.
4.2 Objective #1: Identification of L- and T-type Ca2+ channel subtypes in rat cerebral
arterial smooth muscle cells and their roles in myogenic tone development.
Our study (Chapter 2) revealed the presence of mRNAs and all three α1-subunit proteins
of CaV1.2 (L-type) and CaV3.1/3.2 (T-type) in rat cerebral arteries (Abd El-Rahman et al., 2013),
consistent with the findings of Kuo et al. (Kuo et al., 2010). We used Ba2+ as the conducting ion
rather than Ca2+ in order to enhance the whole cell current amplitude (Abd El-Rahman et al.,
2013) and a pharmacological approach to distinguish between L- and T-type channels (Abd El-
Rahman et al., 2013). We found that nifedipine reduced the whole-cell current and the remaining
current showed a hyperpolarized shift characteristic of T-type Ca2+ channels (Iftinca & Zamponi,
2009; Perez-Reyes, 2003). This result is consistent with the findings of Nikitina et al. (Nikitina et
al., 2007) and Kuo et al. (Kuo et al., 2010). We found that T-type Ca2+ channel blockers
(mibefradil, efonidipine, and kurtoxin) reduced the nifedipine-insensitive current as expected for
T-type current. Furthermore, the activation/inactivation time constants (τ) for T-type currents
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were smaller (indicating faster activation and inactivation) than those of the L-type current,
which agreed well with previously published values for CaV3.1/3.2 and CaV1.2 (Kuo et al., 2010;
Perez-Reyes, 2003). Functional measurements were also carried out in which cerebral arteries
were pressurized and the changes in diameter of arteries were observed in control conditions and
in the presence of L-type Ca2+ channel blocker (nifedipine) and T-type Ca2+ channel blockers
(mibefradil, kurtoxin and efonidipine). The functional contribution of L-type Ca2+ channels to
arterial tone increased with pressurization, as cerebral arteries became more depolarized (-35
mV, ~80 mmHg); (Abd El-Rahman et al., 2013). This finding is consistent with the biophysical
properties of the L-type channel (Bannister et al., 2009; Knot & Nelson, 1998). Results of these
experiments are consistent with the interpretation that the contribution of T-type Ca2+ channels to
development of myogenic tone is greater when the vessel is hyperpolarized (-55 mV, ~20
mmHg) (Abd El-Rahman et al., 2013; Kuo et al., 2010; Nikitina et al., 2007; Perez-Reyes,
2003). Computational modeling of the arterial network showed that a slight change of ~ 5 to
15% in arterial diameter, similar to that observed with T-type Ca2+ channel blockade, could
possibly lead to an increase in network perfusion of ~ 20 –50%. Such results indicate that T-type
channels could have a significant effect on blood flow (Abd El-Rahman et al., 2013).
In summary, the main conclusions are: (i) both L-type (CaV1.2) and T-type (CaV3.1 and
CaV3.2) Ca2+ channels are expressed in smooth muscle cells from middle/posterior cerebral
arteries; (ii) T-type Ca2+ channels contribute to induction of the myogenic response in
hyperpolarized vessels.
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4.3 Objective #2: CaV3.2 and the induction of a negative feedback response in cerebral
arterial smooth muscle.
Based on our observation that two subtypes of T-type Ca2+ channels (CaV3.1, CaV3.2) are
present in rat cerebral artery smooth muscle cells and the suggestion that T-type channels may
mediate vasoconstriction or vasodilation we were motivated to try to separate the contribution of
these two channel types to vascular tone. In agreement with Chen et al. (Chen et al., 2003) we
observed the constriction of pressurized rat cerebral arteries upon application of the CaV3.2
blocker Ni2+. Ni2+ has an IC50 for CaV3.2 of 12 µM, and was used at a concentration (50 µM) that
would block ~ 98% of those channels, without affecting CaV3.1 or CaV1.2 (Table 1; Perez-
Reyes, 2003).
