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This is to certify that the
dissertation entitled

T-TYPE CALCIUM CURRENTS IN SPINAL SENSORY
INTEGRATION

presented by

Wen-hsin Ku

has been accepted towards fulfillment
of the requirements for the

PhD. degree in Physiology

 

 

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T-TYPE CALCIUM CURRENTS IN SPINAL SENSORY INTEGRATION
By

Wen-hsin Ku

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY
Physiology

2010

ABSTRACT
T-TYPE CALCIUM CURRENTS IN SPINAL SENSORY INTEGRATION
By
Wen-hsin Ku

The spinal cord’s dorsal horn receives sensory inputs encoded by primary sensory
neurons that innervate the body and internal organs and is the origin of ascending
pathways toward the brainstem and thalamus. Neurons in this region exhibit distinct
intrinsic properties that are important for the integration of diverse sensory information.
In this study, I focused on T-type Ca2+ currents (IT) and their role on the intrinsic
properties of these neurons. Whole-cell patch-clamp recordings were performed on dorsal
horn neurons in a slice preparation obtained from the lumbosacral spinal cord of hamster
pups (P9—P14). Cells were examined under both voltage- and current-clamp conditions,
and IT was isolated pharmacologically. Two T-type Ca2+ channel blockers Ni2+ and
mibefradil were applied extracellularly in some experiments to verify the involvement of
IT in the intrinsic properties. Single cell reverse transcription polymerase chain reaction
(RT-PCR) was performed after patch-clamp recordings in an attempt to uncover the
molecular identity of IT in individual dorsal horn neurons. The data show that IT recorded
in dorsal horn neurons had voltage-dependent activation and inactivation and similar
sensitivity to the blockers, as did the T currents recorded in other neurons. Although IT
was expressed prevalently in the dorsal horn, its amplitude and kinetics varied widely
from cell to cell. IT had the highest current density in a group of phasic firing cells that
are preferentially activated by rapid membrane depolarization resulting from direct

current injection or fast moving skin stimuli. These cells also had the strongest expression

of a fast inactivating IT component (ITJ‘) revealed by an exponential stripping procedure.
For both IT and IT; , their current amplitude was correlated with the slope of
depolarizing membrane ramps. This rate-dependent activation may underlie the ability of
dorsal horn neurons to sense time-varying stimuli on the Skin. IT expression was
associated with a rebound discharge after a hyperpolarization; further analysis indicates
both Inf and a slowly inactivating IT component (1T,s) contribute to this postinhibitory
rebound behavior. Single-cell RT-PCR experiments indicate that 1T,f is mediated by the
Cav3.1 channel subtype. To summarize, my study confirms the presence of T-type Ca2+
currents in the spinal dorsal horn and suggests that this low-voltage-activated current is
involved in differential capabilities of dorsal horn neurons to distinguish between static

and moving skin stimuli by the mechanism of rate-dependent current activation.

To my parents

iv

 

ACKNOWLEDGEMENTS .:

I would like to thank my committee, Drs. William Atchison, Steven Heidemann,
David Kreulen, and Birgit Zipser, for their guidance from constructing the proposal to
completing this dissertation. I am especially gratefiil to my mentor, Dr. Stephen 1
Schneider, for his sensible advice, kindly support, and constant patience throughout these
years. I would like to give a big thank you to Tracy Walker for her warm friendship and
help both inside and outside the lab. I also want to extend thanks to Dr. Wenling Zhang
and Timothy Houchin for providing a friendly and supportive working environment.

The single cell RT-PCR experiments cannot be done without the assistance from a
number of individuals, and I would like to thank: Dr. Xiaoling Dai and Christine Harman
for sharing valuable experiences and protocols, Dr. Laura McCabe for her counsel, Drs.
Edward Perez-Reyes (University of Virginia) and Ravindra K. Hajela for providing
materials for pilot experiments, and colleagues from Kreulen and Weber labs for
technical support. Also, this study has been made possible through the National Institute
of Neurological Disorders and Stroke grand NSZS771.

Pursuing this degree has been quite a journey, and I am glad and truly thankful to

have family and friends along the way, supporting me and encouraging me.

TABLE OF CONTENTS

LIST OF TABLES .................................................................................... ix
LIST OF FIGURES .................................................................................. x
ABBREVIATIONS ................................................................................. xii
CHAPTER ONE
INTRODUCTION .................................................................................... l
A. The role of the spinal dorsal horn in somatic sensory system ......................... 2
B. Intrinsic properties of spinal dorsal horn neurons and their ionic mechanisms. . ...4
C. T-type calcium channels .................................................................... 7
Discovery ..................................................................................... 7
Electrophysiology ........................................................................... 8
Molecular characterization ................................................................ 10
T—type calcium current in spinal dorsal horn neurons ................................. 13
D. Single cell reverse transcription polymerase chain reaction in combination with
patch clamp recording .............................................. i ....................... 15
Specific Aims .................................................................................... 17
CHAPTER TWO
T-TYPE CALCIUM CHANNELS CONTRIBUTE TO SIGNALING OF MEMBRANE
DEPOLARIZATION RATE BY SPINAL DORSAL HORN NEURONS .................. 18
Abstract .......................................................................................... 19
Introduction ...................................................................................... 20
Methods .......................................................................................... 23
Slice preparation ........................................................................... 23
Recording procedures ..................................................................... 24
Drugs and chemicals ....................................................................... 26
Statistical analyses ......................................................................... 26
Results ............................................................................................ 27
T-type calcium currents in hamster dorsal horn neurons .............................. 27
Rate-dependent activation of T-type calcium currents in spinal dorsal horn
neurons ....................................................................................... 32
T-type calcium currents in spinal dorsal horn neurons with identified membrane
and discharge properties .................................................................. 34
Discussion ....................................................................................... 44
T-type calcium currents and postinhibitory rebound (PIR) ........................... 46

vi

 

Ionic basis of phasic dorsal horn neuron responses to rapid membrane

depolarizations .............................................................................. 47
Functional significance .................................................................... 50
CHAPTER THREE

A FAST INACTIVATING T-TYPE CALCIUM CURRENT IS INVOLVED IN
SENSING MEMBRANE DEPOLARIZATION RATE BY SPINAL DORSAL HORN

NEURONS ........................................................................................... 52
Introduction ...................................................................................... 53
Methods .......................................................................................... 55

Slice preparation ........................................................................... 55
Electrophysiological recording ........................................................... 56
Drugs and chemicals ....................................................................... 58
Single cell reverse transcription polymerase chain reaction (RT-PCR) ............ 58
Statistics ..................................................................................... 62
Results ............................................................................................ 63
Fast and Slow T-type calcium currents in spinal dorsal horn neurons ............... 63
Rate-dependent activation of fast T-type calcium currents in spinal dorsal horn
neurons ....................................................................................... 67
Expression of fast and slow T-type calcium currents in spinal dorsal horn neurons
with identified membrane and discharge properties ................................... 70
Expression of Cav3 subunits in spinal dorsal horn neurons exhibiting fast and
slow T-type calcium currents ............................................................. 75
Discussion ....................................................................................... 78
Fast and slow T-type calcium currents .................................................. 78
Molecular identity of fast and slow T-type calcium channels ........................ 80
Concluding remarks ........................................................................ 82

CHAPTER FOUR

SUMMARY AND DISCUSSION ................................................................ 83
A. Summary of experiments .................................................................. 84
B. Technical issues in single cell RT-PCR experiments ................................. 85
C. T-type calcium currents in spinal sensory integration... ....................................... 88

IT in lowering the threshold for firing ................................................... 88
IT potentiation .............................................................................. 91
IT de-inactivation ........................................................................... 92
IT activation ................................................................................. 95
IT in synaptic function ..................................................................... 97
D. Remarks on the sensory functions of the spinal dorsal horn ......................... 99
E. Future studies .............................................................................. 101
vii

APPENDIX
SUPPLEMENTAL DATA ....................................................................... 103

REFERENCES .................................................................................... 1 12

viii

LIST OF TABLES
Table 2-1. Comparison of 1T and intrinsic membrane properties of spinal dorsal horn
neurons in this study ................................................................................ 36

Table 3-1. Comparison of 1T,f and IT,s in spinal dorsal horn neurons with different firing
pattems...................... .......................................................................... 73

Table 3-2. Summary of the expression of T-type calcium channels in twelve spinal dorsal
horn neurons .......................................................................................... 76

Table A-1. Comparison of action potentials from neurons in slices prepared using
sucrose- vs. glycerol-based dissection solution ................................................ 104

Table A-2. Comparison of inward currents recorded from neurons in slices prepared
using sucrose- vs. glycerol-based dissection solution ......................................... 105

Table A-3. Comparison of IT from neurons in superficial vs. deep spinal dorsal horn...106

Table A-4. A contingency table showing the relationship between the use of silanized
pipettes and the detection of CaV3 mRNA in single cell RT-PCR experiments .......... 107

Table A-5. Comparison of IT and intrinsic membrane properties in Spinal dorsal horn
neurons with and without the capability of generating ‘low-threshold-current-mediated
action potentials’ (LTAPs) ........................................................................ 108

Table A-6. A contingency table Showing the relationship between the exhibition of

postinhibitory rebound (PIR) and the ability to generate low-threshold-current-mediated
action potentials (LTAPs) in twenty-eight spinal dorsal horn neurons ..................... 109

ix

 

LIST OF FIGURES
Figure 1-1. A structural model of T-type Ca2+ channels ....................................... 12
Figure 2-1. Isolation of low-voltage-activated Ca2+ currents ................................. 28

Figure 2-2. Voltage-dependent activation and inactivation of T-type Ca2+ currents in

spinal dorsal horn neurons ......................................................................... 30
Figure 2-3. Effect of T-type Ca2+ channel blockers ................................ ~ ............ 31
Figure 2-4. Activation of IT in dorsal horn neurons with ramp depolarizations ............ 33
Figure 2-5. Firing patterns of hamster dorsal horn neurons .................................... 37
Figure 2-6. Transient depolarization in phasic cells ............................................ 39
Figure 2-7. Effect of Ni2+ and mibefradil on phasic firing .................................... 40
Figure 2-8. Postinhibitory rebound in hamster dorsal horn neurons ......................... 41
Figure 2-9. 1T in neurons with identified firing patterns... . ................................... 43

Figure 3-1. Analysis of T-type Ca2+ current components by using an exponential
stripping procedure ................................................................................. 59

Figure 3-2. Distribution of the decay time constants of IT current components in spinal
dorsal horn neurons ................................................................................. 64

Figure 3-3. Voltage-dependent activation and inactivation of Inf and 1T,s in spinal dorsal
horn neurons .......................................................................................... 66

. 2
Figure 3-4. Inhibition of IT; and 1T,s in spinal dorsal horn neurons by T-type Ca +

channel blockers ..................................................................................... 68

Figure 3-5. Rate sensitivity of IT,f in spinal dorsal horn neurons ............................. 69

Figure 3-6. Firing patterns and IT of spinal dorsal horn neurons ............................. 71
x

Figure 3-7. Comparison of IT; and IT,s in spinal dorsal horn neurons with and without
PIR. ................................................................................................... 74

Figure 3-8. Detection of Cav3.1 subunit mRN A from a spinal dorsal horn neuron
expressing IT with one [7; component .......................................................... 77

Figure 4-1. A hypothetical mechanism for dorsal horn neurons to distinguish the direction

of a moving skin stimulus .......................................................................... 94

Figure 4-2. T-type Ca2+ channels in spinal sensory integration .............................. 98

Figure A-l. Images of the dorsal horn and its neuron recorded in this study .............. 110

Figure A-2. Expression of IT in phasic- and tonic-firing neurons ........................... 111
xi

ABBREVIATIONS

4-AP, 4-arninopyridine

ACSF, artificial cerebrospinal fluid

ADP, afterdepolarization

AHP, afterhyperpolarization

D, delay cells

dV/dt, membrane depolarization slope

EPSP(S), excitatory postsynaptic potential(s)
GAPDH, glyceraldehyde 3-phosphate dehydrogenase
HVA, high-voltage-activated

INaP: persistent Na+ current

IPSP(s), inhibitory postsynaptic potential(s)

IT, T-type Ca2+ current,

1T,f, fast inactivating T-type Ca2+ current

1T,s, slowly inactivating T-type Ca2+ current

L, Rexed’s lamina

LTAP(S), low-threshold-current—mediated action potential(s)
LVA, low-voltage-activated

MIB, mibefradil

Non-PSB, non-short burst phasic cells

P, phasic cells

PSB, Short burst phasic cells

xii

, PIR, postinhibitory rebound

RA, rapidly adapting

RF(s), receptive field(s)

RT, reverse transcriptase or reverse transcription
RT-PCR, reverse transcription polymerase chain reaction
SA, Slowly adapting

T, tonic cells

T-channel(s), T-type Ca2+ channel(s)

T-current(s), T-type Ca2+ current(s)

TEA, tetraethylammonium chloride

TTX, tetrodotoxin

If, decay time constant of fast inactivating T-type Ca2+ current

. . . . 2+
rs, decay t1me constant of slowly Inactlvatmg T-type Ca current

xiii

CHAPTER ONE

INTRODUCTION

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A. The role of the spinal dorsal horn in somatic sensory system L‘:

The dorsal horn of the spinal cord receives terminations from primary sensory neurons
innervating the body and internal organs, and is the origin of pathways projecting to the
brainstem and thalamus. AS the first central relay of the somatic sensory system, the
dorsal horn plays an important role in integrating and processing the information
transmitted by peripheral sensory receptors. Histological Studies (Rexed, 1952; Molander
et al., 1984) Show that the dorsal horn is divided into Six layers or laminae (I-VI), which
have important functional implications. Lamina I neurons receive inputs carried by fine
afferent fibers, both myelinated (A5) and unmyelinated (C), and mainly respond to
thermal, noxious (painful), or both stimuli. Lamina II, also known as the substantia
gelatinosa, primarily receives its input from slowly conducting, usually unmyelinated,
fibers and contains nociceptive, thermoreceptive, and low-threshold mechanoreceptive
neurons; this part of the dorsal horn is mostly regarded for its role in pain processing.
Laminae I and II together are called the “superficial dorsal horn”. Laminae III-VI (the so-
called “deep dorsal horn”) consist of low-threshold, nociceptive specific (or high-
threshold), and wide dynamic range neurons that respond to a broad intensity range of
stimulation applied on primary afferent fibers. This area receives direct input from large,
myelinated (AB) fibers that carry signals encoded by muscle proprioceptors and a variety
Of cutaneous mechanoreceptors with distinct adaptation properties and differential
sensitivity to skin indentation and hair movement. Thus, the deep dorsal horn is directly
involved in integrating and processing mechanosensory information about the body
(Willis and Coggeshall, 2004).

Sensory information relayed at the dorsal horn is subsequently conveyed to higher

levels of the neuraxis, i. e. the brainstem and thalamus, through three major ascending
pathways. The spinothalamic pathway is common in all orders of vertebrates and most
highly developed in primates, including humans (Willis & Coggeshall, 2004). The cells
of origin of this pathway are located in laminae I and IV-VI of the spinal cord and project
their axons to the contralateral thalamus through the anterolateral system. These dorsal
horn neurons receive afferent fibers that innervate skin, muscle, and viscera, and are low-
threshold, high-threshold, or wide dynamic range cells based on the responses to
mechanical stimuli applied on their cutaneous receptive fields (Chung et al., 1979). Some
of the lamina I cells are therrnoreceptive and project to the medial thalamus (Craig &
Kniffki, 1985). The postsynaptic dorsal column pathway originates from neurons of the
spinal laminae III-V and ascends to the dorsal column nuclei through the dorsal funiculus
(Rustioni & Kaufman, 1977; Brown & F yffe, 1981). These dorsal horn neurons often
receive convergent inputs from skin, muscle, and joint afferents and respond to both
innocuous and noxious mechanical stimuli (Brown et al., 1983). The spinocervico-
thalamic pathway originates from laminae III and IV of the spinal cord, relays in the
lateral cervical nucleus by way of the dorsolateral funiculus, and then ascends
contralaterally to the thalamus. Dorsal horn neurons that project through this pathway can
be excited by stimulation on cutaneous afferent fibers of all sizes, as well as high-
threshold muscle and joint afferent fibers, and are primarily nociceptive based on their
responses to skin stimuli (Brown, 1973). This pathway provides a rapidly conducting
system for transmission of information from the skin to the cerebral cortex, especially in
cats (Catalano & Lamarche, 1957; Mark & Steiner, 1958). Despite a lot being known

about primary afferent inputs and ascending pathways, how diverse sensory information

is integrated by the spinal cord remains unclear. By studying the intrinsic properties of

dorsal horn neurons, their role in somatosensory integration may be better understood.

B. Intrinsic properties of spinal dorsal horn neurons and their ionic mechanisms
Based on their axonal projections, dorsal horn neurons are divided into two groups.
Projection cells have axons that extend outside the spinal cord and ascend through the
pathways mentioned above, while intemeurons have propriospinal axons that ramify
locally within the cell’s dendritic tree in some cells and are further extensive
longitudinally in others. Intemeurons outnumber projection cells by a large margin and
make up maximally 97% of the population (Willis & Coggeshall, 2004). As a major
element of spinal local circuits, dorsal horn intemeurons play an important role in spinal
sensory processing and have diverse intrinsic properties (Russo & Hounsgaard, 1999).
Among them, the difference in firing patterns is a much noticeable characteristic and has
been described in several studies (Thomson et al., 1989; Lopez-Garcia & King, 1994;
Jiang et al., 1995; Russo & Hounsgaard, 1996a, b; Hochman et al., 1997; Grudt & Perl,
2002; Prescott & De Koninck, 2002; Ruscheweyh & Sandkiihler, 2002; Schneider, 2003;
Graham et al., 2004). Two types of firing patterns have been commonly documented in
dorsal horn neurons even when different animal preparations and recording
configurations were used (Thomson et a1. , 1989; Lopez-Garcia & King, 1994; Hochman
et al., 1997; Grudt & Perl, 2002; Prescott & De Koninck, 2002; Ruscheweyh &
Sandkiihler, 2002; Schneider, 2003; Graham et al., 2004). Tonic cells fire continuously
throughout the duration of current stimulation, whereas phasic cells respond with only a

Short period of discharge. The degree of spike frequency adaptation, a decrease in firing

 

 

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frequency during sustained current injection, is markedly different for both cell
categories; tonic cells typically have weak spike frequency adaptation (small changes in
frequency with time) while the discharge frequency of phasic cells decreases much more
rapidly. In some studies, a third group of neurons, called ‘delay cells’, are reported as
responding to depolarizing current steps with an irregular or accelerating firing following
an apparent delay to the first action potential in both superficial (Grudt & Per], 2002;
Prescott & De Koninck,. 2002; Ruscheweyh & Sandkiihler, 2002; Graham et al. , 2004)
and deep dorsal horn (Hochman et al., 1997; Schneider, 2003). This type is relatively
fewer in number than tonic and phasic cells, especially in deep laminae (Hochman et al. ,
1997; Ruscheweyh & Sandkfihler, 2002; Schneider, 2003).

In the deep dorsal horn, phasic cells are found to be preferentially activated by
fast membrane depolarization (> 13—34 mV/s) with the number and frequency of the
action potentials positively correlating with the slope of stimulating current ramps,
whereas the discharge frequency of tonic cells is unrelated to current trajectory
(Schneider, 2003). In a hemisected spinal cord-skin patch preparation, phasic cells
selectively respond to rapid brush flicks applied on their cutaneous receptive field and
have strongest excitation in response to Skin displacement that exceeds 8 urn/ms. Tonic
cells, in contrast, respond to both moving and stationary cutaneous stimulation and are
best activated by static skin indentation or contact (Schneider, 2005). Both studies
(Schneider, 2003, 2005) suggest that these two cell types possess differential capabilities
of signal processing. A similar concept is also reported in lamina I neurons: tonic cells
appear to linearly transduce stimulus intensity into firing frequency and can act as

“integrators” (see next paragraph for its ionic mechanisms), whereas phasic cells, with

rapid spike frequency adaptation, are able to follow high frequency stimulus and function
as “coincidence detectors” (Prescott & De Koninck, 2002). In addition, based on their
responses to Skin stimuli, tonic cells appear to be activated by a wide range of mechanical
stimulation, suggesting a mixture of low and high threshold inputs, while phasic cells
likely receive sensory information from a rather narrow range of mechanoreceptors
(Lopez-Garcia & King, 1994; Schneider, 2005). Taken together, these data indicate a role
of different firing patterns in sensory information processing, and a better understanding
of these properties, e. g. the underlying ionic mechanisms, can further advance the
knowledge of sensory integration by dorsal horn neurons.

A few ionic mechanisms have been suggested to mediate different firing patterns
in the spinal dorsal born. A non-inactivating Na+ current that activates at subthreshold
potentials and has dendritic distribution (Safronov, 1999) is proposed to amplify and, by
working synergistically with a slowly-activating persistent Ca2+ current, prolong
synaptically-evoked depolarization in spinal lamina I neurons exhibiting tonic-firing. As
a result, these tonic cells are capable of encoding stimulus intensity through temporal
summation and act as integrators (Prescott & De Koninck, 2005). In lamina II of the
spinal cord, a basic pattern of tonic firing is generated by a fast activating and
inactivating voltage-gated Na+ current and a delayed-rectifier K+ current (Melnick et al. ,
2004b) while a lower expression Of the Na+ current contributes to the spike frequency
adaptation of phasic firing neurons (Melnick et al. , 2004a). In spinal lamina V neurons,
an accelerating discharge following a delayed onset is mediated by the slow inactivation
of a K+ current that is highly sensitive to 4-aminopyridine (4-AP) (Morisset & Nagy,

1998); on the other hand, the activation of a transient K+ current is suggested to underlie

 

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the delayed firing pattern in superficial dorsal horn neurons by other studies (Yoshimura
and Jessell, 1989; Ruscheweyh & Sandkiihler, 2002; Ruscheweyh et al., 2004). Both
high- and low-threshold-activated Ca2+ current are documented in dorsal horn neurons
(Huang, 1989; Ryu & Randic, 1990); among them, the T-type current is proposed to
mediate a burst firing following membrane hyperpolarization in a subpopulation of turtle
dorsal horn neurons (Russo & Huguenard, 1996a). In this study, the roles of this current
in the spinal dorsal horn were investigated, and some previous reports about T-type

current and critical to the present study are reviewed in the last part of the next section.

C. T-type calcium channels
Discovery

Coexistence of multiple Ca2+ conductances is first reported back in 1975 by
Hagiwara and colleagues, who performed two-microelectrode voltage-clamp recording
from starfish eggs. They noted that one of the two inward currents, channel I as they
called it, was activated and inactivated at more negative membrane potentials and
decayed faster, which was different from the properties of “typical”, presumably high-
voltage-activated (HVA), Ca2+ channels reported previously (Hagiwara et al., 1975).
Later in 1978, Moolenaar and Spector revealed a similar Ca2+ current in mouse
neuroblastoma cell line NIB-115 using a Na+-free external solution containing 15 mM
TEA and an elevated Ca2+ concentration of 10 mM. This Ca2+ current was activated at
-55 mV from a holding potential of -80 mV and reached the maximum magnitude at -20
mV with a time to peak of 8—10 ms. Its inactivation was also voltage-dependent and

virtually competed within 100—150 ms. Application of Co2+ (5-10 mM) reversibly

 

 

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abolished the current, which was otherwise TTX—resistant. Moolenaar and Spector’s work
is the first report of mammalian Iow-voltage-activated (LVA) channels, and the same
group confirmed the presence of this transient Ca2+ current later and found a second
Ca2+ current with a much longer inactivation time constant of 2000 ms in the same and
another cell line NG108-15 (Fishman & Spector, 1981). By mid 19805, the existence of
LVA CaZ+ current across various tissues had been firmly established thanks to the efforts
of many laboratories, including those led by Armstrong (Armstrong & Matteson, 1985),
Bean (Bean, 1985), Carbone and Lux (Carbone & Lux, 1984, 1987), Feltz (Bossu et al.,
1985), Kostyuk (Fedulova et al., 1985), and Tsien (Nilius et al., 1985; NoWycky et al.,
1985). In 1985, the term “T-type” was proposed by Nowycky, Fox, and Tsien for this
low-threshold current for its relatively “tiny” and “transient” unitary activity, while the
designators L and N were assigned to a “long-lasting” HVA current and the third
conductance that was “neither” T- nor L-type and found exclusively in “neurons”,
respectively. Since then, T-type Ca2+ channels have become the interest of an ever-
expanding research field. These channels have hitherto been implicated in the
mechanisms of cancer growth, cell development and proliferation, epilepsy, fertilization,
hormone secretion, mechanosensation, nociception, sleep, and vascular tone control
(Nilius et al., 2006) Three T-channel (1] subunits, am, am, and an or Cav3.1, Cav3.2,
and Cav3.3 in the current nomenclature (Ertel et al., 2000), were cloned by the end of the
903 (Cribbs et al., 1998; Perez-Reyes et al., 1998; Lee et al., 1999a), which enabled
systematic studies of channel distribution, as well as examination of their patho-

physiological roles using molecular tools (N ilius et al., 2006).