Our structural results are consistent with the presence of discrete microdomains
consisting of caveolae and SR, in which CaV3.2 and RyR, respectively, reside, which indicate
that CaV3.2 could mediate a localized Ca2+ influx rather than a global rise in [Ca2+]i.
Furthermore, computational modeling predicted, on a theoretical level, that Ca2+ influx through
T-type Ca2+ channels could possibly elicit repetitive Ca2+-induced Ca2+ release events. These in
turn could induce Ca2+ spark-like events through the activation of RyR, which initiate the
activation of large conductance Ca2+-activated K+ channels (BKCa), and hence elicit
hyperpolarization in cerebral arteries (Perez et al., 2001; Perez et al., 1999). This mechanism in
part agrees with that proposed by Kotlikoff (Kotlikoff, 2003) and Ji et al. (Ji et al., 2006), who
reported CICR in smooth muscle cells, but as “loose coupling” events (Collier et al., 2000;
Kotlikoff, 2003), where RyR opening is tied to Ca2+ channel activity in the sarcolemma.
However, CaV1.2 was proposed as the Ca2+ source mediating rises in [Ca2+]i (Collier et al.,
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2000). The biophysical properties of T-type Ca2+ channels (CaV3.2) that activate at lower
voltages, and that rapidly inactivate, suggest that they could be the source of Ca2+ that induces
CICR events. Our model predicts a complete blockade of Ca2+ sparks by blocking RyR channels
as expected. The model predicts a ~ 50 - 60% attenuation upon CaV3.2 channel blockade.
Moreover, direct measurements of spontaneous transient outward currents (STOCs) using the
perforated patch technique showed ~ 60% reduction or elimination of STOCs in the presence of
Ni2+ (CaV3.2 blocker) or paxilline (BKCa blocker). In addition, about a 50% reduction of Ca2+
spark events occurred upon the addition of Ni2+. In functional studies in which cerebral artery
diameter was measured, we found that the addition of paxilline after Ni2+ could not further
increase tone. Similarly, the addition of Ni2+ after paxilline caused no further increase in tone.
These results support and highlight the idea that these two channels are linked serially in a single
sequential pathway. These results support the hypothesis that BKCa channels are regulated by
CaV3.2 indirectly through RyR, and consequently elicit vasodilation in cerebral arteries. Our
proposed feedback mechanism is somewhat analogous to that of Earley et al., who suggested that
Ca2+ spark generation and subsequently BKCa channel activity in cerebral arterial smooth muscle
is triggered by TRPV4 channel activation (Earley et al., 2005). However, in our proposed
scenario, CaV3.2 is the main channel involved, which is compatible with its biophysical
properties of fast activation and inactivation at low voltages, and slower deactivation.
In summary, the main conclusion from Objective #2 is that CaV3.2 voltage-gated Ca2+
channels elicit vasodilation by mediating the influx of Ca2+ into a restricted microdomain to
activate RyR and sequentially cause Ca2+ sparks, which in turn activate BKCa channels and
induce arterial hyperpolarization (Figure 24).
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Figure 24: Diagram highlighting the proposed function of CaV3.2 channels in the
regulation of arterial tone. We propose that Ca2+ influx through CaV3.2 channels mediates a
dilatory response by activating ryanodine receptors (RyR) and initiating Ca2+ sparks, localized
events that activate large conductance Ca2+-activated K+ (BKCa) channels (made by Rasha Abd
El-Rahman).
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4.4 Future directions, experimental considerations, clinical relevance, and conclusion.
This thesis addressed crucial questions concerning the expression and functional role of
VGCCs in cerebral arterial smooth muscle cells. This study suggests that CaV3.1 mediates global
increases in intracellular calcium as suggested in other studies (Braunstein et al., 2009; Hansen et
al., 2011; Kuo et al., 2010; Poulsen et al., 2011) while CaV3.2 may work to alter [Ca2+]i in a
localized manner, as opposed to mediating global increases in [Ca2+]i.
In the preceding chapters additional experiments were proposed to address experimental
concerns and limitations. The future directions section below describes experiments that could be
carried out to expand on the results presented in Chapters 2 and 3. This section is followed by a
discussion of the clinical relevance of the results and a conclusion.
4.4.1 Future directions.
An additional experiment that could be added to the myography experiments in Chapters
2 and 3 would be to measure arterial diameter in the presence of an L-type channel blocker (e.g.
nifedipine) plus Ni2+ (to block CaV3.2 channels as described in Chapter 3) and to follow this by
adding one of the T-type channel blockers used in Chapter 2 (i.e. mibefradil, kurtoxin,
efonidipine). This should allow any effects of the blockers on CaV3.1 and other channels that
might be present in the preparations to be determined in the absence of the contribution of
CaV3.2 channels.
Our work could also be extended to investigate the functional role of CaV3.1 and CaV3.2
in mesenteric and pulmonary arteries and renal arterioles. Both channels have been identified in
these tissues (Braunstein et al., 2009; Hansen et al., 2011; Pluteanu & Cribbs., 2011; Poulsen et
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al., 2011). This would determine if CaV3.1 and CaV3.2 have similar roles as vasoconstrictors or
vasodilators in other vascular beds. This would also be of interest because mesenteric arteries
undergo phasic contractions compared to the tonic contraction seen in cerebral vascular and renal
arteries. The experimental approaches described in Chapters 2 and 3 (e.g. electrophysiological,
functional and structural approaches) could be applied in these studies to investigate the types of
inward Ca2+ currents in these arteries and to examine the possible expression of different T-type
channel subtypes, and their role in the development of myogenic tone. Careful experiments
might make it possible to differentiate between different splice variants of CaV3.1 and CaV3.2
and their biophysical characteristics (conductivity, channel gating, and window current
properties) in these arteries and to understand how the expression of different subtypes might be
reflected in their functional role/s.
Vessel myography could be used to study rat cerebral and other arteries that have not
been denuded of endothelium to investigate the influence that endothelial cells (e.g. through the
release of endothelium-derived hyperpolarizing factors (EDHF)) might have on the development
of myogenic tone in intact blood vessels. This work may also provide information about how the
endothelium could affect the negative feedback that we proposed might occur through the
interaction of CaV3.2 and BKCa channels in cerebral arteries.
Along with a study of the effect of the presence of the endothelium on the myogenic
response of the cerebral and other arteries discussed above and the experiments suggested above,
another line of research could be followed to investigate the presence of CaV3.1 and CaV3.2 in
endothelium. The endothelium of mesenteric and cerebral arteries expresses mRNA for CaV3.1
and CaV3.2 Ca2+ channels, whereas pulmonary micro-vascular endothelial cells express CaV3.1
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(Townsley et al., 2006; Zhou & Wu, 2006). This work could lead to further information about
the mechanism involved in the control or limitation of vasoconstriction in smooth muscle
(Harraz & Welsh, 2013; Zhou et al., 2010; Kuo et al., 2010). The influx of Ca2+ through the T-
type Ca2+ channels into the cytoplasm of the endothelial cells could activate the production of
EDHFs such as nitric oxide (NO) which would induce vasodilation of smooth muscle cells.
These proposed experiments could involve PCR, western blots, immunohistochemistry, and
electrophysiology. It should be noted, however, that patch-clamp studies of endothelial cells
from a variety of vessels failed to find any evidence for T-type Ca2+ channel current (Braunstein
et al., 2009; Kuo et al., 2010). The data available in this field of study is still incomplete,
therefore more work needs to be done, including electrophysiological and structural studies to
determine if T-type channels are present and if they have a functional role in endothelial cells.