Electrophysiology

 

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T-type Ca2+ channels (T-channels) are widely expressed throughout the body and
exhibit a number of signature features in electrophysiology (Perez-Reyes, 2003), which
are described as follows. First, T-channels begin to Open afier small membrane
depolarizations (usually between -75 and -60 mV), a characteristic giving them the
name of “low-voltage-activated” channels. Second, the current mediated by these
channels is transient, even during a prolonged stimulus. Third, native T-channels have
tiny but equal single-channel conductance for Ba2+ and Ca2+; at physiological Ca2+
concentration (2 mM), the value is expected to be ~ 1 p8 (Talavera & Nilius, 2006).
Fourth, they close slowly when the membrane potential is repolarized from a depolarized
level, generating a slowly deactivating tail current. Fifth, the steady-state inactivation of
T-channels occurs in a voltage range that partially overlaps that of the activation (the
overlap usually between -70 mV and -40 mV), implicating the presence of a “window
current” (Crunelli et al. , 2005). Last, T-channels are much less sensitive to HVA channel
blockers, such as dihydropyridines, and can be identified and distinguished from HVA
channels by their sensitivity to Ni2+ and low concentration of mibefradil1 (Feltz, 2006),
both of which were used in my studies.

The distribution and expression of T-type Ca2+ current (T—current) in the body has
been carefully reviewed (Perez-Reyes, 2003; Yunker & McEnery, 2003), and

heterogeneity is observed in native LVA current across different tissues even under the

 

lMibefradil (Ro 40-5967), a tetralol derivative, was previously marketed as the
antihypertensive and antianginal drug Posicor but was voluntarily withdrawn by Roche in
early June 1998 due to its potentially fatal reactions with other drugs. Because of its
inhibitory effect on the liver enzymes cytochrome P450 2D6 and 3A4, mibefradil can
result in an increased plasma concentration of concomitantly administrated drugs to a
dangerous level to the body (SoRelle, 1998).

 

 

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same recording conditions (Yunker & McEnery, 2003). Differential expression of Cav3
subunits may partially contribute to these variations (Perez-Reyes, 2003; Yunker &
McEnery, 2003) Since distinct biophysical properties have been reported among three
Cav3 subunits (reviewed by Talavera & Nilius, 2006). Cav3.l, Cav3.2, and CaV3.3
isoforms show larger, smaller, or equivalent current amplitude, respectively, in 2 mM
Ca2+ than in isotonic Ba2+ concentration (McRory et al., 2001) though the underlying
mechanism is still uncertain (Talavera & Nilius, 2006). Recombinant Cav3.1 and CaV3.2
channels display similar activation and inactivation kinetics to native T—channels
(Talavera & Nilius, 2006), while the Cav3.3 subtype has slower dynamics with an
activation time constant of 30 ms (vs. 6‘8 ms for Cav3.l and 3.2) and an inactivation
time constant of 137 ms (vs. 30-47 ms for the other two) (Kldckner et al. , 1999; Chemin
et al., 2002). Ni2+ is especially efficacious for the current mediated by the CaV3.2
subunit, with the 1C50 of 6-12 PM in 10 mM Ba2+ vs. 167—250 IIM and 87-216 PM for
Cav3.l and Cav3.3, respectively (Lee et al., 1999b). Mibefradil doesn’t have
discriminative blockade on different Cav3 isoforms (ICSO = 1.1-1.5 IIM in 10 mM
Ba2+), but its affinity for the channels increases when Ca2+ is used as the charge carrier-
(Martin et al., 2000). Besides expression of different (1, subunits, the subtype variants
resulting from alternative slicing or single nucleotide polymorphism can also contribute
to the heterogeneity in native T-current (Yunker & McEnery, 2003; Perez-Reyes, 2006;
Perez-Reyes & Lory, 2006).
Molecular characterization

“In silica cloning”, cloning based on computer searches of genetic database, led to

the discovery of three T-type Ca2+ channel 01] subunits: Cav3.1/Ol] G (Perez-Reyes et al. ,

10

 

at

If:

En.

(J)
(I.

ha
air;

the

1998), Cav3.2/C1]H (Cribbs et al., 1998), and Cav3.3/Olu (Lee et al., 1999a). These pore-
forming subunits are comprised of a large polypeptide (> 200 kDa), as observed in HVA
Ca2+ and Na+ channels, and have four domains of six transmembrane segments (SI-S6),
a structure originally found in voltage-gated K+ channels. The S4 segment from
individual domains acts as the voltage sensor, P (pore)-loops between the SS and S6
segments form the selectivity filter, and the S6 segments surround the pore forming its
inner wall (Figure 1-1). The .Cav3 channel proteins show 70°/o—95% sequence
conservation among different species, but the homology within the channel family is only
40%, with the most highly conservative found in the membrane spanning regions (Perez-
Reyes, 2006).

As a result of in silico cloning, no T-channel-specific auxiliary subunits are reported.
Studies indicate that the 13 subunits of HVA Ca2+ channels are unlikely attached to the
LVA channels, and whether other (125 and Y subunits are associated with T-channels in
vivo is still left unanswered. Even if T—channels possess auxiliary subunits, they may not
act in similar ways as they do in HVA Ca2+ channels, e. g. regulating membrane
trafficking, current amplitude, and voltage-dependent gating (Arikkath & Campbell,
2003; Perez-Reyes, 2006).

Three T—channel genes have been identified: CA CNAIG, encoding CaV3.1; CA CNA 1H,
encoding Cav3.2; and CACNA 11, encoding CaV3.3, and they are 70—130 kb long with
35—38 exons. Alternative splicing occurs in all three genes, and most splicing variants
have tissue-specific expression and different kinetics and voltage dependence, which may
affect their ability to influence cell excitability. Some single nucleotide polymorphisms in

the coding region of CA CNA 1 H are found to generate Cav3.2 channels with different

11

Domain I II III IV

 

Extracellular
__.[% A, V A _—_.A. _

Intracellular 7V V V V V \_/' V C
N

Figure 1-1. A structural model of T-type Ca2+ channels

The protein has four domains (I-IV), each one consisting of six transmembrane segments
(from left to right: Sl-S6; shown as the cylinders) and the corresponding links. The S4
segments (labeled in gray shade) act as the voltage sensor, P-loops between the SS and S6
segments form the selectivity filter, and the S6 segments surround the pore and form its
inner wall. N and C indicate the amino- and carboxyl-terminus of the protein,
respectively.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

12

 

"'3
f\

(1.11
alt»:
tech
med
ans
COnd
mole
in xi;
mudc

anClll

gating properties and linked to absence epilepsy (Perez-Reyes & Lory, 2006).

T-type calcium current in spinal dorsal horn neurons

LVA (T-type) Ca2+ current has been reported to coexist with HVA Ca2+ current in the
spinal dorsal horn neurons (Huang, 1989; Ryu & Randic, 1990), consistent with the
findings in many other neuronal cells. The participation of Ca2+ conductances in the
firing of these cells is first suggested by Murase and Randic (1983). They identified two
distinct types of Ca2+-dependent potentials that were evoked by modest and strong
depolarizing current commands, respectively. The low-threshold Ca2+ electroresponsive-
ness was enhanced by hyperpolarized membrane potentials and showed fast inactivation
(Murase & Randic, 1983), similar to what had been reported in inferior olivary neurons
(Llinas & Yarom, 1981b, a). A transient low-threshold Ca2+ current was later recorded
along with two high-threshold components in dorsal horn neurons using voltage-clamp
technique (Huang, 1989; Ryu & Randic, 1990). The low-threshold current is presumably
mediated by T-type Ca2+ channels because of the activation and inactivation kinetics and
sensitivity to 200 — 400 MM Ni2+, whereas the high-threshold counterparts are likely
conducted by L- and N-type Ca2+ channels (Huang, 1989; Ryu & Randic, 1990). The
molecular compositions of T—channels are also examined in the spinal dorsal horn using
in situ hybridization (Talley et al., 1999). Cav3.l mRNA is present in all laminae at
moderate levels. The expression of Cav3.2 transcripts is primarin restricted to laminae I

and II while the distribution of Cav3.3 mRN A is more prominent in laminae III and IV.

The fimction of T-type Ca2+ current in the spinal dorsal horn is uncertain. The T-
current has been suggested to underlie a burst firing in response to an electrically- or

synaptically-evoked membrane hyperpolarization in a subpopulation of turtle dorsal horn

l3

 

CXII
duts
and.
35 D
TCKll':

SCDST

neurons, a phenomenon that may contribute to sensing the directionality of skin
stimulation (Russo & Hounsgaard, 1996a). Nevertheless, T-current has gotten more
attention recently because of its newly discovered role in peripheral and central pain
processing (Todorovic & Jevtovic-Todorovic, 2006). In the spinal dorsal horn, the
possible involvement of T-current in sensory transmission is indicated by both in vivo and
in vitro studies. Behavioral experiments showed that the analgesic effect mediated by u
opioid receptors at the spinal level was potentiated by the. T-channel blocker mibefradil,
(Dogrul et al., 2001). Also, T-current is found to be essential for the long-term
potentiation of synaptic activity evoked by nociceptive afferent fibers in lamina I
projection neurons, a possible cellular mechanism of central sensitization and
hyperalgesia (Ikeda et a1. , 2003). Matthew and Dickenson (2001) reported that the T-
channel blocker ethosuxirnide depressed both mechanically and thermally evoked
extracellular responses recorded from the spinal dorsal horn of anesthetized rats. In the
dorsal root ganglia, a large T-current is expressed in medium diameter sensory neurons
and carried by the Cav3.2 channel subtype. Although the functional identity of these cells
as D-hair mechanoreceptors (Heppenstall and Lewin, 2006) or nociceptors (Jevtovic-
Todorovic and Todorovic, 2006) is still controversial, an emerging role of T-current in

sensory transmission is clear.

T-type Ca2+ current recorded in the lateral geniculate nucleus is preferentially
activated by depolarizing voltage ramps with the slope exceeding 30 mV/sec (Crunelli et
al. , 1989), and the low threshold Ca2+ spikes mediated by this current are also only
evoked by the current ramps that produce a sufficient rate Of membrane depolarization

(Gutierrez et al. , 2001). This is a great interest of mine because phasic dorsal horn

14

 

pat

C01

6211

TC!

\1“.

neurons are selectively activated by fast current ramps and rapid skin movements
(Schneider, 2003, 2005). Given that T-type Ca2+ current has been reported in the dorsal
horn (Huang, 1989; Ryu & Randic, 1990), whether it is responsible for the rate sensitivity
of phasic cells was one of the questions that I wanted to answer in this study (Specific
Aim 1 and Chapter 2). Furthermore, since all three (11 subunits of T-channels are
expressed in the spinal dorsal horn (Talley et al. , 1999), the possible heterogeneity of T-

current was also investigated (Specific Aim 2 and Chapter 3).

D. Single cell reverse transcription polymerase chain reaction in combination with
patch clamp recording

The use of single cell reverse transcription polymerase chain reaction (RT-PCR)
combined with patch clamp recording has been performed to study single neurons since
early 19903 (Sucher & Deitcher, 1995). Generally, a whole-cell patch clamp recording is
performed on cells in culture or slice preparations, and before termination of the
recording, the cell content is collected by the electrode pipette. RNA is reverse
transcribed into cDNA, which is then amplified by PCR with primers specific for the
gene of interest. PCR product can be confirmed by electrophoresis and DNA sequencing
or used for a second round of PCR. As the development of fluorescence-based real-time
PCR techniques, the amplification can be detected while the reaction is still in progress,
also allowing relative or absolute quantification of targeted cDNA molecules (Liss &
Roeper, 2004). This experimental approach enables gene expression profiling of
individual cells that are anatomically and functionally identified, as well as simultaneous

detection of multiple genes from single cell samples (Phillips & Lipski, 2000). The

15

 

ch.
lPi

IC’C

Scr
lse

2115

061

CO]

uncertainty as to whether mRN A detected by single cell RT-PCR are translated into
functional proteins can be alleviated by performing whole-cell recording prior to cell
harvesting,'which provides functional assessment of membrane proteins, e.g. ion
channels, and indirectly correlates mRNA analysis with protein expression in single cells
(Phillips & Lipski, 2000). By combining single cell RT-PCR with whole-cell patch clamp
recording, Cav3 channels are found to mediate the LVA Ca2+ currents in spermatocytes
(Sakata et al., 2001), and Cav3.1 is identified as the T-channel subtype underlying a low-
threshold spike in hypothalamic paraventricular nucleus neurons (Lee et al. , 2008).
Spinal dorsal horn neurons are heterogeneous in anatomic locations, primary
sensory inputs (see Section A in this chapter), as well as electrophysiological properties
(see Section B in this chapter), which makes it appropriate, if not favorable, to study them
at single cell level. Whole—cell patch clamp recording was used in this study to examine
the roles of T-type Ca2+ current (IT) in the intrinsic firing properties of dofsal horn
neurons (Specific Aims 1 and 2), and single cell RT-PCR was performed following that to
confirm expression of the channels and to determine their molecular identity (Specific

Aim 3).

16

 

e
0.:

6X

EX:

IE;

att

Specific Aims
In this study, I tried to answer three questions:
1. Does IT contribute to the sensitivity of phasic-firing dorsal horn neurons to rapid
membrane depolarization?
2. Is IT in dorsal horn neurons composed of kinetically distinct current components?
So, which current component is responsible for the rate sensitivity of phasic cells?
3. What is the molecular identity of IT components in individual dorsal horn
neurons?

To answer these questions, I first re-examined the basic properties of IT in hamster
dorsal horn neurons from a spinal cord Slice preparation under voltage-clamp conditions.
The rate-dependent activation of IT, as previously reported in thalamic neurons (Crunelli
et al., 1989), was also investigated. These results are presented in Chapter 2, in which a
general description of IT expression in dorsal horn neurons is also included. The
experiments directly addressing Specific Aims 1 and 2 were performed by measuring IT
in the cells whose intrinsic firing properties were characterized in current-clamp mode. T-
channel blockers Ni2+ and mibefradil were added to the external media in some
experiments to verify the involvement of 1T in different properties. The analyses
regarding Specific Aim 1 are presented and elaborated in Chapter 2 while the data for
attending to Specific Aim 2 are Shown in Chapter 3. To address Specific Aim 3, single
cell RT-PCR experiments were conducted using the cell content from neurons that were
previously examined in patch clamp recordings, and the results are also shown in Chapter

3. Chapters 2 and 3 were both written in the format of journal articles.

17

CHAPTER TWO
T-TYPE CALCIUM CHANNELS CONTRIBUTE TO SIGNALING

OF MEMBRANE DEPOLARIZATION RATE BY SPINAL DORSAL
HORN NEURONS

18

5;

[11“.

C3
Ilt‘
d".

M"

Abstract

Spinal dorsal horn neurons differ in their responses to membrane depolarization rate and
time-varying afferent sensory inputs. Using whole-cell patch-clamp recordings in slice
preparations of hamster spinal cord, we investigated whether the transient, low-threshold
calcium current (1T) plays a role in generating rate-sensitive responses of dorsal horn
neurons to changes in membrane potential. IT was evoked by depolarizing voltage steps
and ramps having slopes exceeding 46 mV/s. The peak amplitude of IT activated by
voltage ramps increased with increasing ramp slope over the range of 240 mV/s to 2400
mV/s. The time-to-peak of IT decreased with increasing rate of membrane depolarization.
Bath application of 200 uM Ni2+ depressed the ramp-activated IT in a manner similar to
IT activated by step depolarizations. IT was widely, though not uniformly, expressed by
neurons in Rexed’s lamina I—IV. Peak amplitude of IT was correlated with the pattern of
firing evoked under current clamp conditions: IT in neurons responding to constant
membrane depolarization with one or two action potentials was 2.8- and 7-fold larger
than 11- in cells with tonic or delayed-firing patterns, respectively. Cells exhibiting a
postinhibitory, rebound excitation had significantly larger IT than those without. These
results are evidence that IT probably contributes to the ability of phasic-firing dorsal horn
neurons to signal rapid membrane depolarizations and thereby discriminate between

afferent sensory inputs that encode high- and low-frequency stimulus information.

19

 

 

lnl

Tn

116'

V1:

f.)

Introduction

The dorsal horn of the spinal cord is a structure of major importance for processing
sensory information from the body. It receives synaptic terminations of primary sensory
neurons that innervate a variety of tissues, including skin, muscle, tendons, joints and
viscera (Willis & Coggeshall, 2004). In this region, diverse sensory information is
integrated by local neural circuits, which are made from synaptic connections between
multiple, physiologically distinct types of intemeurons (Lu & Perl, 2003, 2005; Santos et
al., 2007; Schneider, 2008), and then relayed to segmental reflex circuitry and ascending
sensory pathways to the brain and brainstem.

Differences in electrophysiological properties of neurons can influence the
transmission of sensory information within dorsal horn circuits. Three or four cell types
have been identified in this region based on firing characteristics (Thomson et al., 1989;
Hochman et al., 1997; Prescott and De Koninck, 2002; Ruscheweyh and Sandkiihler,
2002; Schneider, 2003) and several studies have begun identifying the underlying ionic
conductances. Repetitive discharge of tonically firing cells has been reported to be
regulated by a combination of voltage-gated Na+ and KJr channels and persistent Ca2+
currents (Melnick et al., 2004b; Prescott and De Koninck, 2005). Other evidence suggests
that a transient, low-voltage-activated (T-type) Ca2+ conductance underlies a rapidly
adapting, “phasic” discharge Of other dorsal horn neurons (Russo and Hounsgaard,
1996a)

T-type Ca2+ channels are widely expressed in the central nervous system (Talley
et al., 1999; McKay et al., 2006; Nilius et al., 2006) where they promote burst firing,

. . . . . . . . . . 2+ .
Intrinsrc oscrllatory actrvrty, and partrcrpate In synaptrcally-evoked Ca Influx

20

 

pit

net
Sim

pha

Iha};
(Cry
oscH
fh}fl
chan.
dEpg;

38080

(Huguenard, 1996). Evidence of a Ca2+ current mediated by T-type channels has also
been reported in dorsal horn neurons (Huang, 1989; Ryu and Randic, 1990). Our interest
in the contribution of T-type channels to spinal somatosensory processing was prompted
by two observations. First, fast depolarizing current ramps activate a transient membrane
depolarization and discharge in many phasic-firing dorsal horn neurons, with the
depolarization amplitude and firing frequency being graded with the ramp slope
' (Schneider, 2003). Second, phasic cells in the dorsal horn region where cutaneous
mechanoreceptive afferents terminate respond selectively to mechanical stimuli that
produce rapid deformations in the skin (Schneider, 2005), suggesting that this class of
neurons may be part of specialized spinal circuits for encoding velocity information. A
similar observation has been made by Prescott and De Koninck (2002), who found that
phasic cells in dorsal horn lamina I are driven by high frequency stimulus trains and
argued that they function to detect simultaneous occurrence of separate afferent inputs. In
thalamic neurons, T-type currents are selectively activated by fast voltage ramps
(Crunelli et al., 1989) and generate large, transient Ca2+ depolarizations that contribute to
oscillatory behavior (Gutierrez et al., 2001). Given the importance of T-type channels to
rhythmic firing behavior in thalamic neurons, we considered the possibility that these
channels also shape responsiveness of dorsal horn neurons to dynamic membrane
depolarizations induced by direct current injection or synaptic activation by primary
sensory receptors.

In this study, we used whole-cell patch-clamp recordings in a rodent spinal cord
slice preparation to isolate and re-examine T-type Ca2+ currents of dorsal horn neurons.

We investigated activation of T-type currents by voltage ramps and compared current

21

amplitude to neuronal intrinsic membrane and firing properties characterized under
current clamp conditions. Our observations indicate that dorsal horn neurons with phasic
firing patterns express high T-type Ca2+ current density and provide evidence that a T-
type current is instrumental in tuning the selective responses of these cells to rapid
membrane depolarizations. Selective sensitivity of phasic-firing dorsal horn neurons to
fast depolarizations may underlie detection of transient afferent stimuli and encoding of

the stimulus velocity.

22

 

TE

SI;

TB]
110

frt

flu
Na

18”

Methods

All protocols involving the use of live animals were performed in accordance with the
National Institutes of Health Guide for the Care and Use of Laboratory Animals and were
approved by the Institutional Animal Care and Use Committee at Michigan State

University.

Slice preparation

Spinal cord tissue was obtained from 9- to-14-day-old Syrian hamsters of both sexes
under urethane anesthesia (1.5 mg/ g, intraperitoneal), followed by exsanguination. After
removing vertebrae and meninges, a block of lumbosacral spinal cord was glued to the
stage of a vibrating microtome (Vibratome 3000, St. Louis, MO) and 300-um-thick
transverse slices were made as previously described (Schneider, 2003). Dissection and
slicing steps were carried out at 4—8°C in a solution containing (mM) 179 sucrose, 2.5
KCl, 0.2 CaClz, 10 MgClz, 1.25 NaH2P04, 26 NaHCO3 and 25 glucose (pH 7.35—7.45,
290—310 mOsm/l) equilibrated with 95% 02-5% C02. In some experiments, sucrose was
replaced by equi-molar glycerol in the dissection solution (Ye et al., 2006). There were
no differences in resting potential or action potential amplitude and half-width recorded
from neurons in slices prepared using sucrose- or glycerol-based dissection solutions
(Table A-1 in Appendix). Slices were incubated in an oxygenated artificial cerebrospinal
fluid (ACSF) containing (mM) 120 NaCl, 2.5 KCl, 2.5 CaClz, 1.5 MgSO4, 1.25
NaH2P04, 26 NaHCO3, and 10 glucose (pH 7.35—7.45; 290—310 mOsm/l) at room

temperature for at least one hour prior to electrophysiological recording.

23

 

Ret‘or
For cl

mount

CORCI
IQCOI

16m p

band
iHun
Clam
(Fig!
nulli
COnf
comi

(MI)

Recording procedures

For electrophysiological recording, slices were transferred to a chamber (volume, 1.5 ml)
mounted on a fixed-stage microscope (Olympus BX51WI) and continuously perfused at
5—6 ml/min with ACSF bubbled with 95% 02—5% C02. Patch-pipette recording
electrodes with a resistance of 4—7 M!) were fabricated from borosilicate glass (N-51A,
Drummond Scientific, Broomall, PA). The internal pipette solution contained (mM) 130
K-gluconate, 5 NaCl, 1 CaClz, 1 MgClz, 11 EGTA, 10 HEPES, 2 Mg-ATP, and 0.1 Li-
GTP (pH 7.3, 280—285 mOsm/l). For isolating T-type Ca2+ currents under voltage-clamp,
we switched to a modified extracellular solution with the following constituents (mM):
107 NaCl, 3 BaClz, 2 CsCl, 1.3 MgClz, 26 NaHCO3, 10 glucose, 20 TBA, 0.5 uM TTX,
and 10 uM CdClg (pH 7.3 5—7.45; 290—3 10 mOsm/l). In some experiments, Ba2+
concentration was lowered to 2 mM and Cd2+ was raised to 20 M or 40 uM. All
recordings took place within 4—5 h after tissue dissection and were performed at a bath
temperature of 27°C to promote slice viability.

Recording sites were first identified at low power (10x) relative to the translucent
band corresponding to the substantia gelatinosa (Rexed’s lamina II), visible under trans-
illumination (Figure A- 1A in Appendix). Individual neurons were then targeted for patch-
clamping using a 40x water immersion objective and infrared differential contrast optics
(Figure A-lB in Appendix). Recordings were initiated in voltage-clamp mode after
nulling pipette offset and capacitance transients. After establishing whole-cell recording
configuration, cell capacitance (12—53 pF) and series resistance (15—62 M0) were
compensated. Signals were amplified (0—5 kHz bandwidth) by an Axopatch 1D amplifier

(MDS Analytical Technologies, Toronto, Canada) and saved to hard drive with a

24

 

TI

1)

CK

CUI

mo

P01

UNI
dun
by;
”fin:
P051
Chan

mem

Digidata 1320A data acquisition system running pCLAMP software (MDS Analytical
Technologies). Membrane current exceeding -0.05 nA at a holding potential of —60 mV
was indicative of an unstable recording or poor cell condition, and these recordings were
excluded from analysis. For analyses, voltage-activated currents were low-pass filtered at
500 Hz (Gaussian). P/N leak subtraction was used to correct for passive membrane
current. Recordings targeted neurons spanning a range of Sizes visible in slices. Whole-
cell capacitance was used as an indicator of cell size and read directly from the
potentiometer after nulling capacity transients. In most cases, currents are expressed as
current density (pA/pF) by dividing by cell capacitance.

In some experiments, membrane potential recordings were made in current-clamp
mode to classify neuronal intrinsic properties. Neurons were stimulated to produce action
potentials by applying depolarizing current pulses (3 8) through the recording pipette for
analyses of spike-frequency adaptation patterns. Neuronal input resistance (Rin) was
measured by recording passive membrane responses to pulses Of hyperpolarizing current.
Responses of neurons to time-dependent membrane depolarizations were determined
using ramp-hold waveforms (4.5 s) that produced linear changes in membrane potential
during the ramp phase. The sensitivity to rate of membrane depolarization was estimated
by presenting a series of waveforms having increasing ramp slopes and determining the
minimum rate of membrane potential change that activated action potentials.
Postinhibitory rebound excitation, a property associated with low-threshold T-type
channels, was examined by applying hyperpolarizing current steps (500 ms) at the resting

membrane potential, sometimes followed by a sub-threshold depolarizing current pulse.

25

 

Dru

dill)

tea;

cor.
ml)?
cor

Th.

Drugs and chemicals
TTX was purchased from MP Biomedicals (Irvine, CA). Mibefradil (Ro 40-5967,
dihydrochloride hydrate) was purchased from Sigma-Aldrich (St. Louis, MO). All other

reagents were purchased from Sigma-Aldrich or Mallinckrodt Baker (Phillipsburg, NJ).

Statistical analyses

Numerical data are presented as mean i SEM, unless noted otherwise. Statistical analyses
were performed using InStat 3 (Ver 3.06, GraphPad Software). Statistical significance of
difference between two groups was assessed using the unpaired t test with Welch
correction or Mann-Whitney test as appropriate. Comparison of means among three or
more groups was conducted using one-way ANOVA with Tukey-Kramer multiple
comparisons or Kruskal-Wallis one-way AN OVA with Dunn’s multiple comparisons.

The criterion for statistical significance was a value of P < 0.05.

26

 

Wide

bUI c

Results

Whole-cell recordings were obtained from a total of 97 dorsal horn neurons in spinal
Slices prepared from 66 hamsters. Neurons had resting membrane potentials more
negative than —50 mV under current-clamp conditions. Membrane potential was held at
—60 mV during voltage-clamp recordings, unless otherwise noted for specific

experiments.

T -type calcium currents in hamster dorsal horn neurons

We first re-examined T-type Ca2+ current (IT) in dorsal horn neurons to enable direct
correlations with spike discharge patterns. Voltage-clamp recordings were performed in
the presence of media containing TEA, TTX and Ba2+ under ionic conditions appropriate
for isolating Ca2+ currents. In the absence of Cd2+, depolarizing the membrane potential
from a hOlding potential of —90 mV to —30 mV activated an inward current with two
components, an initial rapidly decaying phase followed by a late sustained phase (Figure
2-1A). Addition of 10 uM Cd2+ reduced the amplitude of the initial phase and almost
completely abolished the late sustained current within 3 min of application. Increasing
the Cd2+ concentration to 20 uM had no additional effect on the current (Figure 2-1),
suggesting that there was little or no contribution from high-voltage-activated Ca2+
channels using the lower Cd2+ concentration. Therefore, subsequent experiments were
performed in the presence of 10 uM Cd2+. Under these conditions, we found that IT was
widely expressed by neurons recorded within Rexed’s laminae (L) I-IV (98%, n = 41/42),

but current density varied considerably from cell to cell [6 :t 4 pA/pF (SD)]. IT was not

27

 

Fig

em
5mg
Cd~
PCS"
(73C
Ellie:
mt‘a
{Al m

Auk

mil.

-30 mV

-3oo-- . . ..
o. ..O .. .d’°°°.

_200 . Peak

0

O . .
0° mwomoo o“ I .v‘
-1 00 ‘ o O

0
Residual

 

Current amplitude (pA)

 

 

0 2 4 6 810121416
Time elapsed (min)

 

Figure 2-1. Isolation of low-voltage-activated Ca2+ currents

A: inward currents recorded from a representative dorsal horn neuron were activated
every 20 s by a ZOO-ms voltage step at ~30 mV from a holding potential of -90 mV for
500 ins (uppermost). Aa+: each of the superimposed traces (black, before application of
Cd ; gray, 10 11M Cd and 20 uM Cd ) was averaged from four consecutive current
resppnses (marked with crosses in B). Ab: inward current that was sensitive to 10 1.1M

Cd was Obtained by subtraction. Ac: inward current that underwent additional blockade
after increasing Cd from 10 11M to 20 11M. Dotted lines indicate the baseline for current
measurements. B: the amplitude of peak and residual current was plotted against the time
elapsed. Cursor positions fqr measuring peak (a) and residual (O) current are shown in
Aa. 10 uM and 20 uM Cd were sequentially added into the external solution as
indicated by shading.

28

 

arrival
to -30
mV he
holding

.cmbr
plotted
(1 lm:
for halt

“ETC SI

different for neurons in the superficial (LI/II) and deep (LIII/IV) dorsal horn laminae (6 :t
1 pA/pF vs. 7 :1: 1 pA/pF; P = 0.46; Table A-3 in Appendix).

We next examined the voltage-dependence of steady-state inactivation and
activation of IT. Inactivation kinetics were analyzed by stepping the membrane potential
to —30 mV every 20 s in 10 mV increments from holding potentials of —120 mV to —40
mV held for 500 ms. An example is Shown in Figure 2-2A. IT was largely inactivated at
holding potentials positive to —60 mV and this inactivatiOn was removed as the
membrane potential approached —-100 mV. Averaged normalized currents (l/ Imax) were
plotted as a function of holding potential (n = 4) and fitted with the Boltzmann equation
(1/1max = 1/{1 + exp [(V— V1/2) / k]}) having a slope factor (k) of 12 mV and voltage
for half-maximal current (V1/2) of —74 mV (Figure 2-2C). Activation properties of IT
were studied by delivering 200-ms depolarizing voltage pulses from a holding potential
of —90 mV to —10 mV in 10 mV increments. As shown in Figure 2-23, IT appeared in the
low-voltage range with an activation threshold near —60 mV and reached a maximum at a
membrane potential near —20 mV. Values for I/ [max were plotted against the membrane
potential to generate an activation curve yielding k = 8 mV and V1 /2 = —47 mV when
fitted with the Boltzmann equation (Figure 2-2C). Overall, activation and inactivation
parameters were consistent with 1T reported previously under similar conditions
(Huguenard and Prince, 1992; Huang, 1989; Ryu and Randic, 1990).

Having established that IT in hamster dorsal horn neurons resembled T-type Ca2+
currents described by previous studies, we then determined the sensitivity of the current
to IT antagonists. As illustrated in Figure 2-3A, 200 uM Ni2+ added to the bathing

solution depressed IT by 52% i 6% (n = 6) after 4 to 9 min of perfusion. The effect was

29

 

 

 

 

:20 1-01 '1-0
0.8- -O.8
X X
“I a
_E 0.6- ~O.6 ~E
D \ \
"" -
B vtest': -10 mV 0.4- »O.4
'90 -90 02< _o.2
I
0 - . . ' . . .fi.o
. 100 pAI 420400-80 -60 -40 -20 0
25 ms Voltage (mV)

Figure 2-2. Voltage-dependent activation and inactivation of T-type Ca2+ currents
in spinal dorsal horn neurons

A: IT was activated in a representative dorsal horn neuron by a 200-ms voltage step to
-30 mV following a series of 500-ms holding potentials (Vh) ranging from -120 mV to
—40 mV at increments of 10 mV. B: the membrane potential of the same dorsal horn
neuron was held at -90 mV for 500 ms before 11- was activated by a series of ZOO-ms
testing potentials (VTest) ranging from -90 mV to -10 mV at 10-mV increments. C: IT
from four dorsal horn neurons was examined using the voltage commands described in A
and B. The current amplitude was normalized to the maximum level (I / Imax), averaged,
and plotted with V}, for the inactivation curve (:1) or VTest for the activation curve (a).
The curves were fitted with the Boltzmann equation. Errors bars represent SD. (n = 4).

30

IT amplitude (pA)

 

0246810121416

Time elapsed (min)

 

-200
2 If."
3 -150
O
'u
,3 -100
Q o
60 e

E -50 ° '.'~6

0

0 5 10 15 20

 

Time elapsed (min)

F igurc 2-3. Effect of T-type Ca2+ channel blockers

Effect of T-type Ca2+ channel blockers Ni2+ (A) and mibefradil (B) on IT in spinal dorsal
horn neurons. IT was activated every 20 s by stepping the membrane potential to ~30 mV
for 200 ms following a 500-ms holding potential of -90 mV. The peak current amplitude
is plotted on the right as a function of time elapsed (filled circle symbols in Aa and Ba
indicate cursor position for current measurements). Shading indicates when drug was
present in the external solution. Aa, Ba: the black trace was averaged from three +
consecutive measurements marked with a white cross before the application of Ni or
mibefradil (MIB), and the gray trace was averaged from three white-cross-marked points
near the end of the drug presentation. The inward currents that were sensitive to either of
the blockers, obtained by subtraction, are shown in Ab and Bb.

31

 

16'
11 a:
of 5.
perl

initi

Rate
Afte
\I‘an‘
depc

(Cm

mm

mm
pulse
incre;
al‘plic
3cm;
00m ]
Both 1

VOIIaQ

reversible with the current recovering to 78% :t 10% of control after 4 to 16 min of
washout. The IT antagonist nribefradil (MIB) was tested on three cells at a concentration
of 5 11M and resulted in a 47% i 4% reduction in IT amplitude afier 6 to 10 min of
perfusion (Figure 2-3B). The effect of MIB on IT in spinal slice neurons persisted after

initiating drug washout.

Rate-dependent activation of T -type calcium currents in spinal dorsal horn neurons
After determining steady state voltage-dependence of IT in dorsal horn neurons, we
wanted to know if the magnitude of IT was also related to the slope of membrane
depolarization (d V/dt), a characteristic that has been described for IT in thalamic neurons
(Crunelli et al., 1989; Gutierrez et al., 2001 ). This is important because of our view that
IT may contribute to the differential responses of dorsal horn neurons to rate of
membrane depolarization and time-varying sensory stimuli (Schneider, 2003, 2005).

IT activation dynamics were examined by using voltage ramps to depolarize
membrane potential at different rates to -30 mV following a 500-ms hyperpolarizing pre-
pulse to -90 mV. As can be seen by the example shown in Figure 2-4A, IT amplitude
increased along with dV/dt between ramp slopes of 400 mV/s and 2400 mV/s. Bath
application of Ni2+ depressed ramp-activated currents in a manner similar to those
activated by step depolarizations (Figure 2-4A, gray). Best-fit regression lines calculated
from plots of 1T amplitude versus ramp slope for seven cells are shown in Figure 2-4B.
Both the x-intercept and slope varied widely within our sample of cells recorded under

voltage clamp. Threshold for IT activation, as estimated from the x-intercept values,

32

 

tn (ms)

ll £3

2+

2400 _ Ni
mV/s V 100 pA L— 400— 250
Control 50 ms 200.

1500 < 150.
“100-

50-
200— 0

 

300—
1091

W
W
857*n‘tyfinwnzahwwv
“"§upfluufl“"".
W

 

 

   
  

 

10 100 1000

mV/s
1 00- /

' rY' I I Y r

400 1T ””"1'0 . 100 ”1000'
-90 va Ramp slope (mVls)

600

IT amplitude (pA)

 

 

 

20« §__

 

 

fil

200 600 1000 1400 1800 2200 2600
Ramp slope (mVls)

Figure 2-4. Activation of IT in dorsal horn neurons with ramp depolarizations

A: IT in a representative dorsal horn neuron was activated by using voltage ramps t0
depolarize the membrane potential at different rates (indicated to the left of each trace)
from a 500-ms holding potential of —90 mV to —30 mV. 1 +before (control, black traces)
and during (Ni , gray traces) presentation of 200 uM Ni is shown. B: linear regression
analysis between IT amplitude and ramp 510 e is Shown for seven dorsal horn neurons
recorded in media containing 2 or 3 mM Ba +. Inset shows representative linear fit to
data points for b. The coefficients of determination (r ) from regression analyses (0.72 to
0.99) indicate that IT amplitude and ramp slope are highly correlated. Individual
responses to ramp depolarizations for regression line a are shown in A. C: the averaged IT
time-to-peak (tp) from the seven cells in B plotted against the ramp slope at which IT was
activated. Error bars represent SEMM. (n = 7).

33

 

ail

(dc

slit
99C

“11

inv

that
Odh
fiur
COnl

ant

ffl'

>4

(IE-4'
."1

dfic

CUrr,
IESp.

lSch

averaged 46 i 21 mV/s (range 2 to 138 mV/s). The sensitivity of IT to ramp dV/dt
(defined as slope of the regression lines, pA-s/mV) varied almost five-fold, from 51 to
242 pA-s/mV, for the cells studied. Despite the cell-to-cell variability, regression analyses
showed that 87 :t 4% of the current amplitude was determined by dV/dt (range 72% to
99%). Taken together, the data suggest that dorsal horn neurons are quite heterogeneous
with respect to activation by voltage ramps.

Since IT activation appeared to be dependent on the slope of the voltage ramp, we
investigated the rate-dependence of IT kinetics. As shown in Figure 2-4C, IT time-to-
peak (tp) was dependent on dV/dt over the range of 240 mV/s to 2400 mV/s, suggesting
that rate of activation increased with increasing speed of membrane depolarization. Four
other neurons were examined using a similar protocol ramping the membrane voltage
from a holding potential of—100 mV ~ —90 mV to —30 mV ~ —20 mV, but with the Ba2+
concentration lowered to 2 mM Ba2+ and Cd2+ increased to 40 M. The maximum
amplitude, threshold and rate sensitivity of IT under these circumstances were not

significantly different from data obtained using the standard conditions described above.

T -type calcium currents in spinal dorsal horn neurons with identified membrane and
discharge properties

Although IT appeared to be a ubiquitous property of dorsal horn neurons, the
current density varied considerably from cell-to-cell under similar experimental
conditions. This variability might be expected if IT contributes to the differential
responses of dorsal horn neurons to static and time-varying membrane depolarizations

(Schneider, 2003, 2005). We therefore examined IT in a population of dorsal born

34

 

in fir
PIPCI
sumn

contil

neurons following initial characterization of intrinsic firing properties under current-
CIMp in standard ACSF solution (see METHODS). Measurements of IT were made by
depolarizing the membrane potential to —30 mV from a potential of -90 mV held for 500
ms and subtracting residual current activated from a holding potential of —40 mV to
minimize contribution from any unblocked high-voltage-activated Ca2+ current.

Dorsal horn neurons recorded under current clamp exhibited distinct differences
in firing behavior when activated by depolarizing current applied.through the recording—
pipette, as described previously (Schneider, 2003). The results from the present study are
summarized in Tabel 2-1 and Figure 2-5. Tonic cells (42%, n = 16) fired action potentials
continuously throughout the period of depolarization (Figure 2-5Ab). A second, much
smaller group of neurons (8%, n = 3) resembled tonic cells but responded to depolarizing
current following a marked delay to the first action potential (Figure 2-5Aa, open
arrowheads). A third group, phasic cells, generated a rapidly adapting discharge when
activated by a constant level of depolarizing current and amounted to half of our sample
(50%, n = 19). Since phasic cells as a group displayed a fairly wide range of spike
frequency adaptation (Schneider, 2003), we further divided them into two subgroups
(rightmost columns in Tabel 2-1, Figure 2-5Ac and d). Short burst phasic cells (P33),
responded to depolarizing current injection with only one or two action potentials
superimposed on a transient membrane depolarization (Figure 2-5Ac, filled arrowheads).
The remaining phasic cells (non-P33) typically responded with 3 or more action
potentials at rapidly diminishing frequency to depolarizing current, and the number of
spikes increased with stimulus amplitude (Figure 2-5Ad). The transient depolarization

activated in non-P53 cells was smaller than in PSB cells (compare upper trace in Figure

35

 

«i=—

 

 

 

Table 2.1. |
horn neuri

Firing p111

 

Number of

l] IPA ’

Resting
lm\

Rinl‘

Ramp 1h
1m

Postinhi

rebor

\

All \‘aerS
1’8er 15 gl

SlgnjfiCi
+ Signlllc;
I Signlllca

Table 2-1. Comparison of IT and intrinsic membrane properties of spinal dorsal
horn neurons in this study '

 

 

Non-short
. , Short burst .
Firing pattern Delay (D) Tonic (T) Phasrc (P) hasic (P ) burst phasrc
p SB (non-P53)
Number of cells 3 16 19 7 12
11-(pA/pF) 2e1 5:1 9e; 14e4*’l’I 6il
' V t ,
Remng m -58i4 -56d:2(14) -65i 1* -68:t 1* H -63:1: 1*
(mV)
9 9 9T
Rin (MO) 651i77 654i 34 319:1: 311*,r 261 ”6*7 352e46*
*9T *9T
Ramp threshold 45 i 5 *,'l' 46 d: 7
7 d: I 9 :1: 3 12 43 :l: 3
(mV/s) ( ) (I8) (I l)
P ti ' '
OS nhrbrtory 0% (0/2) 64% (9/14) 94% (16/17) 100% (7/7) 90% (9/10)
rebound _

 

All values are listed as mean :1: SEM. The number of measurements used to calculate each

value is given in parentheses if different from the number of cells for the group.
* Significantly different from T cells.

T Significantly different from D cells.
I Significantly different from non-P33 cells.

36

 

Aa

Ba

Aa b c d
D cell T cell P33 cell Non-P33 cell

k illi-

     
   
  

1111:1111111

 

 

 

 

   

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

‘1.
50 mV|__
-61W58 6% 132 AL-BG 210m
52 A 909A - L.
r p L. »_1 1 T l_ 1 L
Ba b c d
I_ III I L. I L.
AL #1.. I...
40mV[__1s
1— _/ l_ _/L L_
.../ L. _/ L
36 pA 60 pA 120 pA 150 BA
._..—-—--*""" ..._._—-—---—"'r _./“—‘—‘—'L_ M

 

 

 

 

 

Figure 2-5. Firing patterns of hamster dorsal horn neurons

A: responses of a representative cell from each category (top two traces) to depolarizing
current pulses applied through the recording pipette (lowermost trace). (Categories: D,
delay; T, tonic; PSB» short burst phasic; Non-P33, non-short burst phasic; see text for
details.) Resting membrane potential (Vm) for each neuron is indicated to the left of the
first response. Delay between pulse onset and first action potential for the D cell is
indicated by the open arrowhead. Filled arrowheads in c indicate transient membrane
depolarizations characteristic of P33 neuron activation. B: responses of the same cells to
ramp-hold current commands that depolarized Vm at different rates (waveforms are
superimposed on individual traces). Slope of membrane depolarization (d V/dt) for lower
and middle traces: 6 mV/s and 9 mV/s for D; 5 mV/s and 10 mV/s for T; 18 mV/s and 37
mV/s for P88 25 mV/s and 55 mV/s for non-P33. Upper trace: rectangular current step to

the same level of depolarization (upper trace). Action potential responses in c and d are
indicated by arrows.

37

 

2-5.1t' 11111
wmmmu
potential 1
consistent
current cc
discharge
neurons 11
ramp-hold
b) the mo:
needed to ;
activation .
aclitalion 1
the rtSpong
7..

I‘M N]w ( y
Iou-thresht
T113
rebouhd me.
Currcm l PI g1
cells than 101
filing Pattern

deplllailllnp

F

C

I'Flgllle 284)

2-5Ac with middle trace in Figure 2-5Ad) or absent altogether (Figure 2-6). As
summarized in Tabel 2-1, phasic cells had a more hyperpolarized resting membrane
potential (——65 mV) and lower Rin (319 MO) than tonic cells (—56 mV; 654 MO),
consistent with previous Observations (Schneider, 2003). When depolarized by ramp-hold
current commands, tonic cells discharged primarily during the steady-state depolarization;
discharge frequency was unrelated to the ramp slope (Figure 2-SBb). Delay cells, like
neurons with tonic firing patterns, fired action potentials during the hold phase of the
ramp-hold current command (Figure 2-SBa). Typically, phasic cells were activated only
by the most rapid membrane depolarizations (Figure 2-SBc and d). Depolarization rates
needed to activate phasic cells were five- to Six-times higher than those required for
activation of tonic- and delayed-firing neurons, and were close to the threshold for
activation of IT studied in our voltage clamp experiments. As can be seen in Figure 2-7,
the responses of phasic cells to step or ramp-hold depolarizations were depressed by 200
11M Ni2+ (n = 4) and 5 uM MIB (n = 3), consistent with underlying participation from a
low-tlrreshold T-type Ca2+ current.

Twenty-five of 33 cells tested (76%) exhibited postinhibitory rebound (PIR), a
rebound membrane depolarization and discharge following injection of hyperpolarizing
current (Figure 2-8A). Although PIR appeared to be Observed more frequently in phasic
cells than tonic cells (Tabel 2-1), there was no statistically significant association with
firing pattern (P = 0.067). PIR in hamster dorsal horn neurons was typically activated by
depolarizing the membrane potential to —45 to —60 mV from around —80 mV, and
increased in amplitude when activated from more hyperpolarized membrane potentials

(Figure 2-8A). A time-dependent rectification (voltage “sag” in Figure 2-83) was evident

38

 

Figure 2-
Transient
threshold
typicall} 2
non-P53 c

 

1“

/K r“ A “
eta—f -68 175 pA
__IL

100 pA .
__J

 

 

 

 

 

Figure 2-6. Transient depolarization in phasic cells

Transient depolarizing potentials (arrows, upper traces) were observed in supra- and sub-
threshold responses of PSB cells to depolarizing current pulses (lower traces) but were
typically absent from non-P33 cells at the same Vm (dotted lines). Rin: PSB cell, 344 MO;
non-P33 cell, 194 MO.

39

 

 

Figllre 2.7_ ‘
.4: effect of“)
30011.11 Nr

aflll‘ated lrar
113C682 CUITCr
mVe‘sto 48 In
my": 31 inhi
01 Cllrmm ran
ramPing l 'm 2
Were S”Pfi‘rim
to the left 0 f1.

.Aa
ACSF Ndl

 

 

 

 

 

 

 

500ms
-71 BOpA—L-H L
___l I._ _J L_
.Ab
20va_
/ / 1s
_71 80 A 41%
W mm
Ba
ACSF MIB
20mVL
53 53“““ 5009‘?
- 20pA - J m V I'-
.I L. J L
.Bb
20mm
18

mm

Figure 2-7. Effect of N i2+ and mibefradil on phasic firing

A: effect of Ni + on ramp- and step-activated responses of a representative phasic cell.
200 11M Ni blocked the spike discharge generated by a current step (a) and a ramp-
activated transient depolarization (b, arrows). Upper traces: voltage responses. Lower
traces: current commands. Current commands used in Ab depolarized Vm at rates of 6
mV/s to 48 mV/s and the transient depolarization was activated by a voltage ramp of 48
mV/s. B: inhibitory effect of 5 11M mibefradil on phasic firing evoked by a current step (a)
or current ramps (b) in two other dorsal horn neurons. The cell in Bb was activated by
ramping Vm at 12 mV/s or greater (range 4 mV/s to 31 mV/s). Five consecutive traces
were superimposed and shown in Aa and Ba. Resting Vm for each condition is indicated

to the left of the traces.