The possible involvement of CaV3.1 Ca2+ channels in the regulation of other channels,
such as TRP channels (e.g. TRPM4 channels) would be of interest. TRP channels are involved in
membrane depolarization, activating L-type Ca2+ channels and hence they induce
vasoconstriction in vascular smooth muscle (Figures 25 and 26). TRPM4 channels are permeable
to Na2+ causing membrane depolarization; at the same time they are known to be activated by
Ca2+ release from IP3Rs (Earley et al., 2004; Gonzales et al., 2010a). CaV3.1 could activate IP3Rs
and the Ca2+ released from the SR could then elicit transient inward cation currents (TICCs) or it
could activate chloride (Cl-) channels (e.g. TMEM16A) that induce spontaneous transient inward
currents (STIC) (Harraz & Welsh, 2013). Both channels (TRPM4 and TMEM16A) can cause
membrane depolarization and hence activation of L-type VGCCs that in turn cause a global rise
in [Ca2+]i and constriction (Earley et al., 2004; Gonzales et al., 2010a) (Figures 25 and 26).
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Therefore, CaV3.1 could induce vasoconstriction while CaV3.2 could induce vasodilation.
Application of experimental approaches (patch-clamp, vessel myography) using blockers of
TRPM4 and TMEM16A (9-phenanthrol and T16Ainh-A01, respectively) could be used to
investigate the contribution of these channels to recorded currents and to investigate their role in
myogenic tone development. Structural approaches (e.g. immunogold-labelling, PLA, and
immunofluorescence) could also be used to determine the localization and possible co-
localization of TRPM4 and TMEM16A with other proteins including CaV3.1 and CaV3.2.
It would also be of interest to study the possibility of signaling complexes among T-type
VGCCs (especially CaV3.1) and members of the potassium channels superfamily (e.g. KV4
channels) to regulate the function of the latter channels. Recently it was reported that CaV3-
mediated calcium influx in cerebellar stellate cells assists in the control of the voltage-
dependence of KV4 inactivation at a nano-domain level (Turner et al., 2011). Study of this
possibility in vascular beds could thus uncover a similar nano-domain Ca2+ signaling mechanism
in these beds.
T-type Ca2+ channels might also initiate and/or maintain Ca2+ waves, asynchronous
events dependent on Ca2+ release from the SR (Braunstein et al., 2009; Martin-Cano et al.,
2009). These Ca2+ wave events are more essential in setting myogenic tone at hyperpolarized
rather than depolarized membrane potentials (Mufti et al., 2010) which might be in consistent
with T-type Ca2+ channel involvement. Using confocal microscopy to study the production of
Ca2+ waves in rat cerebral arteries under control conditions, and in the presence of T-type Ca2+
channels blockers (mibefradil, kurtoxin, efonidipine, and Ni2+) would produce information about
possible roles for CaV3.1 and CaV3.2 in Ca2+ wave production.
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- Knockout mice.
A longer term approach would be to perform experiments with knockout mice which are
already commercially available for both CaV3.1 and CaV3.2 (Chen et al., 2003; Stamboulian et
al., 2004). The knockout gene technique involves the replacement of a normal gene with a
mutant gene using a homologous recombination technique that causes either deletion or
inactivation of the normal gene of interest; the gene is said to have suffered a knockout (Zan et
al., 2003). As noted CaV3.1 and CaV3.2 knockout mice have been made, however, the vascular
smooth muscle in these animals has not been studied to any great extent. Although an abnormal
vasodilatory phenotype is observed for the aorta in CaV3.2 knockout mice, their blood pressure is
normal. Examination of CaV3.1 knockout mice indicates that the channel contributes to sinoatrial
node pacemaker activity and atrioventricular conduction (Mangoni et al., 2006), but no work has
been done in the vascular smooth muscle in these animals. To extend the present studies to the
knockouts, it would first be necessary to repeat many of the experiments that were described in
Chapters 2 and 3 on normal mice to determine if regulation of the cerebral arteries in them is
similar to that seen in rats. Depending upon the results with normal mice, experiments could then
be designed to compare normal and knockout mice. If, for example, evidence (e.g. mRNA and/or
presence of channel proteins) was obtained consistent with the presence of both L- and T-type
Ca2+ channels in the cerebral vasculature in mice, then electrophysiological and myography
experiments could be undertaken to determine the effects of the knockout. Experiments designed
to examine the localization and co-localization of these channels with other proteins of interest
could reveal structural changes in smooth muscle cells that develop as a result of the knockout of
one or both of the T-type channels.