40

 

 

Figure 2-
.4: a dors;
lupner Ira
CEII exhil‘.
92 m\' (d
discharge
action p01
hiperpola
dependent
neuron W21
f7mvm
0‘5 101 T1
traces in .4

A B

 

 

 

-40 mV
-57 )— ‘76 -54 mV
'78-‘J ‘ '70
2+
Ni
:> annjBAT—
N”- .. ' _J 25 mV
-92 _l 25 mv \ 100 ms
1ootns -9Z
20 pA .\
50 pA
-50 E 0 E170
C .2+
ACSF TTX TTX + NI

_J 25 mV
-57 mV 2 E9 200 ms
0 pA

MW— WV—

Figure 2-8. Postinhibitory rebound in hamster dorsal horn neurons

A: a dorsal horn neuron was given a depolarizing current pulse from the resting Vm
(upper trace) or following hyperpolarizing current steps (middle and lower traces). The
cell exhibited rebound depolarizations following prior hyperpolarization to —78 mV and -
92 mV (dotted lines indicate resting V +at —57 mV). Both rebound depolarization and
discharge were blocked by 200 uM Ni applied in ACSF (gray traces). B: a rebound
action potential was generated in a dorsal horn neuron depolarized to —54 mV from
hyperpolarized Vm (arrowheads; resting Vm, —70 mV). PIR was independent of a time-
dependent rectification (voltage “sag”, indicated by the arrows). C: another dorsal horn
neuron was hyperpolarized to -83 mV for 500 ms and then returned to the resting level of
-57 mV at rates ranging from 28 mV/s tg+131 mV/s. The PIR responses were resistant to
0.5 uM TTX but blocked by 200 111M Ni .Current commands are shown as lowermost
traces in A—C. Firing pattern type: T cells for A and B; non-P33 cell for C.

41

 

 

in steps I
suggesur
tesedlll
rise of m
8f).sinn
740.1eh,
alnel
:\
character
0=6pA
represent;
dlceHsin
Signlllt‘an]
Wmhc
[hminjoni
in APl‘endi

0.0003 l 1th

in steps to membrane potential below —70 mV in 60% of neurons exhibiting PIR (15/25),
suggesting the presence of a hyperpolarization-activated cation current (1h). For all cells
tested (100%, n = 23), the rebound depolarization increased in amplitude as the rate of
rise of membrane voltage during the return to resting potential became steeper (Figure 2-
8C), similar to the responses of phasic cells to ramp-hold depolarizing currents (Figure 2-
7Ab, left). The rebound depolarization was resistant to TTX but blocked by Ni2+ (n = 2)
(Figure 2-8C) and MIB (n = 2; not shown).

As we expected, IT was observed in the majority of dorsal horn neurons
characterized in current-clamp (95%, n = 36/38) with considerable variation in amplitude
[7 d: 6 pA/pF (SD); range 1 pA/pF to 36 pA/pF]. Examples of IT recorded from
representative cells of each firing pattern class are shown in Figure 2-9, and the data from
all cells in the sample is summarized in Tabel 2- 1. Average IT density in phasic cells was
significantly greater than in delay cells, but comparable to IT in tonic neurons (Tabel 2-1).
Mean IT expressed in PSB cells was 2.3-fold greater than in non-P53 cells and exceeded
that in tonic and delay cells by comparable amounts (Tabel 2-1; also refer to Figure A-2
in Appendix). IT was significantly larger in cells exhibiting PIR (9 i 1 pA/pF, n = 25; P =

0.0002) than in those neurons without PIR (2 d: 1 pA/pF, n = 8).

42

 

Figure
Dorsal I
depolnr:
and 10 1
in norm
median
P SB. Sill
ID): 13:
PA pF (
baseline

 

Figure 2-9. IT in neurons with identified firing patterns

Dorsal horn neurons were voltage clamped at a V}, of —90 mV and IT was evoked by +
depolarizing voltage steps to -30 mV. External bathing solution contained 3 mM Ba
and 10 uM Cd and was added after firing pattern characterization under current clamp
in normal ACSF (see METHODS). Five traces were superimposed and illustrate the
median IT amplitude for each firing pattern classification (categories: D, delay; T, tonic;
PSB, short burst phasic; Non-P33, non-short burst phasic). Median IT amplitudes: 31 pA
(D); 134 pA (T); 244 pA (P33); 116 pA (non-P313). IT density in examples shown: 1
pA/pF (D); 6 pA/pF (T); 8 pA/pF (P33); 4 prF (non-P33). Dotted line indicates the
baseline for inward current measurements.

43

 

Discussior
Our result:
earlier stm
Here. we r
10 depolnri
potentials.
demonstra‘
dependent
l C runelli t’.
phasic-lirir
mOI'ements
and respons

2 .

0P6 Ca” (3
an IlmI-‘Oflan
llIne.(~ar}.inE
We 1
“0101115th
Randic (1991
may reflect £1
IOIIIC median
consistent Wit

00m (Talley 6

suggests lhat c

Discussion

Our results show that dorsal horn neurons possess T-type Ca2+ currents, consistent with
earlier studies (Ryu and Randic, 1990; Huang, 1989; Russo and Hounsgaard, 1996a).
Here, we presented new findings suggesting IT expression is highest in cells that respond
to depolarizing current injection with a rapidly adapting discharge of one or two action
potentials, and lowest in neurons with tonic or delayed-firing properties. Furthermore, we
demonstrated for the first time that activation of IT in dorsal horn neurons is strongly
dependent on the rate of membrane depolarization, similar to IT reported in thalarrric cells
(Crunelli et al., 1989). We believe this property contributes to selective excitation of
phasic-firing dorsal horn neurons by rapid membrane depolarizations and stimulus
movements (Schneider, 2003, 2005). This conclusion is based on our observation that IT
and responses of phasic cells to ramp membrane depolarizations are blocked by the T-
type Ca2+ channel antagonists Ni2+ and mibefradil. Taken together, our results support
an important role for IT in helping dorsal horn neurons differentiate between static and
time-varying afferent input (Schneider, 2005).

We found that nearly all neurons in the hamster dorsal horn (98%) exhibit IT,
which is higher than the proportion of rat LII—V neurons (72%) reported by Ryu and
Randic (1990) to have low-voltage-activated Ca2+ currents under similar conditions and
may reflect a species-dependent difference. Both studies suggest that IT is a ubiquitous
ionic mechanism regulating neuronal excitability in the spinal dorsal horn and are
consistent with widespread expression of T-type Ca2+ channel transcripts in the dorsal
horn (Talley et al., 1999). Our data showing wide variance in IT current density strongly

suggests that expression of IT by dorsal horn neurons is non-uniform. Modeling studies

44

 

have in
influen
ditt‘ercr
electric
measur
dendrit
section

slice sr

pI010C(
arrival
The IT
ranges
(XieSp
(0101115
Was Sh
rat dOr
V‘ollag.
Went;
II 990 )
CUrren.
Studie8

SHIN”)

have indicated that subtle changes in T-type Ca2+ channel density can have a potent
influence on neuronal electrical activity (Chorev et al., 2006). Thus, we might expect that
differences in T-channel density would also have substantial effects on neuronal
electrical behavior and integrative properties of dorsal horn networks. However, our
measurements may not take into account the contribution of T-type Ca2+ channels with a
dendritic distribution (Destexhe et al., 1998) that may have been removed during the
sectioning process, as is typically the case for recordings made from neurons near the
slice surface.

Despite differences in divalent cation concentrations and voltage clamping
protocols, the IT expressed in hamster dorsal horn neurons resembles low-voltage-
activated, transient Ca2+ currents reported in other neurons recorded from brain slices.
The IT in this study has broadly similar steady-state activation and inactivation voltage
ranges as T-type Ca2+ currents found in thalamus (Crunelli et al., 1989), hypothalamus
(Niespodziany et al., 1999), suprachiasmatic nucleus (Kim et al., 2005) and cerebellum
(Mouginot et al., 1997). Steady-state inactivation (V1/2 = —74 mV) in the present study
was shifted by about 12 mV more positive in comparison with that reported previously in
rat dorsal horn neurons (Ryu and Randic, 1990). This shift may reflect the stronger
voltage (-30 mV versus -45 mV) used to activate IT; a relatively more depolarizing
potential for inactivating half of the current is therefore to be expected. Ryu and Randic
(1990) also reported a more potent Ni2+ blockade (91%) of the low-threshold Ca2+
current (versus 52% in the present study). The Ni2+ concentration (200 uM) used in both
studies was sufficient to produce half-blockade of the three known T-type channel

.2
subtypes (Lee et al., 1999b). However, N1 + blockade is voltage-dependent, with less

45

 

block at 11
found that
reported f
2002; Kin
half ( 47° 0
One is lll‘d
Callas tht
cells recur

IICUI’OIlS.

T-(l‘pe cal
Involveme-
lllleclion c
20113; Sun
dorsal hor
hEimsler d.
T‘ll'Pe Ca
Potentials
ofPIR Ill-7
which arm
beevokcd
lhar requlrt

that bIOCk .

block at the -—30 mV test potential used in the present studies (Lee eta1., 1999b). We also
found that potency of the T-channel antagonist mibefradil was lower than what has been
reported for other neurons (Viana et al., 1997; McDonough and Bean, 1998; Lee et al.,
2002; Kim et al., 2005). In the present study, mibefradil (5 uM) decreased IT by about
half (47%). There are two explanations for the lower mibefradil potency in our studies.
One is that mibefradil may be less efficacious as a T-channel blocker when Ba2+ replaces
Ca2+as the charge carrier (Martin et al., 2000). Another possibility is that drug access to
cells recorded in slice preparations may be lower than in preparations using isolated

neurons.

T -type calcium currents and postinhibiton rebound (PIR)

Involvement of low-voltage activated Ca2+ channels in rebound firing following an
injection of hyperpolarizing current has been reported for many neurons (Israel et al.,
2008; Sun and Wu, 2008; Alvifra et al., 2009) and has also been described for turtle
dorsal horn cells (Russo and Hounsgaard, 1996a). We found PIR in the majority (76%) of
hamster dorsal horn neurons we recorded and associated this property with activation of a
T-type Ca2+ current for the following reasons. First, PIR was activated at membrane
potentials of —45 mV to —60 mV, falling in the range for IT activation. Second, generation
of PIR typically required prior membrane hyperpolarizations near —80 mV, a voltage at
which around 60% of IT is available for activation (Figure 2-2C). Third, PIR could only
be evoked by fast membrane depolarizations (Figure 2-7B) with dV/dt consistent with
that required to activate IT. Finally, PIR was blocked by Ni2+ and MIB at concentrations

that block T-type channel activation. It Should be noted that hyperpolarization-activated

46

 

catior
lh-lik
2002:
a mi
perfo:

that I]

Ionic!
[T has

cation currents (1h) may also contribute to rebound firing (van Welie et al., 2006) and an
Ih-like current has been reported in rat superficial dorsal horn neurons (Grudt and Perl,
2002). We also observed an Ih-like sag in voltage responses to hyperpolarizing current in
a majority cells with PIR (Figure 2-8B). Since our current clamp recordings were
performed without blocking Na+, K+ or Ca2+ currents, we cannot rule out the possibility
that 1h also contributes to rebound firing documented in our experiments. However, 1h
seems unlikely to be the only ionic mechanism of PIR since rebound firing could be
present in the absence of voltage sag.

The functional significance of PIR in spinal dorsal horn neurons is uncertain.
Low—threshold Ca2+ channel-mediated PIR has been suggested to influence rhythmic
firing behavior of neurons in the thalamus and other brain regions (Huguenard, 1996).
Rhythmic firing can be recorded from dorsal horn neurons in vivo (Sandkiihler and
Eeblen-Zajjur, 1994) but is not normally characteristic of hamster dorsal horn neurons
under standard in vitro conditions (Schneider, unpublished observations). From the
perspective of encoding afferent information related to stimulus velocity, it is possible
that activation of local inhibitory pathways (Lu and Per], 2003; Yasaka et al., 2007;
Schneider, 2008) could amplify the sensitivity of dorsal horn neurons to rapid membrane
depolarizations by hyperpolarizing the membrane potential and removing inactivation of

the T-current.

Ionic basis of phasic dorsal horn neuron responses to rapid membrane depolarizations
IT has been implicated for a long time in the production of oscillatory behavior of CNS

neurons (see Huguenard, 1996). Our present findings strongly suggest that IT also

47

   

to the skin
excet3ded -
Phaslc'firi
sensitiVe a
with 3530C
channel an
phasic n0”?
rampS and j
The

bear3 3 SH 01

3+

Ca curren
depolallzflll‘
olfactory m
exceeding 3‘-
(Cmnelli eh

reverse the R

 

are fewer eh

amplitude at

7
JJOI). I! shot

 

dorsal horn n‘

for thalamic t

contributes to the striking ability of phasic-firing dorsal horn neurons to respond
selectively to fast membrane depolarizations and signal rapid mechanical stimuli applied
to the skin (Schneider, 2003, 2005). First, activation of IT began only when dV/dt
exceeded ~ 46 mV/s and increased in amplitude with the speed of depolarization. Second,
phasic-firing cells expressed the highest IT density in the dorsal horn. Third, rate-
sensitive activation of IT under voltage clamp conditions and a transient depolarization
with associated action potential discharge under current clamp are blocked by the T-
channel antagonists Ni2+ and MIB. These results are in line with previous findings that
phasic neurons in Rexed’s LIII—V are selectively excited by fast depolarizing current
ramps and rapidly-moving stimuli applied to the skin (Schneider, 2003, 2005).

The activation of 1T by voltage ramps that we describe for dorsal horn neurons
bears a strong resemblance to T-type Ca2+ currents characterized in other cells. T-type
Ca2+ currents that are activated from hyperpolarized resting potentials by rapid ramp
depolarizations have also been described in glomerulosa cells (Vérnai et al., 1995) and
olfactory receptor neurons (Kawai and Miyachi, 2001). Membrane depolarizations
exceeding 30 mV/s are required for activating an IT in cat lateral geniculate neurons
(Crunelli et al., 1989). In thalamic cells, voltage ramps of lower velocity appear to
reverse the removal of T-channel inactivation by a prior hyperpolarization, so that there
are fewer channels available for activation, resulting in a diminishing of the current
amplitude at slower membrane depolarizations (Crunelli et al., 1989; Guiterrez et al.,
2001). It should be noted that cell-to-cell variability in IT activation threshold for hamster
dorsal horn neurons (46 d: 21 mV/s) is greater than that reported by Crunelli et al. (1989)

for thalamic cells (30 :t 2 mV/s). Furthermore, the sensitivity of dorsal horn neurons to

48

 

rate of me
48). These
differing i
Th

certain do:
ionic mecl
membrane
than for ac
suggesting
3'1). Satin
accounting
resembles z
1998: Mag-
reported an
activated h)
Current has
Contribmc t:
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The
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rate of membrane depolarization showed considerable variability between cells (Figure 2-
4B). These differences could reflect differential expression of T-channel subunits
differing in activation kinetics (Chemin et al., 2002; Talley et al., 1999).

The present findings support a major involvement of IT in shaping the ability of
certain dorsal horn neurons to signal dynamic membrane depolarizations, although other
ionic mechanisms may also contribute to this ability. Our results showed that the rate of
membrane depolarization for activating P813 and non-P53 cells was significantly higher
than for activating tonic and delay cells. However, IT was elevated only in PSB cells,
suggesting that another mechanism may underly rate sensitivity in non-P33 cells (Tabel
2-1). Safronov et al. (1997) described a slowly-inactivating, TTX-sensitive Na+ current
accounting for ~ 5 - 20% of total Na+ current in rat dorsal horn neurons that broadly
resembles a persistent Na+ current (INap) activated near ~ —65 mV (Parri and Crunelli,
1998; Magistretti and Alonso, 1999; Magistretti et al., 2006). Recent studies have
reported an [Nap in spinal lamina I neurons and ventral horn motoneurons that are
activated by voltage ramps (Prescott and De Koninck, 2005; Kuo et al., 2006). This
current has a slower threshold (~ 10 mV/s) for ramp activation than [T but could
contribute to neuronal excitation over the range of ramp depolarizations used in the
present study.

The sensitivity of phasic-firing dorsal horn neurons to rate of membrane
depolarization may also be influenced by activation of K+ currents. A low-threshold,
dendrotoxin-sensitive K+ conductance has been reported to underlie selective responses
of phasic-firing neurons in the ventral cochlear nucleus to rapid depolarizations

(Ferragamo and Oertel, 2002). Other voltage-gated K+ channels (M-type) that are

49

ho

SU

811

 

inhibited by the cholinergic agonist muscarine have been reportedly expressed by dorsal

horn neurons (Rivera-Arconada and Lopez-Garcia, 2005). Recent modeling studies
suggest that activation of M-type currents can increase spike-frequency adaptation and
enhance neuronal responses to fast inputs (Prescott and Sejnowski, 2008). Because of its
relatively slow kinetics, the M-current may only permit high frequency signals to drive

membrane depolarizations leading to firing.

Functional significance
Our recordings establish the ubiquity of a low-threshold, transient Ca2+ current in spinal
dorsal horn neurons. However, it is clear from the data that IT is most strongly expressed
within a subpopulation of phasic-firing cells. IT may thus be essential for the signaling of
information related to stimulus velocity within the dorsal horn intrinsic network
(Schneider, 2003, 2005). Our previous studies showed that cells in dorsal horn LIII—V
that are selectively excited by rapid membrane depolarizations are intemeurons with
short, longitudinally oriented axons and establish local circuit connections with
neighboring cells (Schneider, 2003, 2008). However, IT may be also expressed to a lesser
extent in other functional types of dorsal horn neurons, as low-threshold Ca2+ currents
have also been reported in output cells having ascending axonal projections to the
thalamus (Huang, 1989; Ikeda et al., 2003).

The molecular counterparts of IT that participate in velocity-sensitive responses of
dorsal horn neurons have not been identified. However, mRNA transcripts of the T-type
Ca2+ channel subfamily, Cav3.1, Cav3.2 and Cav3.3, have been reported in rat spinal

dorsal horn (Talley et al., 1999). Cloned Cav3.1 and Cav3.2 channels generate large,

50

rapidly inactivating currents (Chemin et al., 2002) that are well-suited to the short burst
firing exhibited by phasic dorsal horn neurons sensitive to high rates of dV/dt and thus
may contribute to the proposed velocity encoding function of these cells. Interestingly,
the Cav3.2 channel isoform has been found to be expressed by D-hair receptors, 3 class
of vertebrate cutaneous mechanoreceptor, and could underlie their high sensitivity to
moving stimuli applied to the skin (Shin et al., 2003). Our results suggest that T-type
channels may be integral components of specialized central dorsal horn circuits for
processing information related to stimulus velocity.

Finally, the present results should be taken in context with other studies that have
reported persistent cation currents in spinal neurons. Our findings do not exclude the
possibility that activation of [Nap contributes to the responses of phasic—firing dorsal horn
neurons to dynamic membrane depolarizations. However, [Nap appears to function
principally in spinal neurons by enhancing responses to sustained excitatory inputs (Kuo,
et al. , 2006). [Nap expression is greater in tonic-firing dorsal horn neurons than in phasic-
firing cells and prolongs responses to transient inputs, leading to an idea that persistent
inward currents facilitate encoding of stimulus intensity (Prescott and De Koninck, 2005).
Our results suggest that differential expression of transient and persistent inward currents
in dorsal horn cells may contribute to differential processing of stimulus intensity and
velocity information by functionally-defined spinal circuits (Schneider, 2005). These
currents could be subject to actions by neurotransmitters, thus modulating integration of
sensory information within the networks (Mittmann and Alzheimer, 1998; Gorelova and

Yang, 2000; Chemin et al., 2006).

51

CHAPTER THREE
A FAST INACTIVATING T-TYPE CALCIUM CURRENT IS

INVOLVED IN SENSING MEMBRANE DEPOLARIZATION RATE
BY SPINAL DORSAL HORN NEURONS

52

Introduction
The spinal dorsal horn, first central relay of somatic sensory system, plays an important
role in integrating diverse sensory inputs encoded by primary sensory neurons
innervating skin, muscle, joints, and internal organs (Willis & Coggeshall, 2004). The
neurons in this area exhibit various electrophysiological properties, which as well as the
intrinsic ionic conductances, determine how the sensory information is processed (e. g.
Prescott and De Koninck, 2005). It has been shown previously that expression of T-type
Ca2+ currents is associated with the selective responses of some phasic-firing dorsal horn
neurons to rapidly-changing membrane depolarization caused by direct current injection
(Schneider, 2003; Chapter 2) or fast-moving cutaneous stimuli (Schneider, 2005), and the
rate-dependent activation of this current may be the underlying mechanism (Chapter 2).
Three pore-forming (1] subunits (Cav3.1, Cav3.2, and Cav3.3) of T-type Ca2+
channels have been cloned (Cribbs et al., 1998; Perez-Reyes et al., 1998; Lee et al.,
1999a), and when expressed in HEK 293 cells, the recombinant channels generate low-
voltage-activated Ca2+ currents with distinctive kinetic properties, which may contribute
differently to neuronal excitability (Chemin et al. , 2002). Similarly, native T currents in
thalamic neurons also show distinct activation and inactivation kinetics, and different
physiological functions are proposed for these currents (Huguenard & Prince, 1992;
Zhuravleva et al. , 2001; Joksovic et al., 2005). In the spinal dorsal horn, mRNAs of all
three Cav3 subunits are detected by in situ hybridization (Talley et al., 1999). However,
whether T currents in dorsal horn neurons exhibited heterogeneous properties is not clear,

and neither were the molecular constituents of the currents in individual cells.

53

Therefore, in this study, I examined T-type Ca2+ currents in dorsal horn neurons
from a hamster spinal cord slice preparation using whole-cell patch-clamp recordings.
The currents were analyzed using an exponential stripping procedure to isolate the
constituent current components. Cells were also characterized under current-clamp mode
to compare expression of the current components and intrinsic firing properties. To
identify the Cav3 subunit that mediate each current component, single cell RT-PCR was
performed after electrophysiological recordings. The data show that a fast inactivating T-
type Ca2+ current was selectively activated by fast membrane depolarization and highly
expressed in rapidly-adapting phasic-firing dorsal horn neurons. This fast T current may
underlie the ability of these cells to detect dynamic mechanical stimulation and is

probably mediated by CaV3.1 channel subtype.

54

Methods
All protocols involving the use of animals were performed in accordance with the
National Institutes of Health Guide for the Care and Use of Laboratory Animals and were

approved by the Institutional Animal Care and Use Committee at Michigan State

University.

Slice preparation

Spinal cord slices were prepared from 9- to 14-day-old Syrian hamsters of both sexes
anesthetized with urethane (1.5 mg/ g, intraperitoneal) as described previously (Schneider,
2003). Briefly, following the removal of vertebrae and meninges from the spine, a block
of lumbosacral spinal cord was dissected and glued on the stage of a vibrating microtome
(Vibratome 3000, St. Louis, MO), and 300-um-thick transverse slices were cut slowly
(<1 mm/min) at the maximum available vibration frequency. Dissection and slicing steps
were carried out in cold (4—8°C) dissection solution containing (in mM) 179 sucrose or
glycerol, 2.5 KCl, 0.2 CaClz, 10 MgClz, 1.25 NaHzPO4, 26 NaHCO3, and 25 D-glucose
(pH 7.3 5—7.45, 290—310 mOsm/l) equilibrated with 95% 02—5% C02. The use of
glycerol to replace sucrose in dissection solution to enhance viability of neurons was
reported previously (Ye et al., 2006), and no difference in neuronal electrophysiological
properties was observed from the slices prepared using either of the solutions (Tables A-1
and A-2 in Appendix). Slices were then incubated in an oxygenated artificial
cerebrospinal fluid (ACSF) containing (in mM) 120 NaCl, 2.5 KCl, 2.5 CaClz, 1.5
MgSO4, 1.25 NaHzPO4, 26 NaHCO3, and 10 D-glucose (pH 7.35—7.45; 290—310

mOsm/l) at room temperature for at least one hour before electrophysiological recording.

55

E lectroph ysiological recording

Four to six transverse spinal cord slices were obtained from each animal, and one dorsal
horn neuron was recorded from each slice. For electrophysiological recording, slices
were transferred to a chamber (volume, 1.5 ml) mounted on a fixed-stage microscope
(Olympus BXSlWI, Central Valley, PA) and continuously perfused at 5—6 ml/min with
ACSF gassed with 95% 02-5% C02. Patch recording electrodes were fabricated from
borosilicate glass pipettes (N -51A, Drummond Scientific, Broomall, PA) and filled with
internal solution containing (in mM) 130 K-gluconate, 5 NaCl, 1 CaClz,l MgClz, 11
EGTA, 10 HEPES, 2 Mg-ATP, and 0.1 Li-GTP (pH 7.3, 280—285 mOsm/l; DC
resistances, 4—7 MQ). T-type Ca2+ currents were isolated under voltage clamp using an
external solution containing (in mM) 107 NaCl, 3 BaClz, 2 CsCl, 1.3 MgClz, 26
NaHCO3, 10 D-glucose, 20 TBA, 0.5 uM TTX, and 10 uM CdC12 (pH 7.35—7.45; 290—
310 mOsm/l) adopted from Ryu & Randic (1990). All recordings were performed at 27°C
maintained by an in-line temperature controller (Warner TC-324B, Hamden, CT) within
445 h after dissection.