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To illustrate some possible experiments that could be done with KO mice. I will make the
assumption that mice and rats have the same complements of VGCCs. If this actually turns out to
be the case, there are several types of experiments that could be done using knockout mice.
Cerebral artery smooth muscle cells from normal and CaV3.2 KO mice could be used in
electrophysiological experiments in which STOCs are measured. If CaV3.2 functions the way we
concluded in Chapter 3, then characteristics of STOCs (e.g. frequency) seen in normal mice
might not be the same as those seen in cerebral arterial smooth muscle cells from CaV3.2 KO
mice.
The effect of Ni2+ (at a concentration that blocks only CaV3.2) could be compared in
cerebral arteries from normal and CaV3.2 KO mice in vessel myography experiments. Ni2+ would
be expected to cause constriction in vessels from normal mice based on our conclusions from the
experiments reported in Chapter 3. In contrast, Ni2+ would be predicted to have no effect in the
cerebral arteries from CaV3.2 KO mice.
Vessel myography experiments, similar to those reported in Chapter 2, could be carried
out using cerebral arteries from normal and CaV3.2 KO mice in order to examine the effects of
T-type channel blockers. Based on the experiments reported in Chapters 2 and 3, a greater degree
of relaxation might be expected to occur in vessels from CaV3.2 KO mice in response to T-type
blockers (e.g. mibefradil, kurtoxin, and efonidipine). In vessels from normal mice, blocking
CaV3.1 channels would have a vasoconstricting effect while blocking CaV3.2 channels would
have a vasodilatory effect. As we showed in Chapter 2 the net effect of the blockers, based on the
predominant role of CaV3.1 channels, is to cause vasodilation. In vessels from CaV3.2 KO mice
only the vasodilatory effect would occur therefore the degree of relaxation might be expected to
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be greater. The results of these experiments could possibly be complicated by differences in the
resting vessel diameters and possible differences in CaV3.1 expression level between normal and
CaV3.2 KO mice.
Ca2+ sparks and Ca2+ waves could be characterized in cerebral artery smooth muscle cells
from normal, CaV3.1 KO mice and CaV3.2 KO mice, to see if Ca2+ waves occur and if their
properties vary between cells from different types of mice. The properties of the waves could
include frequency, amplitude, rate of propagation and distance travelled through the cell.
Vessel myography experiments using cerebral arteries from normal, CaV3.1 KO mice and
CaV3.2 KO mice could be compared using vessels with the endothelium present in order to
understand the role of T-type VGCCs in smooth muscle and endothelial cells. However, these
experiments might be extremely complicated by the possible presence of both CaV3.1 and
CaV3.2 in smooth muscle and endothelial cells in the normal mice.
In smooth muscle cells from both CaV3.1 and CaV3.2 KO mice the inward current
blocked by T-type channel blockers (e.g. mibefradil, kurtoxin, and efonidipine) would be
expected to be less than that blocked in cells from normal mice. If this result was not seen in
experiments then that might indicate that the blockers have non-specific effects.