Recording locations were first identified at low power (10x) by using the
substantia gelatinosa (lamina II), a thin translucent band under transillumination, as a
visual landmark (Figure A-lA in Appendix). Individual cells were then approached under
a 40x water immersion objective and infrared differential contract optics (Figure A-lB in
Appendix). All recordings were initiated in voltage-clamp mode after nulling pipette
offset and capacitance transients. After whole-cell recording configuration was
established, cell capacitance [26 :t 9 pF (SD), n = 52] and series resistance [30 x 9 M9

(SD), n = 52] were compensated. Signals were amplified (0—5 kHz bandwidth) by an

56

Ax0patch 1D amplifier (MDS Analytical Technologies, Toronto, ON, Canada) and saved
to hard drive with a Digidata 1320A data acquisition system running pCLAMP 8
software (MDS Analytical Technologies). Membrane current exceeding -0.05 nA at a
holding potential of —60 mV was usually indicative of an unstable recording
configuration or poor cell condition, and these recordings were excluded from analysis.
Prior to the isolation of T-type Ca2+ currents, the recordings were temporarily switched
to current-clamp mode to examine the intrinsic properties of dorsal horn neurons using
the procedure described previously (Chapter 2). T-type currents were isolated and
recorded after voltage-clamp recordings resumed. Passive membrane current was
corrected by the implementation of PM leak subtraction, in which the cell is applied with
a command waveform (or pulse “P”) that is scaled down by a factor of l/“N” for “N”
times. For analysis, whole-cell currents were low-pass filtered at 500 Hz (Gaussian). In
most cases, currents were expressed as current density QiA/pF) by dividing by whole-cell
capacitance, an indicator of cell size and read directly from the potentiometer after
neutralizing capacity transients.

To analyze T-type Ca2+ current components and estimate their amplitude, an
exponential stripping procedure developed by Zhuravleva and colleagues (1999) was
adopted. T-type current was activated at —30 mV from a potential of -90 mV held for
500 ms, and an averaged current trace from 3-5 trials was used for the analysis. In ten
cells, a sustained residual current resembling high-voltage-activated Ca2+ current was
recorded from a holding potential of —40 mV and therefore subtracted from the current
activated from —90 mV before the analysis. The current trace was then converted to a

semi-logarithmic plot of the current amplitude in a logarithmic scale along with the linear

57

time course. The current decay phase was fitted with linear functions, and the slope of the
linear regression was used to determine the decay time constants of individual current
components. The amplitude of a current component with slower decay was estimated by
extrapolating the linear function to the current peak, and the amplitude of a faster-
decayed current component was approximated by subtracting the slow current amplitude

from the peak T—type current, as shown in Figure 3-1.

Drugs and chemicals

TTX was purchased from MP Biomedicals (Irvine, CA). Mibefradil (Ro 40-5967,
dihydrochloride hydrate) was purchased from Sigma-Aldrich (St. Louis, MO). All other
chemicals were purchased from either Sigma-Aldrich (St. Louis, MO) or Mallinckrodt

Baker (Phillipsburg, NJ).

Single cell reverse transcription polymerase chain reaction (R T -PCR)

To identify the Cav3 subunit mediating T-type Ca2+ currents in dorsal horn neurons,
single cell RT-PCR was carried out on the cells recorded in electrophysiological
experiments. Gloves were worn throughout these experiments. Patch pipettes were
sterilized, and the internal solution was made with nuclease-free water (Fisher Scientific,
Fair Lawn, NJ). Following the completion of whole-cell patch clamp recordings, the cell
content of the neuron was aspirated into the patch pipette without loss of the seal (Figure
A-lC in Appendix). The pipette tip was broken, and the contents (5—8 ul) were expelled
into a PCR tube containing 5 ul of nuclease-free water (Fisher Scientific), 1 ul of

dithiothreitol (DTT, 0.1 M, Invitrogen, Carlsbad, CA), 1 ul of RNase inhibitor (40 U/ul,

58

-30 mV

 

 

 

 

Current amplitude (pA)

 

 

01°
0 50 100 150 200

 

Time (ms)

Figure 3-1. Analysis of T-type Ca2+ current components by using an exponential
stripping procedure

A, a T-type Ca + current (IT, lower trace) composed of a fast (1T9f) and a slow current
component (1T,s) was activated from a representative dorsal horn neuron by the voltage
command shown above the trace. A single-term exponential fitted to 1T,s decay is
superimposed on the current trace and extended to the beginning of the inward current.
The vertical dotted line indicates the time of the current peak, at which the amplitude of
IT,s is estimated by extrapolation of the fit. The amplitude of IT; is obtained by
subtracting the 1T,s amplitude from the peak amplitude of IT. B, the current trace in A was
converted to a semi-logarithmic plot, and the current decay phase was fitted with two
linear functions. The decay time constant of IT; (rf) and IT,s (rs) was derived from the
slope of the linear functions and indicated on the plot.

59

Promega, Madison, WI) and 1 ul of random hexanucleotides (0.5 ug/ul, Promega). The
tube was placed immediately in dry ice and then stored at -80°C. Reverse transcription
was carried out as previously reported (Han et al., 2005) with minor modification. The
cell content mixture was heated to 70°C for 10 min and quickly chilled on ice. After brief
centrifugation, the solution was mixed with 4 ul of 5X F irst-Strand Buffer (250 mM
Tris—HCl, 375 mM KCl, 15 mM MgC12,Invitrogen), 1 ul DTI‘ (0.1 M, Invitrogen), 1 ul
mixed deoxynucleotide triphosphates (dNTPs, 10 mM, Invitrogen), and 1 ul
SuperScriptTM II reverse transcriptase (RT, Invitrogen). Following 10-min incubation at
room temperature, the solution was warmed to 42°C for 50 min for cDNA synthesis. The
reaction was terminated by heating at 70°C for 15 min, and RNase H (2 U/ul, Invitrogen)
was added to remove RNA from the RNA-DNA hybrid. The single cell cDNA was then
stored at —20°C until used in PCR.

The presence of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) cDNA was
first examined using real-time PCR to confirm success in cell harvesting and reverse
transcription. For GAPDH-positive single cell samples, a two-step PCR protocol was
carried out to detect the cDNA of three known Cav3 subunits. The first amplification for
all Cav3 cDNAs was performed on a TGradient therrnocycler (Biometra, Goettingen,
Germany), and the product was diluted 10 times and used in the subsequent real-time
PCR using an inner forward primer specific to one of the Cav3 subunit cDNAs.

For the first-step of the Cav3 PCR, one 50-ul reaction mixture was prepared for
each single cell sample and contained 25 ul of PCR Master Mix (Promega), 5 ul of single
cell cDNA template, and the outer primers: Cav3 forward, 5’-CGCTCCAGCCGGAACA

GC-3’; Cav3 reverse, 5’-GCAATGGTGATACAGTTGAGGA-3’. The reaction went

60

through 40 cycles of 95°C for l min, 53°C for 1 min, and 72°C for 1 min. Real-time PCR
was performed with the Stratagene Mx3000P® QPCR system (Agilent Technologies,
Santa Clara, CA) using SYBR® GreenERTM qPCR SuperMix (Invitrogen). Triple 25-ul
reactions with 2 ul of cDNA or diluted PCR product template were prepared for each
cDNA examined. The amplification proceeded with the following profile: 50°C for 2 min
and 95°C for 10 min, followed by 45 cycles (50 cycles for GAPDH) of 95°C for 30 s,
60°C for 1 min, and 72°C for 30 s. The sequences of the primers used areas follows:
GAPDH forward, 5’-AGCAATGCATCCTGCACCACCA-3’; GAPDH reverse, 5’-i
ACAGCCTTGGCAGCACCAGT-3’; Cav3.1 forward, 5’-GCTTCCTGCCTGTTGCCG
AG-3’; Cav3.2 forward, 5’-TGCTCGAACCCTATGCTCC CC-3’; Cav3.3 forward, 5’-
GCAGAATGCCCAACATCGCCA-3’. The Cav3 reverse primer was paired up with the
subunit-specific forward primers and used in the second-step semi-nested Cav3 PCR. All
primers were designed by Primer3 (Rozen & Skaletsky, 2000) using reported cDNA
sequences [DDBJ/EMBL/GenBank accession number: GAPDH, U10983; Cav3.l,
AF027984 (rat); Cav3.2, AF290213 (rat); Cav3.3, AF086827 (rat)] and are intron
spanning. The identities of the PCR-generated fragments were verified by electrophoresis
and sequencing, and the melting temperature of the fragments was used for confirmation
of specific amplification in real-time PCR. The sizes of the amplicons are (bp): GAPDH,
209; Cav3.1, 153; Cav3.2, 166; Cav3.3, 277. No amplification of GAPDH was detected
in no-RT controls from reverse transcription. Negative controls without cDNA template
and positive controls using total hamster spinal cord cDNA template were included in
each PCR. Detection of a transcript was confirmed if the linear phase of the amplification

th th
plot crossed the fluorescence intensity threshold before reaching the 40 (or 45 for

61

GAPDH) amplification cycle (Ct < 40 or 45).

Statistics

Numerical data are presented as mean a: SEM unless noted otherwise. Statistical analysis
was performed using InStat 3 (Ver 3.06, GraphPad Software, Inc., San Diego, CA).
Nonparametric methods were adopted when one or more of the compared groups had less
than five values or failed Kolmogorov-Smirnov normality test. Two groups were
compared using the unpaired t test with Welch correction or Mann-Whitney
(nonparametric) test as appropriate. Comparison of the means among three or more
groups was conducted using one-way ANOVA with Tukey-Kramer multiple comparisons
test or Kruskal-Wallis (nonparametric) one-way ANOVA with Dunn’s multiple

comparisons test. The criterion for statistical significance was a value of P < 0.05.

62

Results

Whole-cell recordings were made from a total of 52 dorsal horn neurons in spinal cord
slices prepared from 35 hamsters. Membrane potential was held at ~60 mV during
voltage-clamp recordings unless otherwise noted for specific experiments. Neurons had
resting membrane potentials of —50 mV or more hyperpolarized under current-clamp

conditions.

Fast and slow T -type calcium currents in spinal dorsal horn neurons

T-type Ca2+ currents (IT) are widely expressed in dorsal horn neurons (Ryu and Randic,
1990; Chapter 2) though the current density varies considerably from cell to cell (Chapter
2). I also observed substantial variations in IT kinetics and surmised that more than one
type of T-channel mediates IT in dorsal horn neurons. In addition, the inactivation of IT
was better described with a double or triple exponential in most of the cases (77%, n =
33), also suggesting that multiple types of T-channels co-exiSt in the same cells. By using
the exponential stripping procedure (see METHODS, Figure 3-1), two IT components were
revealed in 29 (67%) out of 43 cells, while a third component was isolated from four
other neurons, accounting for 9% of the sample. Ten (23%) of the analyzed cells had IT
inactivating in a single exponential term and presumably mediated by only one type of T-
channel. The analysis yielded a total of eighty decay time constants (r), and the
distribution is shown in Figure 3-2A. This frequency histogram suggests that two types of
1T Components are expressed in dorsal horn neurons. A fast-inactivating current
cOmponent (IT,f) had an averaged decay time constant (If) of 12.0 i 1.1 ms (n = 48,

Figure 3-2B), and the slow current component (1T,s) inactivated with a longer decay

63

 

 

 

U)
C
.9
a
(U
E
0
U)
Q
0
t.—
O
L.
w
.0
E
3
z
r(ms)
B C
E12 312
0
E110 2310
$8 $8
96 96
O O
'64 '54
h- 5
£2 32
E
:0 a
2 z0

0 100 200 300 400

If (ms) Ts (ms)

Figure 3-2. Distribution of the decay time constants of IT current components in
Spinal dorsal horn neurons

A, eighty decay time constants (r) of IT current components analyzed from 43 dorsal horn
neurons divide into two groups: If (range, 2.7—38.2 ms) and rs (range, 81—339 ms). B, the
smooth line indicates the best fit of IT distribution to the Weibull probabilitg density
function (r = 0.94). C, 1:5 is also best fitted with the Weibull distribution (r = 0.88). Bar
intervals: 10 ms for A; 2 ms for B; 40 ms for C.

64

time constant (rs) of 177 2 12 ms (n = 32; vs.. rf, P < 0.0001; Figure 3-2C'). The
activation of individual components was assessed by measuring the 10% to 90% rise time
010-90) of the inward current in the cells expressing only one current component, and the
t10-90 for I'Lf and IT,s was 5.2 2 0.8 ms (n = 7) and 14.3 t 3.0 ms (n = 3), respectively.
The amplitude of IT’f was 4.6 x 0.7 pA/pF (n = 48), and IT,s had an average amplitude of
3.5 2 0.4 pA/pF (n =32). Interestingly, two IT; components with two different If were
sometimes observed in the same cells (n = 8) with or without the companion of a slowly-
inactivating current component. Therefore, a number of components, rather than a
number of cells, was given with the presentation of IT; statistics.

Afier establishing a way to analyze IT; and IT,s in dorsal horn neurons, I next
examined the voltage-dependence of steady-state inactivation and activation of these
current components. Inactivation kinetics were analyzed by stepping the membrane
potential to —30 mV every 20 s in 10 mV increments from holding potentials of -120 mV
to -70 mV held for 500 ms. Activation properties were studied by delivering 200—ms
depolarizing voltage pulses (testing steps) in 10 mV increments from —50 mV to -10 mV ’
following a holding potential of —90 mV. When activated from a holding potential more
positive than —70 mV or at a testing step more negative than —50 mV, IT was too small to
be analyzed by the exponential stripping procedure and therefore absent from the results
presented. Average normalized currents (I / Imax) were plotted as a function of holding
potentials for inactivation or testing steps for activation (n = 3 for 1135 n = 2 for IT,s) and
fitted with the Boltzmann equation (I / [max = 1 / {l + exp [(V— V] /2) / k]}) (Figure 3-3).

The half-inactivation potentials for IT; and IT,s were -82 mV and —66 mV, respectively,

65

 

 

 

 

 

 

 

1.0. . 1.0. l
0.8-l L 0.84 -

a: a
E0.6" - 80.6- .
2 0.4. . : 0.4. c
0.24 - 0.2. .
o J I I I I I I I' O “ I I I f r I '—IL
-120-100-80 —60 -40 -20 0 420400 -80 -60 -40 -2o 0

Voltage (mV) Voltage (mV)

Figure 3-3. Voltage-dependent activation and inactivation of 11"; and IT,s in spinal
dorsal horn neurons

Activation (O) and inactivation curves (D) were fitted with the Boltzmann equation (I /
[max = 1 / 1 + exp [(V— V1/2) / k]). V“; and k are shown next to each curve. Error bars

represent SEM (n = 3 for IT; and n = 2 for 1T,s)-

66

It

While the voltage required to activate half-maximal current was -45 mV for 1T,f and —51
mV for IT’S. These results are similar to what has been reported in a study using the same
analysis strategy (Zhuravleva et al., 2001).

Besides variations in current kinetics, 1T,f and 1T,s also showed differential
sensitivity to Ni2+ blockade (Figure 3-4). Activated at —30 mV from a holding potential
of —90 mV, the amplitude of [T,f was reduced by 51% t 5% (n = 5) after 2—4 min
exposure to 200 uM Ni2+, while only 15% :9: 11% (n = 5) of 1T,s was blocked. Another IT
antagonist mibefradil generated comparable inhibitory effect on both current components
(Figure 3-4); Treating dorsal horn neurons with 5 uM mibefradil for 5—8 min decreased

In~ by 52% z 3% (n = 2) and IT’S by 37% i 5% (n = 2).

Rate-dependent activation of fast T -type calcium currents in spinal dorsal horn
neurons

It has been shown that the amplitude of IT in dorsal horn neurons is dependent on the rate
of membrane depolarization (d V/dt), a characteristic contributing to the ability of phasic-
firing neurons to signal rapid membrane depolarization (Chapter 2). However, whether
1T,f and 1T,s behaved in the same way was unclear. IT of three dorsal horn neurons
possessing one IT; component and one 1T,s component were activated by ramping the
membrane potential from a 500-ms hyperpolarizing pre-pulse of —90 mV to -30 mV at
different rates ranging from 400 mV/s to 2400 mV/s. After they were isolated using the
exponential stripping procedure, IT; showed rate-dependent activation, while the

amplitude of 1T,s appeared independent of dV/dt (Figure 3-5). Linear regression analysis

67

 

2+

 

 

 

 

Ni MIB
100- 100.
2\: 80. 80-
i:
.9 60- 3 60- .
f: -t- T
g 40- . 40. o
c I ’
;.-_- 20‘ 20.
° 0
*5 0 q 0 .
8
I. -20‘ -20~
g n = 5 . n = 2
‘40“ I I ‘40‘ l 7
IT r IT,s ITJ IT,s

Figure 3-4. Inhibition of IT; and 11‘s in spinal dorsal horn neurons by T-type Ca2+
channel blockers

Each point was obtained from one dorsal horn neuron treated with either 200 uM Ni2+ or
5 uM mibefradil (MIB). The short horizontal bar indicates the average value of inhibition.

68

300 2 2

  

 

  

 

 

 

 

 

i I r = 0.89
V 250 + TJ
0
'3 200 -
.2: 2
a 150 . r = 0.96
E 2 _
.. 100 -
E
g 50 -
3
o 0 J I I I I
300 1000 2000 3000
2 300 -
a. I
V 250 - T,s
0
'2 200 -
.2:
Ta. 150 . 0 o 2
g A A 0 r2 = 0.02
...- 100 r O U r = Q13
: 0 A r2 = 0.35
2 50 -
L-
3
o o ‘ r I I I I I fit I I
300 1000 2000 3000
Ramp slope (mVls)

Figure 3-5. Rate sensitivity of IT; in spinal dorsal horn neurons

The relationship between the amplitude of IT current components and the ramp slope is
described by a linear function for three dorsal horn neurons. The coefficient of
determination (r2) from linear regression analysis indicates that the IT; amplitude and the

slope are highly correlated, but the amplitude of 1T,s does not parallel the change of the
ramp slope.

69

 

 

indicated that 90% z 3% (n = 3) of the 1T,f amplitude was determined by dV/dt, and the 'i
activation threshold in these three cells, estimated from the x-intercepts, was 533 z 83
mV/s (n = 3), higher than the value observed for the total T current (46 t 21 mV/s)

reported previously (Chapter 2).

Expression of fast and slow T -type calcium currents in spinal dorsal horn neurons with
identified membrane and discharge properties

Having shown that 1"” could be the major current component responsible for the rate-
sensitivity of IT, I next examined the expression of IT’f and IT,s in dorsal horn neurons
with different firing patterns. Non-uniform expression of IT has been reported previously,
with the highest current density in dorsal horn neurons responding to constant membrane
depolarization with one or two action potentials (Chapter 2). It would be interesting to
know if IT; was also expressed more abundantly in these cells, which are preferentially
activated by rapid membrane depolarization (Chapter 2).

Forty dorsal horn neurons used in the IT component analysis above were
categorized based on their firing behavior when activated by depolarizing current steps
applied through the recording pipette. Three distinct discharge patterns were exhibited
among these neurons as described previously (Schneider, 2003, 2005; Chapter 2). Tonic
(T) cells (35%, n = 14) fired action potentials continuously throughout the period of
depolarization (Figure 3-6A b), while the prolonged discharge of delay (D) cells (8%, n =
3) followed a noticeable delay to the first action potential (Figure 3-6A a). Phasic (P) cells
(58%, n = 23) responded to depolarizing current steps with a rapidly adapting firing and

Were further divided into two subgroups based on the degree of spike frequency adaption.

70

A3 b c d

D T P33 Non-P$8

’ litttttgiuttuumtuuiout}
. turnout

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Figure 3-6. Firing patterns and IT of spinal dorsal horn neurons

A, membrane responses of a representative cell from each group (top two traces) to
depolarizing current pulses (lowermost trace) are shown. (Firing pattern groups: D, delay
cells; T, tonic cells; PSB, short burst phasic cells; Non-P53, non-short burst phasic cells)
The arrowheads in a indicate the delay between pulse onset and first action potential in
the delay cell. The transient membrane depolarization characteristic of PSB cells is
indicated by the arrows in c. B, 17 recorded from the same dorsal horn neurons in A was
analyzed by an exponential stripping procedure, revealing one IT; component and one
1T,s component. The fit of 1T,s decay with a single exponential overlays the current trace.
The horizontal dotted line indicates the baseline for inward current measurement, and the
vertical dotted line locates the peak of the current. Current density of IT,f- 2.5 pA/pF for
D; 2.7 pA/pF for T; 13.3 pA/pF for P88; 3.7 pA/pF for non-P33. 17,5 density: 2.5 pA/pF
for D; 3.3 pA/pF for T; 5.4 prF for P88; 2.9 pA/pF for non-P33.

7l

Short burst phasic (P33) cells (n = 7) were characteristic of one or two action potentials
superimposed on a transient membrane depolarization when activated by depolarizing
current injection (Figure 3-6Ac). The remaining non-P53 phasic cells (n = 16) typically

responded with 3 or more action potentials at rapidly diminishing frequency to

depolarizing current pulses, and the number of spikes increased with stimulus amplitude
(Figure 3-6Ad). IT of these cells was activated at -30 mV from a holding potential of —90
mV, and the current components were analyzed by the exponential stripping procedure.
The results of this analysis are summarized in Table 3-1, and examples of IT current
components recorded from representative cells of each firing pattern group are shown in

Fig 3-6B. As expected, PSB cells had two- to six-times higher I’Lf current density than D,

T, and non-P33 cells did, while 1T,s amplitude was fairly comparable among different
firing pattern groups.
I also compared IT; and 1T,s expression in dorsal horn neurons with and without

exhibition of postinhibitory rebound (PIR), a rebound membrane depolarization and

discharge following hyperpolarizing current injection that has been associated with

activation of T-type Ca2+ currents (Russo & Hounsgaard, 1996a; Chapter 2). Both IT;

and IT,s were significantly larger in cells showing PIR than in those neurons without PIR

(Figure 3-7).

72

Table 3-1. Comparison of 1'” and IT,s in spinal dorsal horn neurons with different

firing patterns

 

 

 

Firing
pattern D T P PSB Non‘PSB
n 3 14 23 7 16
15.33226 10.0i0.8 10.3:t 1 0
9.0 :1: 1.5 3 . a: '
Tf (ms) ( ) (16) (26) 9 3 1.2 (9) (17)
IT f density 5.1 3c 1.1 9.0 :t 2.7
’ 1.6 :1; 0.5 (3) 4.2 :t 0.9 16 l 1,2," 3.0 :t 0.3
(prF) ( ) (26) (9) ’ (1 7)
1's (ms) 171 $23 (2) l74i18(8) 180:1:18(20) 167:1:26 (7) 187i25 (13)
IT,s density
l.9:1:0.7 2 3.8i0.9 8 3510.6 20 3.3:t0.8 7 3.6:t0.8 13
(WP) ( ) ( ) < > ( > < )

 

Data presented as mean :5 SEM (cell number or component number for If and [T f

density). 1Significantly different from D cells.

Significantly different from non-P33 cells.

73

Significantly different from T cells.

,8

CD
1

 

 

 

 

 

 

 

 

 

 

 

 

 

 

fl: 8' *-

a.

Z

3 6- 6+

.E‘

‘2 4- 4~

a:

'u

E 2- 21 T

S

U o.
PIR No PIR PIR No PIR
(n=34) (n=10) (n=24) (n=4)

Figure 3-7. Comparison of 1131- and IT,s in spinal dorsal horn neurons with and
without PIR

Bars show the mean current density of each group. Error bars represent SEM. * indicates
the difference between two means is statistically significant.

74

Expression of Ca V3 subunits in spinal dorsal horn neurons exhibiting fast and slow T -
type calcium currents

In an attempt to identify the subunit type of T-type Ca2+ channels that mediate 1T,f and
IT,s in dorsal horn neurons, 1 performed single cell RT-PCR using the cells recorded in
whole-cell configuration (see METHODS). Twenty-nine neurons were successfully
harvested based on the detection of GAPDH cDNA. Cav3 subunit cDNAs were detected
in 12,out of 27 cells expressing IT in voltage-clamp recordings, and the results are
summarized in Table 3-2. IT from three cells was composed of only one IT; component,
and Cav3.1 cDNA was detected in all these neurons. In nine other cells, both IT; and 1T,s
were expressed, but only one species of Cav3 cDNAs was amplified. An example of a
Cav3.l amplification plot resulting from a second-step real-time PCR is shown in Figure
3-8 along with IT recorded from the same cell. No Cav3 subunit cDNA was detected in

two cells in which 1T was absent in voltage-clamp recordings.

75

Table 3-2. Summary of the expression of T-type calcium channels in twelve spinal
dorsal horn neurons

 

 

Cell 11",: In, Cav3.l Cav3.2 Cav3.3
0807074 + - + — —
080708-1 + — + — —
080709-2 + — + - -
080523-1 + + + — -
0805304 + + + — —
080619-1 + + + — —
080620-2 + + + — —
0807184 + + + — -
080716-1 + + — + —
080605-3 + + — — +
0807094 + + - - +
0902124 + + — - +

 

IT was recorded in whole-cell patch clamp experiments, and the current components IT;
and IT,s were assessed by using an exponential stripping procedure. The presence of
Cav3 subunit mRNAs was examined using single cell RT-PCR. “+” indicates detection
of a current component or subunit, while “-” shows no identification of the element.

76

 

> Positive control 1

Single dorsal
horn neuron

. Positive control 2

Fluorescence (an)

 

 

) Negative control

 

 

 

Figure 3-8. Detection of Cav3.1 subunit mRNA from a spinal dorsal horn neuron
expressing 1T with one IT; component

Amplification plots of Cav3.l subunit fragments were generated from a real-time PC R
experiment. Each measurement was averaged from three replicate reactions. The
fluorescence signal normalized to the internal passive reference dye (ROX) is shown, and
plots were base-lined by the Mme software. Hamster spinal cord cDNA with (positive
control 1) and without a previous Cav3 PCR amplification (positive control 2) was used
as template in positive control. cDNA template was replaced with water in negative
control. The threshold fluorescence value (indicated by the horizontal gray line) was
determined by the MxPro software using an amplification-based algorithm, resulting in a
threshold cycle (C1) of 23, 31, and 35 for positive control 1, the single dorsal horn neuron,
and positive control 2, respectively. Inset: the decay of IT (lower trace) recorded from the
same dorsal horn neuron can be fitted with a single exponential with requal to 20 ms.The
voltage command used to activate the current is shown above the trace.

77

Discussion

In this study, I showed that spinal dorsal horn neurons possessed two types of T—type
Ca2+currents, differing in the time course of current decay. The fast inactivating current
IT; was expressed strongly in phasic-firing cells responding to depolarizing current
injection with a brief discharge of one or two action potentials, while the expression of
the slowly decaying current 1T,s was rather uniform among dorsal horn neurons with
different firing patterns. The amplitude of IT,f, but not IT’S, was dependent on the rate of
membrane depolarization. This rate-sensitive property is shown by IT in dorsal horn
neurons (Chapter 2) and is associated with selective activation of phasic-firing cells by
rapid membrane depolarization caused by direct current injection (Schneider, 2003) or
peripheral sensory input (Schneider, 2005). Taken together, the results presented in this
study suggest that expressing a fast-inactivating T-type Ca2+ current contribute to the
ability of dorsal horn neurons to differentiate between afferent sensory inputs encoding
static and dynamic cutaneous stimuli, and single cell RT-PCR experiments suggest that

this current is mediated by Cav3.1 channels.

Fast and slow T-type calcium currents

Two kinetically and pharmacologically distinct T-type Ca2+ currents have been reported
in thalamic neurons by different groups (Huguenard & Prince, 1992; Tarasenko et al. ,
1997; Zhuravleva et al., 1999, 2001; Joksovic et al., 2005). In this study, I showed that a
‘fast’ and a ‘slow’ T-type Ca2+ current coexisted in spinal dorsal horn neurons. These
tW0 current types also had discrete activation and inactivation properties, as well as

different sensitivity to Ni2+ blockade. IT; showed ‘classic’ fast activation and

78

inactivation resembling to T currents recorded in other preparations, and the voltage-
dependence of current kinetics was also similar to what has been reported for native low-
voltage-activated Ca2+ currents (reviewed by Perez-Reyes, 2003). In contrast, 1T,s rose
and decayed more slowly (~3 and ~15 times longer for activation and inactivation,
respectively, compared to Inf), and the steady-state activation and inactivation curves
(Figure 3-3, right) indicates a larger ‘window current’ arising from these slow channels,
similar to a slowly-inactivating T current reported in thalamic local'circuit neurons
(Zhuravleva et al. , 2001). In thalamic cells, window T current underlies membrane
potential bistability, an important element of rhythmogenesis (Crunelli et al. , 2005).
Spinal dorsal horn neurons are also capable of producing oscillatory activity under both
in vivo and in vitro conditions (Sandkiihler & Eblen-Zajjur, 1994; Ruscheweyh &
Sandkt'ihler, 2003; Asghar et al., 2005); however, the contribution of T current to
rhythmic behavior in these cells remains unknown. Alternatively, since most dorsal horn
neurons had fairly comparable 1T,s regardless of their ability to distinguish between
different sensory inputs, this current could play an essential role in basic neuronal
functionality, e. g. dendritic expansion and formation of new synaptic contacts
(Zhuravleva et al., 2001; Lory et al., 2006).

Neuronal T currents show different sensitivity to Ni2+ blockade (Yunker, 2003),
possibly due to heterogeneous Cav3 subunits constituting native T channels (Lee et al.,
1999b). In this study, I also found Ni2+ was more potent in blocking Ipfthan IT,s:
Consistent with what was reported in reticular thalamic neurons (Joksovic et al., 2005).
However, when treated with 100 MM Ni2+, the slowly inactivating T current in thalamic

local circuit cells was much more susceptible to blockade (56% vs. 15% inhibition by 200

79

11M Ni2+ in this study) (Zhuravleva et al. , 2001). Besides an easier access of Ni2+ to
isolated cells used in Zhuravleva and colleagues’ study, this discrepancy in Ni2+
sensitivity between different neuronal cells may also result from tissue-specific post-
translational modifications and regulatory influences from auxiliary subunits. In fact, a
slow T current mediated by cloned Cav3.3 channels also shows varied Ni2+sensitivity
when expressed in different cells (Lee et al., 1999b). On the other hand, mibefradil had
similar effects on IT; and 1T,s in this study, whereas it has been reported that this drug is
equally efficacious on all three Cav3 subtypes, especially when Ba2+ is used as the
charge carrier (Martin et al. , 2000).

Another difference between IT; and 1T,s found in this study is the relationship
between the current amplitude and the rate of membrane depolarization that activated the
current. The [TI amplitude was correlated with the slope of depolarizing membrane
potential ramps, while the magnitude of IT,s appeared to be rate-independent. I attributed
this rate-sensitive property of 1T,f to the same characteristic reported in IT previously
(Chapter 2), with one caveat. The activation threshold of IT by voltage ramps is 46 mV/s
(Chapter 2), but the threshold for IT,f, estimated in the same way, was 533 mV/s. The
discrepancy between these two results implies that IT,s might be also responsive to time-
varying membrane depolarization with a slower range of rate. Given that IT,s had a longer

decay time constant and voltage-dependent inactivation, this idea seems reasonable;

however, further studies are required to validate it.

Molecular identity of fast and slow T-type calcium channels

80

When expressed in HEK 239 cells, Cav3.1 and Cav3.2 recombinant channels generate
T-type Ca2+ currents with classic fast kinetics, while a slowly activating and inactivating
low—voltage-activated current is generated by T channels with cloned Cav3.3 subunit
(McRory et al., 2001; Chemin et al. , 2002). In this study, the results of single cell RT-
PCR experiments suggest that IT,f was mediated by Cav3.1 channels, and this is also
supported by some findings obtained from voltage-clamp recordings. First, the time
constant of IT; decay is consistent with what has been reported for cloned Cav3.1
channels (McRory et al., 2001; Chemin et al., 2002). In addition, the sensitivity of IT,f to
Ni2+ blockade is close to the current produced by the Cav3.l channel isoform (ICso =
~200 uM) (Lee et al., 1999b). However, the 10% to 90% rise time of IT,f activation was
longer in this study and is similar to the Cav3.2 subtype (Chemin et al., 2002). The
potential needed for half-activation and half-inactivation in IT,fIS also comparable to that
of cloned Cav3.2 channels reported by a previous study (McRory et al., 2001). This
contradiction can be explained by the discrepancies between native and cloned T-type
channels that result from post-translational modifications and expression of splicing
variants or auxiliary subunits, as suggested by previous studies (Zhuravleva et al., 2001;
Joksovic et al., 2005). Alternatively, IT; may rise from more than one type of T channels.
Indeed, eight cells in this study had two Ipfcomponents, and Cav3.l and Cav3.2
mRNAs were both detected in cells possessing IT,f. Therefore, this rapidly inactivating
current could be mediated by the T-type Ca2+ channels with Cav3.1 or Cav3.2 subunit.
Unlike IT,f, 1T,s so far has only one possible molecular counterpart being reported. Given

that Cav3.3 mRNA was detected in dorsal horn neurons exhibiting 1T5, it is probable that

81

this current was mediated by Cav3.3 channels, though it cannot be ruled out that a novel

. . . . 2+
channel type also generates a slowly inactlvatmg low-voltage-actrvated Ca current.

Concluding remarks

The present study shows for the first time that two kinetically distinct T-type calcium
currents are expressed in spinal dorsal horn neurons, and the fast current I’Lf contributes
to the ability of some phasic cells to sense rapidly-moving mechanical stimuli applied on
the skin. The role of IT,s in dorsal horn neurons, however, is unclear at this point, though
this slowly inactivating current may underlie the rebound depolarization or firing after the
membrane potential of these cells returned from a hyperpolarized level. This rebound
behavior suggests a mechanism for short-term plasticity based on previous neuronal
activity (Uebachs et al. , 2006), and in the spinal dorsal horn, it has been proposed that the
cells with T-current-mediated rebound bursting are capable of sensing the directionality
of skin stimulation (Russo & Hounsgaard, 1996a). In conclusion, this study provides the
evidence of a rapidly and a slowly inactivating T-type Ca2+ current co-existing in dorsal
horn neurons. These currents may play different roles in the somatosensory fimctions

carried out by these cells.

82

CHAPTER FOUR

SUMMARY AND DISCUSSION

83

1‘57???” . .~ ..

A. Summary of experiments

My study on the role of T-type Ca2+ currents (IT) in the sensory function of the spinal
cord started with a re-examination of 1T in hamster dorsal horn neurons since this current
has been reported in the same area in rats (Huang, 1989; Ryu & Randic, 1990). IT from
hamster cells, isolated using an external solution modified from a previous report (Ryu &
Randic, 1990), had voltage-dependent activation and inactivation (Figure 2-2) and was
blocked by Ni2+ and mibefradil (Figure 2-3) to the extent similar to other neuronal cells
(reviewed by Perez-Reyes, 2003). Although IT was present in the majority of dorsal horn
neurons (> 95%), the current amplitude and kinetics varied from cell to cell. This finding
led to the discovery of differential IT expression among cells exhibiting distinct discharge
properties and two 1T current components using an exponential stripping procedure
(Zhuravleva et al., 1999). IT and its fast inactivating current component, IT,f, could only
be activated by depolarizing voltage ramps with slopes steeper than 46 mV/s and 533
mV/s, respectively, and the current amplitude was related to the rate of membrane
depolarization (Figure 2-4A,B and Figure 3-5). In addition, the highest current density of
IT, as well as 1T,f, was found in phasic-firing dorsal horn neurons that fired one or two
action potentials responding to depolarizing current injection (Tables 2-1 and 3-1). Phasic
cells are preferentially responsive to rapid membrane depolarization that results from
direct current injection (Schneider 2003; Figure 2-5Bc,d) or fast moving mechanical
stimuli applied on the skin (Schneider, 2005). Moreover, the responses of phasic cells to
depolarized membrane potential or ramps were inhibited by Ni2+ and mibefradil (Figure
2-7) at the concentration that also blocked IT and IT; (Figure 3-3). Therefore, the strong

current expression in some phasic cells suggests that IT, or more specifically IT,f,

84

contributes to the ability of dorsal horn neurons to sense dynamic skin stimuli. Single-cell
RT-PCR performed on the cells after whole-cell recordings indicates that IT; is mediated
binav3.l channels. I also found a correlation between high [T density and cells
exhibiting rebound depolarization or discharge following a hyperpolarized membrane
potential. This postinhibitory rebound (PIR) was sensitive to Ni2+, and the activation was
rate-dependent (Figure 2-8), suggesting the involvement of IT. Further analysis indicates
both IT; and a slowly inactivating IT current component IT,s contribute to this
phenomenon. To summarize, my study confirmed the presence of T-type Ca2+ currents in
the dorsal horn neurons of the spinal cord and their participation in a burst of discharge
after membrane hyperpolarization previously reported in turtle dorsal horn neurons
(Russo & Hounsgaard, 1996a). I proposed, based on the data, that this low-voltage-
activated Ca2+ current underlies the differential capability of dorsal horn neurons to
distinguish between static and dynamic skin stimuli by the mechanism of rate-dependent
current activation, and this function is probably carried out primarily by a T-type current

mediated by the Cav3.1 channel isoform.

B. Technical issues in single cell RT-PCR experiments

It has been a decade since three Cav3 subunits were cloned (Cribbs et al. , 1998; Perez-
Reyes et al. , 1998; Lee et al., 1999a); however, the in situ expression of T-type Ca2+
channel proteins has only been reported in a few studies (Latour et al. , 2003; Yunker et
al., 2003; Trevino et al., 2004; Shao et al., 2005; McKay et al., 2006; Molineux et al.,
2006; Zhang et al. , 2006; De Proost et al., 2007; Inagaki et al., 2008), probably owing to

‘in silico’ cloning of these channels (see Chapter 1). Immunolabeling of the Cav3

85

channel isoforms using antibodies developed by either research laboratories or
commercial companies generated unexpected nuclear labeling that cannot be explained
by the current knowledge of the T-type Ca2+ channel as a plasma membrane-associated
protein (McKay et al., 2006; Schneider, unpublished observations). Furthermore, a recent
study demonstrates that an antibody that was anticipated to recognize specifically
recombinant and endogenous Cav3.3 proteins cross-reacts with another neuronal
membrane protein (Chen et al. , 2007), underlining the difficulty of correctly detecting
these rather scarce channels by immunohistochemistry. Since selective blockers for T
'channels are currently unavailable (Huc et al., 2009), in this study, I performed single cell
RT-PCR to examine dorsal horn neurons that expressed IT in voltage-clamp recordings
and determine which Cav3 subunit mediated IT in these cells.

Using a two-step PCR protocol (see METHODS in Chapter 3), I was able to detect
all three subunit transcripts in the spinal dorsal horn, consistent with a previous report
using in situ hybridization (Talley et al., 1999). However, the finding from this analysis is
inconclusive because only one subunit mRNA was identified in nine dorsal horn neurons
that evidently expressed two kinetically distinct T currents. This could be explained by
the degradation of targeted transcripts in the single cell samples owing to the unstable
nature of mRNA molecules. However, some precautions had been taken during prior
electrophysiological recordings. First, 1; limited the recordings to a maximum of 40
minutes since mRNAs are more likely to degrade when they are exposed to an exogenous
solution during a prolonged recording process at room temperature. No difference in fact
was observed in the recording time between the cells with detection of Cav3 mRNAs

(mean 1 SD, 34 :t 4 min, n = 12) and those without identification of these transcripts but

86

expressing IT (35 :t 4 min, n = 15; P = 0.53, unpaired t test), suggesting that setting this
recording time limit is effective. Because mRNAs could also be lost during the process of
cell harvesting, in some experiments I used recording electrodes made from silanized
glass pipettes to prevent mRNAs from adhering to the inner surface of the pipettes (Lin et
al., 2007). Although the detection rate of Cav3 mRNAs was higher when silanized
pipettes were used (58% vs. 33%; Table A-4 in Appendix), contingency analysis
indicates that coating the pipette surface is not the cause of better success in detecting the
transcripts (P = 0.26, Fisher’s exact test).

The inconclusive results could also be attributed to some uncertainty inherent in.
the design of this RT-PCR assay. Hamster spinal cord cDNA was used to optimize the
protocol, but the copy number of each Cav3 subunit cDNA was unknown in these
mixtures. As a result, it is difficult to estimate and therefore improve the efficiency of
individual PCR. Theoretically, even a single copy of cDNA existing in the sample can be
amplified and detected using PCR when given an adequate number of cycles; however, in
reality it does not necessarily happen in this way owing to the Monte Carlo effect2
(Karrer et al., 1995). Therefore, the sparseness of Cav3 subunit transcripts in some single
cell samples could cause the failure of detection. It is also uncertain whether any
interaction, such as competition, occurs in the amplification of three Cav3 subunit

cDNAs from single cell samples during the first step of PCR even though the ‘outer’

 

2 Monte Carlo effect: a limitation inherent in PCR amplification from any complex
template of low abundance due to differences in amplification efficiency among
individual template molecules. As a result, any PCR template diluted past a certain
threshold of copy numbers will have large variations in amplification, and its true amount
will be less likely reflected in the PCR product (Karrer et al., 1995).

87

primers (‘Cav3 forward’ and ‘Cav3 reverse’) had the ability to recognize all three
molecules in the cDNA sample obtained from the spinal cord tissue. Some competition
for amplification among the three subunits seems possible cause because only one
transcript was detected from each dorsal horn neuron, while the first step of PCR was the

determinant of which subunit cDNA that got amplified (Ku & Schneider, unpublished

observations).

C. T-type calcium currents in spinal sensory integration
IT in lowering the threshold for firing

In this study, I hypothesized that IT underlay the selective responses of phasic-
firing dorsal horn neurons to fast membrane depolarizations caused by direct current
injection (Schneider 2003, 2005) or rapidly moving skin stimuli (Schneider 2005). This
idea was prompted by the observation that the amplitude of IT in thalamic sensory
neurons is dependent on the slope of depolarized membrane ramps (Crunelli et al., 1989).
This rate-sensitive activation is also found in IT of dorsal horn neurons (Figure 2-4) and
is attributed to more T-type Ca2+ channels going back to the inactivation state and
becoming unavailable when the membrane potential is depolarized at slower speed
(Crunelli et al., 1989; Gutierrez et al., 2001). In addition, 1T blockers Ni2+ and mibefradil
are able to cease phasic firing and fast-ramp-evoked transient depolarization (Figure 2-7)
at the concentration that also substantially inhibits IT in dorsal horn neurons (Figure 2-3),
further supporting my hypothesis. Meanwhile, these data also suggest that IT may
facilitate the initiation of firing at least in some dorsal horn neurons. It has been proposed

that IT can assist Na+-mediated spike generation in neurons by lowing the threshold for

88

 

firing (Huguenard, 1996). I also noted from current-clamp recordings that a number of
dorsal horn neurons were able to fire action potentials when the membrane potential was
stepped, by direct current injection, to -40 mV or a more hyperpolarized level, at which
no action potential current was recorded in the same cells under voltage-clamp mode.
This phenomenon will be referred to as ‘low-threshold-current-mediated action
potentials’ (LTAPs) in this document, and I speculated that IT also contributed to LTAPs
because of the subthreshold current activation. I think what happened was that the fast,
kinetics of IT generated a depolarizing transient that briefly “leaped” the membrane
potential to pass the threshold for voltage-gated NaJr current activation in the beginning
of an injected current pulse. This caused the cell to fire and left a rather hyperpolarized
membrane potential observed at the end of the current pulse. This speculation was
supported by a higher IT density in dorsal horn neurons that were able to produce LTAPs
than those without this capability (8 1 2 pA/pF vs. 3 1 1 pA/pF; Table A-5 in Appendix).
When applied on phasic cells (presumably IT-rich dorsal horn neurons), Ni2+ (200 11M)
increased the threshold for firing (Ku & Schneider, unpublished observations). In
addition, the observation of LTAPs was strongly associated with a postinhibitory rebound
behavior in dorsal horn neurons (P = 0.0033, Fisher’s exact test; Table A-6 in Appendix),
also suggesting the involvement of IT as a common mechanism. A similar idea has been
reported in amphibian olfactory receptor cells, in which IT is attributed to enhanced odor
sensitivity by lowering the threshold for spike generation (Kawai & Miyachi, 2001).

In this study, not all phasic-firing dorsal horn neurons had the ability to produce
LTAPs, consistent with the observation that phasic cells are a rather diverse group

(Chapter 2). These LTAP-generating phasic cells may have lower threshold for

89

P’fifil—r

mechanical skin stimuli than those without LTAPs. The differential capability of
producing LTAPs, and essentially the non-uniform expression of IT, among phasic cells
could be the reason why different conclusions were drawn from previous studies (Lopez-
Garcia & King, 1994; Schneider, 2005) regarding the functional role of phasic firing
dorsal horn neurons in sensory transmission (nociceptive specific vs. low threshold
mechanical). IT may, in turn, serve as a molecular marker for the dorsal horn neurons that
are involved in processing sensory inputs from a certain type of primary sensory neurons,
e. g. low threshold mechanoreceptors. It has been reported that large 11- mediated by the
Cav3.2 channel subtype is uniquely expressed by D-hair receptors3 and thus a useful
marker for identification of these sensory cells (Heppenstall & Lewin, 2006). In D-hair
receptors, IT is proposed to contribute to the high sensitivity of these mechanoreceptors
to slowly moving stimuli by “amplifying” the receptor potential and eventually leading to
generation of action potentials (Shin et al., 2003; Dubreuil et al., 2004). Since it is
possible that the receptor potentials evoked by slow skin stimuli are initiated by some
mechanosensitive channels other than T-type Ca2+ channels (Heppenstall & Lewin,
2006), the function of IT proposed in D-hair receptors does not necessarily contradict
what was suggested by this present study for spinal dorsal horn neurons. However, it is
also noteworthy that some small dorsal root ganglion cells also have large 1T that

prominently arises from Cav3.2 channels and are C-thermonociceptors or C-

3 D-hair receptors: mechanoreceptors that have an extremely low threshold, large
branching receptive field in the skin, and conduction velocities of A6 fibers. Their name
comes from the observation in cats that these receptors can be selectively activated by
moving small sinus or down hairs in the skin (Heppenstall & Lewin, 2006).

90

mechanonociceptors (Coste et al., 2007), indicating a potential role of IT in pain
transmission (see Jevtovic-Todorovic & Todorovic, 2006 for a review).
IT potentiation

Although this and previous studies have suggested that IT play some role(s) in
spinal sensory processing, a basic, but not so easy, question that has been left unanswered
is whether T-type Ca2+ channels are able to function in a physiological condition. Using
the data presented in this study as an example, the resting membrane potential of dorsal
horn neurons is usually between -55 mV and -70 mV (Table 2-1), at which, by
estimation no more than 10% of T-channels are available (dc-inactivated) for activation
(the inactivation curve in Figure 2-2C). To remove the inactivation, substantial and
prolonged membrane hyperpolarization is required4, making it uncertain whether dorsal
horn neurons exhibit physiologically significant 17 in vivo. A possible answer to this
question has recently been approached in sensory thalamocortical neurons, in which an
ATP-dependent phosphorylation increases IT amplitude and accelerates its inactivation
(Leresche et al., 2004). This potentiating process occurs only when T-charmels are
inactivated and therefore is able to facilitate IT generation at resting membrane potential
(where the majority of T-channels are inactivated) by reducing the number of channels
required to recover from inactivation to produce a substantial IT (Leresche et al., 2004).
IT potentiation operates under the conditions in which few T-channels are available for

activation and only weak depolarization is required (Bessa’fh et al., 2008), making it more

 

4 For native T-type Ca2+ channels, to fully recover from inactivation requires a
hyperpolarization to —90 mV for 100 ~ 3300 ms, depending on the type of different
neuronal cells (Perez-Reyes, 2003).