In considering the use of knockout mice, however, one must be aware of some possible
limitations/difficulties with the technique. Knockout mice are made by using a technique in
which a DNA fragment containing a desired mutant gene is inserted into a vector and introduced
into a special line of embryo-derived mouse stem cells (ES) by homologous recombination. If
the mutant gene prevents the expression of the protein coded by the normal gene a knockout is
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produced. In using this technology, there are several possible problems and limitations that must
be taken into consideration. 1) In about 15% of knockout mice the knockout is either embryonic
lethal or the mice do not grow to adulthood, so the effects of the affected protein cannot be
studied in adult animals. 2) Certain genes are difficult to knockout. 3) Knocking out some genes
will affect the expression of other genes such that a specific characteristic phenotypic cannot be
distinguished. 4) In experiments using knockouts another potential problem is that cells from two
different animals (normal and knockout) are compared. 5) A major issue for the use of knockout
technology is that readily available knockout models are developed in mice. Thus results
obtained from the use of knockouts may not be readily applicable to other animals. This can be a
problem especially if work on another animal model has been extensive and it appears that the
tissues and biological systems differ significantly between these models and the mouse model. 6)
It is still an expensive approach because of the high maintenance costs needed for the mice.
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Figure 25: Diagram highlighting the different proposed possible functions of CaV3.2
channels in the regulation of arterial tone. We propose that Ca2+ influx through CaV3.2
channels mediates a dilatory response by activating ryanodine receptors (RyR) and initiating
Ca2+ sparks, localized events that activate large conductance Ca2+-activated K+ (BKCa) channels.
CaV3.2 could possibly activate TRP channels (e.g. the TRPM4 channels) to elicit transient
inward cation currents (TICCs) (made by Rasha Abd El-Rahman).
.
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Figure 26: Diagram highlighting the possible separate functions of CaV3.1 and CaV3.2
channels in the regulation of arterial tone. Ca2+ influx through CaV3.2 channels mediates a
dilatory response by activating ryanodine receptors (RyR) and initiating Ca2+ sparks, localized
events that activate large conductance Ca2+-activated K+ (BKCa) channels. Furthermore, a role for
CaV3.1 in activating Cl- channels (possibly TMEM16A channels) may be to induce spontaneous
transient inward currents (STIC) or perhaps TRP channels (e.g. TRPM4) to elicit transient
inward cation currents (TICCs) (made by Rasha Abd El-Rahman).
.
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4.4.2 Clinical relevance.
The work presented in my thesis enhanced our current understanding of the L- and T-type
voltage-gated Ca2+ channel function in cerebral arterial smooth muscle. We have proposed a
unique functional role for one of the T-type Ca2+ channels (CaV3.2) in cerebral arteries. As
discussed above there are some limitations to our experiments (the lack of specific blockers for
T-type VGCCs, the use of Ba2+ as the charge carrier when recording the inward Ca2+ current, and
the limitations of the different structural experiments) and I have suggested additional
experiments that could be done. Nevertheless, my investigation has a significant impact on
understanding the functional role of smooth muscle VGCCs in normal cerebral arteries. This new
understanding may help in comprehending the molecular basis of the function of cerebral arteries
in pathological conditions. A major pathological condition is hypertension and VGCCs are one
of the main targets in the treatment of hypertension (Godfraind et al, 1986; Belardetti, et al,
2008; Hansen et al, 2011).
Hypertension is a chronic medical condition in which blood pressure is elevated in the
arteries, hence augmenting vascular tone (Carretero & Oparil, 2000a, 2000b). Therefore,
blocking T-type Ca2+ channels per se may not be effective at treating hypertension, because at
least one of the T-type channels functions in a manner different from L-type channels, the latter
contributing to constriction while CaV3.2 contributes to dilation. To target the CaV1.2 or CaV3.2
separately, drugs that are selective in blocking CaV3.2 or CaV3.1, and CaV1.2, will have to be
developed. It may be that CaV3.2 is down-regulated in hypertension leading to the reduction in
Ca2+ spark generation. It has been noted that Ca2+ sparks are decreased in frequency in
hypertensive rat arterial cells (Amberg et al., 2003); this could be because CaV3.2 expression is
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decreased or channel regulation is altered (possibly by a mutation in the protein). Further work in
this area will be important in examining these possibilities.