91

 

likely a functional mechanism in vivo. Furthermore, IT potentiation also enhances the
occurrence and temporal precision of synaptically evoked burst firing in sensory
thalamocortical neurons (Bessa'r'h et al., 2008). This outcome is especially favorable to
phasic cells, if IT potentiation turns out also to modulate firing in dorsal horn neurons,
since the discharge property of phasic cells is suited for coincidence detection, for which
timely responses are essential (Prescott & De Koninck, 2002).
IT de-inactivation

Although the potentiation mechanism provides a possibility for 1T to function
under physiologically relevant conditions, a preceding hyperpolarization is still an
imperative to de-inactivate the channels and make them available for activation. In vivo,
hyperpolarized membrane potential can result from inhibitory postsynaptic potentials
(IPSPs) or afterhyperpolarization (AHP). Low-threshold Ca2+ current has been proposed
to contribute to fast afterdepolarization (ADP) in spinal lamina II neurons, which
possibly follows, but may not be triggered by the prior AHP (Yoshimura & Jessell,
1989). The slower activation of the current, in contrast to spike-generating Na+ current,
probably causes the presence of the Ca2+-mediated transient depolarization as an
afierpotential, which is still “conditioned” by resting membrane potential before spike
generation (Yoshimura & Jessell, 1989). Although it appears unlikely that T-channels in
dorsal horn neurons are “primed” by AHP, it is still possible that IT-mediated ADP
influences the response of these neurons to synaptically evoked events that occur
concomitantly (Yoshimura & Jessell, 1989).

IT, activated by subthreshold postsynaptic potentials, may contribute to synaptic

integration even before the action potential is generated (Reyes, 2001). In that case,

92

Wit .. _fl-I

IPSPs are likely to be the source of membrane hyperpolarization removing inactivation of
T-channels. IPSPs in dorsal horn neurons can be evoked by volleys in cutaneous afferent
fibers (Sahara et al., 1990; Schneider, 1992) as well as mechanical stimuli on the skin
(Schneider, 2005). They are seemingly polysynaptic since there is no evidence for
inhibitory afferent inputs (Willis & Coggeshall, 2004), and may come from inhibitory
intemeurons in the adjacent or more distant dorsal horn area (Schneider, 1992).
Functional inhibitory connections haveibeen reported between dorsal horn neurons (Lu &
Perl, 2003; Schneider, 2008), further supporting this idea. Stimulating cutaneous afferent
fibers evokes both IPSP-EPSP and EPSP-IPSP sequences in dorsal horn neurons (Sahara
et al., 1990; Schneider, 1992). Given the biophysics of T-channels, the former would
more likely activate IT than the latter does. This is reminiscent of the function of IT
proposed previously for dorsal horn neurons, which is to sense the directionality of skin
stimuli (Russo & Hounsgaard, 1996a). This seems possible because inhibitory receptive
fields (RFs) in dorsal horn neurons, when identified using innocuous cutaneous stimuli,

e. g. brush strokes and hair movements, are located adjacent or eccentric to excitatory
ones (Hongo et al., 1968; Brown et al., 1987; Schneider, 1992, 2005). This spatial
organization of RFs suggests a mechanism for detecting the moving direction of a tactile
stimulus on the skin (Hongo et al., 1968). Due to the prerequisite of membrane
hyperpolarization for priming the channels, IT in dorsal horn neurons could only be
activated when a stimulus moves from an inhibitory skin RF to an excitatory one, but not
in the opposite direction (Figure 4-1A). IT therefore would be a plausible candidate for
the underlying ionic mechanism of direction sensitivity. It has been reported in cat

thalamic ventral posterolateral nucleus, a major relay of somatosensory inputs, that some

93

Receptlve Primary Dorsal horn neurons
Fields sensory

neurons a /
e l: .

9,.) :4

 

 

 

 

B Receptlve Primary Dorsal ho'rn neurons
Fields sensory a
neurons /
. (’9 11 ‘
Q 4

 

 

 

 

. p b
\ l4 _llml
b / 0

Figure 4-1. A hypothetical mechanism for dorsal horn neurons to distinguish the
direction of a moving skin stimulus.

This mechanism can apply to either phasic (P, shown in A) or tonic (T, shown in B) cells,
with an assumption that these cells express substantial IT. To Simplify, only two primary
sensory neurons (X, Y) are shown here to project to dorsal horn neurons. Neuron X with
a direct connection with the cell recorded and therefore forms an excitatory receptive
field (in gray shade) for the dorsal horn neuron. On the other hand, Neuron Y projects to
an inhibitory dorsal horn intemeuron (I), which synapses with the P or T cell. As a result,
IPSPs will be produced in the cell recorded when Neuron Y is activated; therefore, its
receptive field (in white) is inhibitory for the P or T cell. The direction of moving skin
stimuli are indicated by arrows next to the receptive fields, and the membrane responses
evoked are shown next to the dorsal horn neuron, indicated by a corresponding
alphabetical designator (a, b). See text for details.

94

cells can only be excited by the stimulus of light pressure moving proximally in the long
axis of an opposite forelimb (Gordon & Manson, 1967). IT potentially can contribute to
this phenomenon since its expression has been reported in this region (Leresche et al.,
2004; McKay et al., 2006). In the dorsal horn of the spinal cord, phasic firing neurons are
especially responsive to moving skin stimuli (Schneider, 2005) and express large IT
(Chapter 2), so these cells could be direction sensitive. However, tonic cells may also
have this ability because they are activated by brush strokes applied on the skin
(Schneider, 2005), and some of them have substantial IT (Figure A-2) as well. Instead of
differentiating the orientation of skin stimuli by firing or not as phasic cells would do
(Figure 4-1A), tonic cells could generate either tonic firing for a stimulus starting in their
excitatory RF or a short burst of action potentials for a brush stroke moving from an
inhibitory RF to an excitatory one, similar to PIR (Figure 4-lB). Of course, more
experiments using an in vivo preparation or an in vitro one with intact RFs in a limb
(Lopez-Garcia & King, 1994) or a skin patch (Schneider, 1992, 2005) attached to the
spinal cord are needed to test this hypothesis.
IT activation

After T-type Ca2+ channels are de-inactivated, they are available for activation.
However, in a cell, how many channels will actually be activated by an excitatory event,
either an EPSP or a compound of EPSPs, is determined by how fast the membrane
potential is depolarized by the event. As a result, the amplitude of IT (Crunelli et al.,
1989; Chapters 2 and 3) and the IT-mediated transient depolarization or spike (Gutierrez
et al., 2001) is related to the rate of membrane depolarization. Fast and slow EPSPS can

be generated by activating postsynaptic ionotropic and metabotropic receptors,

95

 

respectively (Gutierrez et al., 2001). In the spinal dorsal horn, both types of receptors for ‘ _
glutamate, the major neurotransmitter producing EPSPs, are evident for synaptic
transmission (Willis & Coggeshall, 2004). However, it is currently unknown if EPSPs
mediated by different types of receptors would affect IT activation in the neurons of this
region. In addition, the rate of membrane depolarization is not the only determinant of the
current activation. It has been reported that IT-mediated low-threshold spike can be
evoked even by very slow voltage ramps when leak K+ conductance is blocked
(Gutierrez et al., 2001). This may provide another explanation for the rate insensitivity of
tonic firing dorsal horn neurons found in this and previous studies (Schneider, 2003,
2005). Tonic cells have a two-times higher input resistance than phasic cells, suggesting
less leak K+ current, and a more depolarized resting membrane potential supports this
conclusion (Table 2-1). Therefore, instead of expressing less IT, it is the scarcity of leak
K+ current in these cells that is unable to “mask” IT at the resting membrane potential

and makes tonic cells less sensitive to the rate of membrane depolarization. This idea can
be tested by examining their rate sensitivity in the presence of IT blockers, e. g. N12+. The
firing threshold of tonic cells should increase if a constant window IT is the cause.
Alternatively, without substantial window IT, slow membrane depolarization can still
activate IT if T-channels in dorsal horn neurons are subject to the potentiation mechanism
reported in sensory thalamocortical cells (see IT potentiation above; Leresche et al.,
2004; Bessa‘rh et al., 2008). In that case, IT in tonic cells could be better potentiated
because of more depolarized resting membrane potential, and therefore would have lower

ramp threshold for activation.

96

III-"P—T' 1?:-

Nevertheless, the data presented in this study suggest that IT, possibly its fast
inactivating component, contributes to the rate sensitivity of phasic firing dorsal horn
neurons, which have been proposed to receive more sensory inputs from rapidly adapting
(RA) mechanoreceptors than from slowly adapting (SA) ones (Schneider, 2005) (Figure
4-2A). This idea is further supported by the finding that RA receptors, when activated by
ramp stimuli, respond to increasing velocities of skin indentation with shortening
interspike intervals (Knibestol, 1973), which could, by temporal summation of EPSPS,
lead to a faster depolarization in the postsynaptic membrane. The resulting IT activation
could further depolarize the membrane potential and lead to discharge of the phasic cell
(Figure 4-28). Therefore, by expressing 1T, phasic firing dorsal horn neurons may act as a
‘high-pass filter’, which only allows the transmission of sensory stimuli with a certain
level of velocities, and the ‘cutoff’ level (threshold) could be determined by the current
density as well as the channel subtype expressed.

IT in synaptic function

It has been demonstrated in this study that most dorsal horn neurons possess a
slowly inactivating IT component (Chapter 3), which has been reported to be the
“dendritic” type in thalamic neurons (Zhuravleva et al. , 2001; Joksovic et al., 2005; but
also see Huguenard & Prince, 1992). Currently there is no agreement regarding the
subcellular distribution of the three known Cav3 subtypes, and the localization is
unlikely the same in different neural tissues (McKay et al., 2006). However, axodendritic
contacts formed between primary afferent fibers and dorsal horn neurons as well as
between intemeurons in the region (Schneider et al., 1995; Schneider, 2008) suggest the

dendritic distribution of (at least some) T-channels and their role in synaptic transmission

97

A Skin Primary sensory neurons Dorsal horn

  

I IT, f-mediated channels (Cav3.1 andlor Cav3.2?)
A IT’S-mediated channels (Cav3.3?)

3 Skin Sensory nerve Current
Indentation recording clamp

h/
945 ,
e /

 

 

4

 

Figure 4-2. T-type Ca2+ channels in spinal sensory integration

A, a summary diagram showing that tonic firing dorsal horn neurons (T) and short burst
phasic cells (P53) receive differential sensory inputs from rapidly (RA) and slowly
adapting (SA) mechanoreceptors (Schneider, 2005) and also exhibit different levels of
1T,f, probably mediated by the Cav3.1 and/or Cav3.2 channel subtypes (Chapter 3).
Higher density of 1qu in PSB cells may contribute to their sensitivity to rapidly moving
skin stimuli, and a possible mechanism shown in B. B, the RA receptor responds to a fast
skin indentation with short interspike intervals, which result in a rapid membrane
depolarization (a temporal summation of EPSPs) in the postsynaptic PS B cell (upper).
The rapid depolarization activates T-type channels, and a IT-mediated transient further
depolarizes the membrane and eventually leads to an action potential. When the skin is
indented at a reduced velocity, T-type channels are inactivated by a slowly rising
depolarization, and therefore neither IT-mediated transient nor action potentials are
generated (lower).

98

 

and integration within the dorsal horn network. In cortical and hippocampal pyramidal
neurons, IT has been reported to amplify synaptic input by enhancing the amplitude of
EPSPs (Magee & Johnston, 1995; Magee et al., 1995; Gillessen & Alzheimer, 1997;
Urban et al,1998; de la Pefia & Geijo-Barrientos, 2000). This process may counteract the
passive attenuation of EPSPs from synapses in distal dendrites and increase the chance of
integration when these EPSPs travel toward the soma (de la Pefia & Geijo-Barrientos,
2000). Owing to the transient nature of the channel activation, the enhancement by IT
could be more effective for spatial than for temporal summation (de la Pefia & Geijo-
Barrientos, 2000). IT therefore could assist phasic dorsal horn neurons in the intensity
coding of flutter-vibration sensation, a plausible function for these cells requiring
activation of a group RA receptors (Johnson, 1974) and spatial summation. By contrast,
EPSPs in tonic dorsal horn neurons are amplified and prolonged by persistent Na+ and
Ca2+ currents and consequently tend to be summed temporally (Prescott & De Koninck,
2005). IT may also play a role in synaptic plasticity. It has been reported in spinal lamina
I neurons that IT activation is required for the induction of long-term potentiation at C-
fiber synapses (Ikeda et al., 2003; Ikeda et al., 2006). This may be an underlying
mechanism of hyperalgesia caused by natural noxious stimulation including

inflammation, trauma, and nerve injury (Sandkfihler, 2009).

D. Remarks on the sensory functions of the spinal dorsal horn
It has been a consensus that dorsal horn neurons not only‘transmit but also integrate
various sensory inputs from primary afferents, although the exact mechanisms have yet to

be elucidated. Given the diversity of intrinsic properties found in dorsal horn neurons, it

99

a. by ..v _—_—_

would be no surprise if these neurons turn out to contribute to sensory processing in a

 

er

number of different ways. Several sensory functions have been suggested for dorsal horn
neurons in this and previous studies. The most intuitive and frequently studied function is
to encode stimulus features. For example, Prescott and De Koninck (2002, 2005)
proposed that tonic neurons, which fire repetitively during stimulation and exhibit
prolonged EPSPs, are suited to encode stimulus intensity, while phasic firing neurons
may act like coincidence detectors, relaying sensory inputs with temporal precision. In
this study, I suggest that phasic cells may also signal the velocity of moving tactile
stimuli, while some dorsal horn neurons could be capable of encoding in their discharge
the direction of a stimulus moving across the skin (see Section C). In addition to
extracting a specific feature from a stimulus, dorsal horn neurons may sometimes amplify
the inputs by wind-up. This is an activity-dependent and progressive increase in the
responses to repetitive stimuli, and denotes a potentially noxious event (Coste et al.,
2008). By contrast, dorsal horn neurons can also function by blocking the transmission of
some sensory information. For example, phasic cells, as mentioned previously, may act
like a high pass filter by which the inputs from slow skin indentation or movement are
screened out. Other dorsal horn neurons may “ignore” the stimuli of one type (modality)
when those of another type (modality) happen simultaneously. Lu and Perl (2003)
reported an inhibitory connection between spinal lamina II neurons receiving direct
excitatory input from different afferent C-fibers and proposed that this neural module

may represent nociceptive input being suppressed by innocuous impulses. These dorsal

horn neurons analyze and decipher the significance of peripheral inputs before

100

transmitting the flow of sensory information to higher centers and are therefore often

subject to the modulation of descending control from the brain (Heinricher et al., 2009).

E. Future studies

T-type Ca2+ channel research is a newly emerging but rapidly developing field,
with more than a hundred related articles published each year for the last decade (N ilius
et al., 2006). Many, of the studies were conducted in neurons (N ilius et al., 2006), and the
role of IT in mechanosensation is also recently revealed (Heppenstall & Lewin, 2006).
However, there have been few studies of IT in the spinal dorsal horn, and a general
description of IT-expressing dorsal horn neurons was lacking before this study. Using a
standardized voltage protocol, I examined IT in superficial and deep dorsal horn neurons
whose intrinsic properties were also assessed. The finding that 1T was ubiquitously
expressed in dorsal horn neurons has provoked a great interest in the function(s) of the
current in spinal sensory integration. I have made several conjectures about the functions
throughout this chapter, which can serve as hypotheses for the future studies of this topic.
Some of the proposed functions imply certain subcellular distribution of T-channels. This
can be examined by using immunohistochemistry techniques (McKay et al., 2006) or/and
performing cell-attached patch on different cellular locations, e.g. dendrites and the soma
(Joksovic et al., 2005). The former procedure can also unveil the subtype(s) of the T-
Channel in dorsal horn neurons while the latter method will provide a better estimation on
the channel density. To determine the time point at which IT participates in the firing of
dorsal horn neurons, the utilization of the action potential clamping technique (see Box 2

in Bean, 2007 for an overview) would be appropriate. The idea of IT facilitating action

101

 

 

potential generation can also be tested using this approach. These preceding experiments
can be conducted in spinal cord slice preparations. The physiological role of IT in dorsal
horn neurons, however, needs to be confirmed in in vivo preparations or in vitro ones in
which natural stimulation can be applied, e. g. a hemisected spinal cord-skin patch
preparation (Schneider 1992, 2005). It is also noteworthy that some genetic approaches,
such as gene knock-out by gene targeting or gene knock-down by antisense techniques,
have recently been developed and utilized to delete or block the function of a specific '1‘-
channel subtype (reviewed by Shin et al., 2008). In the spinal dorsal horn, these genetic
tools can be used in combination with electrophysiological recording to study the
function of IT mediated by individual Cav3 subunits. The results of these researches will
not only advance our knowledge of sensory processing in normal physiological
conditions but also benefit future clinical practice as T-type Ca2+ channels have become

an emerging therapeutic target for at least absent seizures and pain management (Shin et

al., 2008).

102

APPENDIX

SUPPLEMENTAL DATA

103

 

Table A—1. Comparison of action potentials from neurons in slices prepared using
sucrose- vs. glycerol-based dissection solution

Fix n ‘I“ '

 

 

Dissection solutions Sucrose-based Glycerol-based pl
Number of cells 20 20
Resting Vm (mV) ’61 i 8 ‘64 i 5 0.29
Action potential amplitude (mV) 71 :1: 12 75 1 10 020
Action potential half-width (ms) 1.2 :l: 0.3 1.1 1 0,3 0,13

 

Data presented as mean 1 SD. 1 Based on unpaired t test.

Action potentials were generated by direct injection of depolarizing current pulses
through the recording pipette. The measurements were made and averaged from three
‘first’ action potentials obtained from each cell.

104

Table A-2. Comparison of inward currents recorded from neurons in slices
prepared using sucrose- vs. glycerol-based dissection solution

 

 

Dissection solutions Sucrose-based Glycerol-based p1
Number of cells 20 20
Half-width (ms) 41 1 36 33 1 25 0.92
Peak current amplitude (pA) 176 1 169 191 1 89 0.19
Time to peak (ms) 16 1 17 10 1 4 0.49

 

Data presented as mean 1 SD. 1 Based on Mann-Whitney test.

Inward currents were recorded in an external solution suitable for isolating T-type Ca2
currents (see METHODS in Chapter 2 or 3) and activated by a 200-ms voltage step to -30
mV from a 500-ms holding potential of -90 mV. Three current traces from each cell were
low-pass filtered at 500 Hz (Gaussian) and averaged before the measurements were made.

+

105

Table A-3. Comparison of IT from neurons in superficial vs. deep spinal dorsal horn

 

 

Laminae I/II III/IV pl
Number of cells 15 28
IT amplitude (pA) 135 1 20 140 1 20 0.74
IT density (pA/pF) 6 1 1 7 1 l 0.46

 

Data presented as mean 1 SEM. 1 Based on Mann-Whitney test.

The location of cells was determined under light microscopy (100x), based on the
residence of the cell soma relative to spinal lamina 11 (see Figure A-lA). IT was recorded
using an IT-isolating external solution (see METHODS in Chapter 2 or 3) and activated by '
a 200-ms voltage step to -30 mV from a 500-ms holding potential of -90 mV. Three to
five current traces from each cell were low-pass filtered at 500 Hz (Gaussian) and
averaged before the measurements were made. The current density was obtained by

dividing the peak amplitude by whole-cell capacitance, an indicator of cell size and read
directly from the potentiometer during patch clamping.

106

Table A-4. A contingency table showing the relationship between the use of silanized
pipettes and the detection of Cav3 mRNA in single cell RT-PCR experiments

 

 

 

 

. . . Non-silanized
S l
r anlzed pipettes pipettes Total
Detection of Cav3 7 5 12
mRNA
No detection 5 10 15
Total 12 15 27
Detection rate 58% 33%

 

 

 

Fisher’s exact test indicates no association between these two variables (P = 0.26). All 27

 

dorsal horn neurons presented here expressed some level of T-type Ca + currents in

voltage-clamp recordings.

107

Table A-5. Comparison of IT and intrinsic membrane properties in spinal dorsal

horn neurons with and without the capability of generating ‘low-threshold-current-
mediated action potentials’ (LTAPs)

 

 

Generation of LTAPs Yes No
Number of cells 20 12 P
Threshold (mV) —46 1 1 —34 1 1 < 0.00011
Resting Vm (mV) -66 i 1 -56 i 2 0.00072
Rin (QM) 297 1 33 680 1 105 0.00412
17 density (pA/pF) 8 i 2 i 3 =1: 1 0.00173

 

Data presented as mean31 SEM. ] Based on unpaired t test. 2 Based on unpaired t test
with Welch correction. Based on Mann-Whitney test.

The threshold (of firing) was determined by measuring the membrane response of cells to
a 3-s depolarizing current pulse of the lowest intensity at which they fired. The
measurement was made during the last half second of the current pulse when action
potentials were not present. Cells were designated to have the capability of generating
LTAPs if the threshold was at -40 mV or a more hyperpolarized level. The measurement
of other variables was described in Chapter 2.

108

Table A-6. A contingency table showing the relationship between the exhibition of
postinhibitory rebound (PIR) and the ability to generate low-threshold-current-
mediated action potentials (LTAPs) in twenty-eight spinal dorsal horn neurons

 

 

 

PIR No PIR Total
LTAPs 1 7 1 1 8
No LTAPs 4 6 10
Total 21 7 28

 

 

 

Fisher’s exact test indicates a significant association between these two variables (P =
0.0033). The designation of possessing PIR and LTAPS was described in Chapter 2 and
Table A-5, respectively.

109

 

Figure A-l. Images of the dorsal horn and its neuron recorded in this study

A shows the magnified image (100x) of a transverse slice prepared from the lumbosacral
section of a hamster spinal cord. In this study, whole-cell patch-clamp recordings were
performed in the cells primarily located at laminae I to IV of the dorsal horn. Lamina 11
(L11) is the translucent band on the dorsal part of the horn, while laminae III and IV
(LIII/LIV) looks darker and cannot be distinguished from each other under light
microscopy. Orienting labels: D, dorsal; V, ventral; L, lateral; M, medial. B shows the
magnified image (400x) of a dorsal horn neuron after the whole-cell configuration was
formed. The same cell was harvested after termination of recording for single cell RT-
PCR analysis. The cell was aspirated into the recording pipette and lifted away fi'om the
slice surface as shown in C. The arrow indicates part of the cell that was not sucked into
the pipette. In order to harvest the whole cell, the pipette tip was broken and preserved
along with the cell in a mixture containing the reagents for reverse transcription. Scale
bars: 0.2 mm forA; 20 pm for B and C.

110

 

 

 

 

5' n=16
4‘ Tcellsflllllllllt
31 r—1
.1 2:2 Vs
m
= 0- .L
33' i: "=12 N P P II J, L
a 3_ OH' SB C95
1=L
: 2‘
3 “Jill
a 0" fl I I r
2 51 -7
"- 34 "‘ PSBceIIsL—c
.4 .r——-1_
2.
1..
0‘ I I I I L

 

0 510152025303540
IT current density (pAlpF)

Figure A—2. Expression of IT in phasic- and tonic-firing neurons

Histograms comparing IT density for phasic- and tonic-firing dorsal horn neurons. Short
burst phasic cells (P33) had significantly greater IT than tonic (T) cells or other phasic
(Non-P53 P) cells did. Current density was calculated by dividing IT amplitude by
whole-cell capacitance. Insets show example voltage responses (upper traces) to
depolarizing current pulses (lower traces).

lll

REFERENCES

112

References

Alvifia K, Ellis-Davies G & Khodakhah K. (2009). T-type calcium channels mediate
rebound firing in intact deep cerebellar neurons. Neuroscience 158, 635-641.

Arikkath J & Campbell KP. (2003). Auxiliary subunits: essential components of the
voltage-gated calcium channel complex. Curr Opin Neurobiol 13, 298-307.

Armstrong C & Matteson D. (1985). Two distinct populations of calcium channels in a
clonal line of pituitary cells. Science 227, 65-67.

Asghar AUR, Cilia La Corte PF, LeBeau F EN, Dawoud MA, Reilly SC, Buhl EH,
Whittington MA & King AE. (2005). Oscillatory activity within rat substantia
gelatinosa in vitro: a role for chemical and electrical neurotransmission. J Physiol
562, 183-198.

Bean B. (1985). Two kinds of calcium channels in canine atrial cells. Differences in
kinetics, selectivity, and pharmacology. J Gen Physiol 86, 1-30.

Bean BP. (2007). The action potential in mammalian central neurons. Nat Rev Neurosci
8, 451-465.

Bessa'lh T, Leresche N & Lambert RC. (2008). T current potentiation increases the

occurrence and temporal fidelity of synaptically evoked burst firing in sensory
thalamic neurons. Proc Natl Acad Sci U S A 105, 11376-11381.

Bossu JL, Feltz A & Thomann JM. (1985). Depolarization elicits two distinct calcium
currents in vertebrate sensory neurones. Pflugers Arch 403, 360-368.

Brown AG. (1973). Ascending and long spinal pathways: dorsal columns, spinocervical
tract and spinothalamic tract. In Handbook of Sensory Physiology Volume II:
Somatosensory System, ed. I ggo A, pp. 315-338. Springer-Verlag, Berlin,
Heidelberg, New York.