4.4.3 Conclusion.
The work described in this thesis used a combination of innovative methods to
investigate the basis of Ca2+ dynamics and arterial tone regulation in vascular smooth muscle
cells. Collectively, three voltage-gated Ca2+ channels have been confirmed to be present in rat
cerebral arterial smooth muscle. Molecular and electrophysiological approaches showed that
both L-type (CaV1.2) and T-type (CaV3.1 and CaV3.2) Ca2+ channels are expressed and
electrically distinctive in cerebral arterial smooth muscle cells. This work also provided evidence
that T-type Ca2+ channels contribute to myogenic tone generation, where L-type (CaV1.2)
showed the greater contribution and T-types the lesser contribution. Furthermore, our findings
point to a unique role for a VGCC (CaV3.2) in driving local Ca2+ events (Ca2+ sparks), through a
CICR process, to regulate arterial tone in vascular smooth muscle cells. Therefore, understanding
of VGCCs should take into consideration the different roles the channels play. Furthermore, our
findings suggest that CaV3.1, identified in rat cerebral arteries, could be responsible for the
modest contribution of T-type Ca2+ channels to myogenic tone. This channel could play a more
prominent role in increasing [Ca2+]i at a more hyperpolarized state of the vessels and therefore
assists in initiating basal myogenic tone.
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APPENDIX
Figure 27. CaV3.1 protein expression in vascular smooth muscle cells.
Figure 28. BKCa protein expression in vascular smooth muscle cells.
Figure 29. IP3R protein expression in vascular smooth muscle cells.
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Figure 27: CaV3.1 protein expression in vascular smooth muscle cells. Cerebral arteries were
labeled with antibodies against smooth muscle actin (green) and CaV3.1 (red). In (A), CaV3.1
labeling (arrowheads) runs parallel to the long axis of smooth muscle cells (long white arrow). In
(B), smooth muscle actin (green) and CaV3.1 (red) are presented in individual panels. Secondary
antibody and peptide control experiments were negative for labeling (C). Secondary antibody
control experiments were done with both primary antibodies absent. Peptide experiment utilized
a peptide that blocks the specific epitope of the primary antibody, and was added to primary
antibodies overnight at a dilution 1 : 1, then added to tissue samples. Photomicrographs are
representative of 3 individual experiments (Performed by Rasha Abd El-Rahman).
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Figure 28: BKCa protein expression in vascular smooth muscle cells. Cerebral arteries were
labeled with antibodies against smooth muscle actin (green) and BKCa (red). In (A), BKCa
labeling (arrowheads) runs parallel to the long axis of smooth muscle cells (long white arrow). In
(B), smooth muscle actin (green) and BKCa (red) are presented in individual panels. Secondary
and peptide control experiments were negative for labeling (C). Secondary antibody control
experiments were done with both primary antibodies absent. Peptide experiment utilized a
peptide that blocks the specific epitope of the primary antibody, and was added to primary
antibodies overnight at a dilution 1 : 1, then added to tissue samples. Photomicrographs are
representative of 3 individual experiments (Performed by Rasha Abd El-Rahman).
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Figure 29: IP3R protein expression in vascular smooth muscle cells. Cerebral arteries were
labeled with antibodies against smooth muscle actin (green) and IP3R (red). In (A), IP3R labeling
(arrowheads) runs parallel to the long axis of smooth muscle cells (long white arrow). In (B),
smooth muscle actin (green) and IP3R (red) are presented in individual panels. Secondary and
peptide control experiments were negative for labeling (C). Secondary antibody control
experiments were done with both primary antibodies absent. Peptide experiment utilized a
peptide that blocks the specific epitope of the primary antibody, and was added to primary
antibodies overnight at a dilution 1 : 1, then added to tissue samples. Photomicrographs are
representative of 3 individual experiments (Performed by Rasha Abd El-Rahman).
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