Brown AG, Brown PB, Fyffe RE & Pubols LM. (1983). Receptive field organization and

reSponse properties of spinal neurones with axons ascending the dorsal columns
in the cat. J Physiol 337, 575-588.

113

Brown AG & Fyffe RE. (1981). Form and function of dorsal horn neurones with axons
ascending the dorsal columns in cat. J Physiol 321, 31-47.

Brown AG, Koerber HR & Noble R. (1987). An intracellular study of spinocervical tract
cell responses to natural stimuli and single hair afferent fibres in cats. J Physio]
382, 331-354.

Carbone E & Lux HD. (1984). A low voltage-activated, fully inactivating calcium
channel in vertebrate sensory neurones. Nature 310, 501-502.

Carbone E & Lux HD. (1987). Kinetics and selectivity of a low-voltage-activated
calcium current in chick and rat sensory neurones. J Physiol 386, 547-570.

Catalano JV & Lamarche G. (1957). Central pathway for cutaneous impulses in the cat.
Am JPhysio] 189, 141-144.

Chemin J, Monteil A, Perez-Reyes E, Bourinet E, Nargeot J & Lory P. (2002). Specific
contribution of human T-type calcium channel isotypes (0110, am and 011 l) to
neuronal excitability. J Physiol 540, 3-14.

Chemin J, Traboulsie A & Lory P. (2006). Molecular pathways underlying the
modulation of T-type calcium channels by neurotransmitters and hormones. Cell
Calcium 40, 121-134.

Chen Y, Sharp AH, Hata K, Yunker AMR, Polo-Parade L, Landmesser LT & McEnery
MW. (2007). Site-directed antibodies to low-voltage-activated calcium channel
Cav3.3 (ALPHAl 1) subunit also target neural cell adhesion molecule-180.
Neuroscience 145, 981-996.

Chorev E, Manor Y & Yarom +Y. (2006). Density is destiny - on the relation between
quantity of T-type Ca channels and neuronal electrical behavior. CNS Neural
Disord Drug Targets 5, 655-662.

Chung JM, Kenshalo DR, Jr., Gerhart KD & Willis WD. (1979). Excitation of primate
spinothalamic neurons by cutaneous C-fiber volleys. J Neurophysiol 42, 1354-
1369.

Coste B, Crest M & Delmas P. 2007). Pharmacological dissection and distribution of

NaN/Nav1.9, T-type Ca + currents, and mechanically activated cation currents in
different populations of DRG neurons. J Gen Physiol 129, 57-77.

114

Coste J, Voisin D, Luccarini P & Dallel R. (2008). A role for wind-up in trigeminal
sensory processing: intensity coding of nociceptive stimuli in the rat. Cephalalgia
28, 631-639.

Craig AD & Kniffki KD. (1985). Spinothalamic lumbosacral lamina I cells responsive to
skin and muscle stimulation in the cat. J Physiol 365, 197-221.

Cribbs LL, Lee JH, Yang J, Satin J, Zhang Y, Daud A, Barclay J, Williamson MP, Fox
M, Rees M & Perez-Reyes E. (1998). Cloping and characterization of alH from
human heart, a member of the T-type Ca channel gene family. Circ Res 83,
103-109.

Crunelli V, LightowleLr S & Pollard CE. (1989). A T—type Ca2+ current underlies low-
threshold Ca potentials in cells of the cat and rat lateral geniculate nucleus. J
Physio] 413, 543-561.

Crunelli V, Toth TI, Cope DW, Blethyn K & Hughes SW. (2005). The 'window' T-type
calcium current in brain dynamics of different behavioural states. J Physiol 562,
121-129.

de la Pefia E & Geijo-Barrientos E. (2000). Participation of low-threshold calcium spikes

in excitatory synaptic transmission in guinea pig medial frontal cortex. Eur0pean
Journal of Neuroscience 12, 1679-1686.

De Proost I, Brouns I, Pintelon I, Timmermans JP & Adriaensen D. (2007). Pulmonary
expression of voltage-gated calcium channels: special reference to sensory airway
receptors. Histochem Cell Biol 128, 301-316.

Destexhe A, Neubig M, Ulrich D & Huguenard J. (1998). Dendritic low-threshold
calcium currents in thalamic relay cells. J Neurosci 18, 3574-3588.

130ng A, Yesilyurt 0, Isimer A & Giizeldemir ME. (2001). L-type and T-type calcium
channel blockade potentiate the analgesic effects of morphine and selective u
opioid agonist, but not to selective 5 and K agonist at the level of the spinal cord in
mice. Pain 93, 61-68.

Dubreuil A-S, Boukhaddaoui H, Desmadryl G, Martinez-Salgado C, Moshourab R,
Lewin GR, Carroll P, Valmier J & Scamps F. (2004). Role of T-type calcium

115

current in identified D-hair mechanoreceptor neurons studied in vitro. J Neurosci
24, 8480-8484.

Ertel EA, Campbell KP, Harpold MM, Hofmann F, Mori Y, Perez-Reyes E, Schwartz A,
Snutch TP, Tanabe T, Bimbaumer L, Tsien RW & Catterall WA. (2000).
Nomenclature of voltage-gated calcium channels. Neuron 25, 533-535.

F edulova SA, Kostyuk PG & Veselovsky NS. (1985). Two types of calcium channels in
the somatic membrane of new-bom rat dorsal root ganglion neurones. J Physiol
359, 431-446.

Feltz A. (2006). Low-threshold-activated Ca channels: from molecules to functions: over
25 years of progress. Crit Rev Neurobiol 18, 169-178.

Ferragamo MJ & Oertel D. (2002). Octopus cells of the mammalian ventral cochlear
nucleus sense the rate of depolarization. J Neurophysiol 87, 2262-2270.

F ishman MC & Spector I. (1981). Potassium current suppression by quinidine reveals
additional calcium currents in neuroblastoma cells. Proc Natl Acad Sci U S A 78,
5245-5249.

Gillessen T & Alzheimer C. (1997). Amplification of EPSPs by low Ni2+- and amiloride-
sensitive Ca channels in apical dendrites of rat CA1 pyramidal neurons. J
Neurophysiol 77, 1639-1643.

Gordon G & Manson JR. (1967). Cutaneous receptive fields of single nerve cells in the
thalamus of the cat. Nature 215, 597-599.

Gorelova NA & Yang CR. (2000). Dopamine Dl/DS receptor activation modulates a
persistent sodium current in rat prefrontal cortical neurons in vitro. J
Neurophysiol 84, 75-87.

Graham BA, Brichta AM & Callister RJ. (2004). An in vivo mouse spinal cord
preparation for patch-clamp analysis of nociceptive processing. J Neurosci
Methods 136, 221-228.

Grudt TJ & Perl ER. (2002). Correlations between neuronal morphology and

electmphysiological features in the rodent superficial dorsal horn. J Physiol 540,
189-207.

116

Gutierrez C, Cox CL, Rinzel J & Sherman SM. (2001). Dynamics of low-threshold spike
activation in relay neurons of the cat lateral geniculate nucleus. J Neurosci 21,
1022-1032.

Hagiwara S, Ozawa S & Sand O. (1975). Voltage clamp analysis of two inward current
mechanisms in the egg cell membrane of a starfish. J Gen Physiol 65, 617-644.

Han SH, Murchison D & Griffith WH. (2005). Low voltage-activated calcium and fast
tetrodotoxin-resistant sodium currents define subtypes of cholinergic and
noncholinergic neurons in rat basal forebrain. Brain Res Mol Brain Res 134, 226-
238.

Heinricher MM, Tavares I, Leith JL & Lumb BM. (2009). Descending control of
nociception: specificity, recruitment and plasticity. Brain Res Rev 60, 214-225.

Heppenstall P & Lewin G. (2006). A role for T-type Ca2+ channels in mechanosensation.
Cell Calcium 40, 165-174.

Hochman S, Garraway SM & Pockett S. (1997). Membrane properties of deep dorsal
horn neurons from neonatal rat spinal cord in vitro. Brain Res 767, 214-219.

Hongo T, Jankowska E & Lundberg A. (1968). post-synaptic excitation and inhibition
from primary afferents in neurones of the spinocervical tract. J Physiol 199, 569-
592.

Huang L-YM. (1989). Calcium channels in isolated rat dorsal horn neurones, including
labelled spinothalamic and trigeminothalamic cells. J Physiol 411, 161-177.

Huc S, Monteil A, Bidaud 1, Barbara G, Chemin J & Lory P. (2009). Regulation of T-
type calcium channels: signalling pathways and functional implications. Biochim
Biophys Acta 1793, 947-952.

Huguenard JR. (1996). Low-threshold calcium currents in central nervous system
neurons. Annu Rev Physiol 58, 329-348.

Huguenard JR & Prince DA. (1992). A novel T-type current underlies prolonged Ca2+-

dependent burst firing in GABAergic neurons of rat thalamic reticular nucleus. J
Neurosci 12, 3804-3817.

117

 

Ikeda H, Heinke B, Ruscheweyh R & Sandktihler J. (2003). Synaptic plasticity in spinal
lamina I projection neurons that mediate hyperalgesia. Science 299, 1237-1240.

Ikeda H, Stark J, Fischer H, Wagner M, Drdla R, Jager T & Sandktihler J. (2006).

' Synaptic amplifier of inflammatory pain in the spinal dorsal horn. Science 312,
1 659-1 662.

Inagaki A, Ugavia S, Yamamura H, Murakami S & Shimada S. (2008). The Cav3.l T-
type Ca channel contributes to voltage-dependent calcium currents in rat outer
hair cells. Brain Res 1201, 68-77.

Israel J-M, Poulain DA & Oliet SHR. (2008). Oxytocin-induced postinhibitory rebound
firing facilitates bursting activity in oxytocin neurons. J Neurosci 28, 385-394.

J evtovic-Todorovic V & Todorovic S. (2006). The role of peripheral T-type calcium
channels in pain transmission. Cell Calcium 40, 197-203.

J iang MC, Cleland CL & Gebhart GF. (1995). Intrinsic properties of deep dorsal horn
neurons in the L6-Sl spinal cord of the intact rat. J Neurophysiol 74, 1819-1827.

Johnson KO. (1974). Reconstruction of population response to a vibratory stimulus in
quickly adapting mechanoreceptive afferent fiber population innervating glabrous
skin of the monkey. J Neurophysiol 37, 48-72.

Joksovic PM, Bayliss DA & Todorovic SM. (2005). Different kinetic properties of two
T-type Ca currents of rat reticular thalamic neurones and their modulation by
enflurane. J Physio] 566, 125-142.

Karrer EE, Lincoln JE, Hogenhout S, Bennett AB, Bostock RM, Martineau B, Lucas WJ,
Gilchrist DG & Alexander D. (1995). In situ isolation of mRNA from individual
plant cells: creation of cell-specific cDNA libraries. Proc Natl Acad Sci U S A 92,
3814-3818.

2 . . .
Kawai F & Miyachi E. (2001). Enhancement by T-type Ca + currents of odor sensrtlv1ty
in olfactory receptor cells. J Neurosci 21, RC144 (141-145).

Kim DY, Choi H], Kim 13, Kim YS, Jeong DU, Shin HC, Kim M], Han H-C, Hong SK
& Kim YI. (2005). Voltage-gated calcium channels play crucial roles in the
glutamate-induced phase shifts of the rat suprachiasmatic circadian clock. Eur J

Neurosci 21, 1215-1222-

118

 

Klockner U, Lee JH, Cribbs LL, Daud A, ,Heschele2r+ J, Pereverzev A, Perez-Reyes E &
Schneider T. (1999). Comparison of the Ca currents induced by expression of

thrge cloned (1] subunits, (11 G, alH and all, of low-voltage- activated T-type
Ca channels. Eur J Neuroscz 11, 4171-4178.

Knibestol M. (1973). Stimulus-response functions of rapidly adapting mechanoreceptors
in human glabrous skin area. J Physiol 232, 427-452.

Kuo 11, Lee RH, Zhang L & Heckman CJ. (2006). Essential role of the persistent sodium
current in spike initiation during slowly rising inputs in mouse spinal neurones. J
Physiol 574, 819- 834.

Latour I, Hamid J, Beed12e+AM, Zamponi GW & Macvicar BA. (2003). Expression of
voltage-gated Ca channel subtypes in cultured astrocytes. Glia 41, 347-353.

Lee JH, Daud AN, Cribbs LL, Lacerda AE, Pereverzev A, Klockner U, Schneider T &
Perez-Reyes E. (1999a). Cloning and expression of a novel member of the low
voltage-activated T-type calcium channel family. J Neurosci 19, 1912-1921.

Lee JH, Gomora JC, Cribbs LL & Perez-Reyes E. (1999b). Nickel block of three cloned
T-type calcium channels: low concentrations selectively block alH. Biophys J 77,
3034-3042.

Lee JH, Kim EG, Park BG, Kim KH, Chi-1 SK, Kong ID, Lee JW & Jeong SW. (2002).
Identification of T-type alH Ca channels (Cav3.2) in major pelvic ganglion
neurons. J Neurophysiol 87, 2844-2850.

Lee S, Han TH, Sonner PM, Stern JE +Ryu PD & Lee SY. (2008). Molecular
characterization of T-type Ca +channels responsible for low threshold spikes in
hypothalamic paraventricular nucleus neurons. Neuroscience 155, 1195-1203.

Leresche N, Hering J & Lambert RC. (2004). Paradoxical potentiation of neuronal T-type
Ca2+ current by ATP at resting membrane potential. J Neurosci 24, 5592- 5602.

Lin DM, Loveall B, Ewer J, Deitcher DL & Sucher NJ. (2007). Characterization of
mRNA expression in single neurons. Methods M0] Biol 399, 133-152.

Liss B & Roeper J. (2004). Correlating function and gene expression of individual basal
ganglia neurons. Trends Neurosci 27, 475-481.

119

Llinas R & Yarom Y. (1981a). Electrophysiology of mammalian inferior olivary

neurones in vitro. Different types of voltage-dependent ionic conductances. J
Physio] 315, 549-567.

Llinas R & Yarom Y. (1981b). Properties and distribution of ionic conductances

generating electroresponsiveness of mammalian inferior olivary neurones in vitro.
J Physiol 315, 569-584.

Lopez-Garcia JA & King AE. (1994). Membrane properties of physiologically classified

rat dorsal horn neurons in vitro: correlation with cutaneous sensory afferent input.
Eur J Neurosci 6, 998-1007.

Lory P, Bidaud I & Chemin J. (2006). T-type calcium channels in differentiation and
proliferation. Cell Calcium 40, 135-146.

Lu Y & Perl ER. (2003). A specific inhibitory pathway between substantia gelatinosa
neurons receiving direct C-fiber input. J Neurosci 23, 8752-8758.

Lu Y & Perl ER. (2005). Modular organization of excitatory circuits between neurons of
the spinal superficial dorsal horn (laminae I and II). J Neurosci 25, 3900-3907.

Magee JC, Christofi G, Miyakawa H, Christie B, Lasser-Ross+N & Johnston D. (1995).
Subthreshold synaptic activation of voltage-gated Ca channels mediates a

localized Ca + influx into the dendrites of hippocampal pyramidal neurons. J
Neurophysiol 74, 1335-1342.

Magee JC & Johnston D. (1995). Synaptic activation of voltage-gated channels in the
dendrites of hippocampal pyramidal neurons. Science 268, 301-304.

Magistretti J & Alonso A. (1999). Biophysical properties and slow voltage-dependent
inactivation of a sustained sodium current in entorhinal cortex layer-II principal
neurons: a whole-cell and single-channel study. J Gen Physiol 114, 491-509.

Magistretti J, Castelli L, Forti L & D'Angelo E. (2006). Kinetic and functional analysis of
transient, persistent and resurgent sodium currents in rat cerebellar granule cells in

situ: an electrOphysiological and modelling study. J Physiol 573, 83-106.

Mark RF & Steiner J. (1958). Cortical projection of impulses in myelinated cutaneous
afferent nerve fibres of the cat. J Physiol 142, 544-562.

120

Martin RL, Lee JH, Cribbs LL, Perez-Reyes E & Hanck DA. (2000). Mibefradil block of
cloned T-type calcium channels. J Pharmacol Exp T her 295, 302-308.

Matthews EA & Dickenson AH. (2001). Effects of ethosuximide, a T—type Ca2+ channel

blocker, on dorsal horn neuronal responses in rats. Eur J Pharmacol 415, 141-
149.

McDonough SI & Bean BP. (1998). Mibefradil inhibition of T-type calcium channels in
cerebellar purkinje neurons. Mol Pharmacol 54, 1080-1087.

McKay BE, McRory JE, Molineux ML, Hamid J, Snutch TP, Zamponi. GW & Turner
RW. (2006). Cav3 T-type calcium channel isoforms differentially distribute to

somatic and dendritic compartments in rat central neurons. Eur J Neurosci 24,
2581-2594.

McRory JE, Santi CM, Hamming KS, Mezeyova J, Sutton KG, Baillie DL, Stea A &
Snutch TP. (2001). Molecular and functional characterization of a family of rat
brain T-type calcium channels. J Biol Chem 276, 3999-4011.

Melnick IV, Santos SF & Safronov BV. (2004a). Mechanism of spike frequency
adaptation in substantia gelatinosa neurones of rat. J Physiol 559, 383-395.

Melnick IV, Santos SF, Szokol K, Szucs P & Safronov BV. (2004b). Ionic basis of tonic
firing in spinal substantia gelatinosa neurons of rat. J Neurophysiol 91, 646-655.

Mittmann T & Alzheimer C. (1998). Muscarinic inhibition of persistent Na+ current in
rat neocortical pyramidal neurons. J Neurophysiol 79, 1579-1582.

Molander C, Xu Q & Grant G. (1984). The cytoarchitectonic organization of the spinal
cord in the rat. I. The lower thoracic and lumbosacral cord. J Comp Neurol 230,
133-141.

Molineux ML, McRory JE, McKay BE, Hamid J, Mehaffey WH, Rehak R, Snutch TP,
Zamponi GW & Turner RW. (2006). Specific T-type calcium channel isoforms
are associated with distinct burst phenotypes in deep cerebellar nuclear neurons.
Proc NatlAcad Sci USA 103, 5555-5560.

Moolenaar WH & Spector I. (1978). Ionic currents in cultured mouse neuroblastoma cells
under voltage-clamp conditions. J Physio] 278, 265-286.

121

Morisset V & Nagy F. (1998). Nociceptive integration in the rat spinal cord: role of non-
linear membrane properties of deep dorsal horn neurons. Eur J Neurosci 10,
3642-3652.

Mouginot D, Bossu JL & Gahwiler BH. (1997). Low-threshold Ca2+ currents in dendritic
recordings from Purkinje cells in rat cerebellar slice cultures. J Neurosci 17, 160-
170.

Murase K & Randic M. (1983). Electrophysiological properties of rat spinal dorsal horn
neurones in vitro: calcium-dependent action potentials. J Physiol 334, 141-153.

Niespodziany I, Derambure P & Poulain P. (1999). Properties of T-type calcium current

in enkephalinergic neurones in guinea-pig hypothalamic slices. Pflugers Arch
437, 871-880.

Nilius B, Hess P, Lansman JB & Tsien RW. (1985). A novel type of cardiac calcium
channel in ventricular cells. Nature 316, 443-446.

Nilius B, Talavera K & Verkhratsky A. (2006). T-type calcium channels: the never
ending story. Cell Calcium 40, 81-88.

Nowycky MC, Fox AP & Tsien RW. (1985). Three types of neuronal calcium channel
with different calcium agonist sensitivity. Nature 316, 440-443.

Parri HR & Crunelli V. (1998). Sodium current in rat and cat thalamocortical neurons:
role of a non-inactivating component in tonic and burst firing. J Neurosci 18, 854-
867.

Perez—Reyes E. (2003). Molecular physiology of low-voltage-activated T-type calcium
channels. Physiol Rev 83, 1 17-161.

Perez-Reyes E. (2006). Molecular characterization of T-type calcium channels. Cell
Calcium 40, 89-96.

Perez-Reyes E, Cribbs LL, Daud A, Lacerda AE, Barclay J, Williamson MP, Fox M,

Rees M & Lee JH. (1998). Molecular characterization of a neuronal low-voltage-
activated T-type calcium channel. Nature 391, 896-900.

122

 

Perez-Reyes E & Lory P. (2006). Molecular biology of T-type calcium channels. CNS
Neural Disord Drug Targets 5, 605-609.

Phillips JK & Lipski J. (2000). Single-cell RT-PCR as a tool to study gene expression in
central and peripheral autonomic neurones. Auton Neurosci 86, 1-12.

Prescott SA & De Koninck Y. (2002). Four cell types with distinctive membrane

properties and morphologies in lamina I of the spinal dorsal horn of the adult rat.
J Physiol 539, 817-836.

Prescott SA & De Koninck Y. (2005). Integration time in a subset of spinal lamina I
neurons is lengthened by sodium and calcium currents acting synergistically to
prolong subthreshold depolarization. J Neurosci 25, 4743-4754.

Prescott SA & Sejnowski TJ. (2008). Spike-rate coding and spike-time coding are
affected oppositely by different adaptation mechanisms. J Neurosci 28, 13649-
1 3 661 .

Rexed B. (1952). The cytoarchitectonic organization of the spinal cord in the cat. J Comp
Neurol 96, 414-495.

Reyes A. (2001). Influence of dendritic conductances on the input-output properties of
neurons. Annu Rev Neurosci 24, 653-675.

Rivera-Arconada I & Lopez-Garcia JA. (2005). Effects of M-current modulators on the

excitability of immature rat spinal sensory and motor neurones. Eur J Neurosci
22, 3091-3098.

Rozen S & Skaletsky HJ. (2000). Primer3 on the WWW for general users and for
biologist programmers. In Bioinformatics Methods and Protocols: Methods in
Molecular Biology, ed. Krawetz S & Misener S, pp. 365-386. Humana Press,
Totowa, NJ.

Ruscheweyh R, Ikeda H, Heinke B & Sandkl'ihler J. (2004). Distinctive membrane and
discharge properties of rat spinal lamina 1 projection neurones in vitro. J Physiol

555, 527-543.

Ruscheweyh R & Sandkilhler J. (2002). Lamina-specific membrane and discharge
properties of rat spinal dorsal horn neurones in vitro. J Physiol 541, 231-244.

123

Ruscheweyh R & Sandkfihler J. (2003). Epileptiforrn activity in rat spinal dorsal horn in
vitro has common features with neuropathic pain. Pain 105, 327-338.

Russo RE & Hounsgaard J. (19963). Burst-generating neurones in the dorsal horn in an in
vitro preparation of the turtle spinal cord. J Physiol 493 (Pt 1), 55-66.

Russo RE & Hounsgaard J. (1996b). Plateau-generating neurones in the dorsal horn in an
in vitro preparation of the turtle spinal cord. J Physiol 493 (Pt 1), 39-54.

Russo RE & Hounsgaard J. (1999). Dynamics of intrinsic electrophysiological properties
in spinal cord neurones. Prog Biophys Mo] Bio] 72, 329-3 65.

Rustioni A & Kaufman AB. (1977). Identification of cells or origin of non-primary
afferents to the dorsal column nuclei of the cat. Exp Brain Res 27, 1-14.

Ryu PD & Randic M. (1990). Low- and high-voltage-activated calcium currents in rat
spinal dorsal horn neurons. J Neurophysiol 63, 273-285.

Safronov BV. (1999). Spatial distribution of Na+ and K+ channels in spinal dorsal horn

neurones: role of the soma, axon and dendrites in spike generation. Prog
Neurobiol 59, 217-241.

Safronov BV, Wolff M & Vogel W. (1997). Functional distribution of three types of Na+

channel on soma and processes of dorsal horn neurones of rat spinal cord. J
Physiol 503, 371 -385.

Sahara Y, Xie YK & Bennett GJ. (1990). Intracellular records of the effects of primary

afferent input in lumbar Spinoreticular tract neurons in the cat. J Neurophysiol 64,
1791 -1 800.

Sakata Y, Saegusa H, Zong S, Osanai M,2Murakoshi T, Shimizu Y, Noda T, Aso T &
Tanabe T. (2001). Analysis of Ca currents in sperrnatocytes from mice lacking
Cav2.3 (1113) Ca channel. Biochem Bi0phys Res Commun 288, 1032-1036.

Sandktihler J. (2009). Models and mechanisms of hyperalgesia and allodynia. Physiol
Rev 89, 707-758.

Sandktthler J & Eblen-Zajjur AA. (1994). Identification and characterization of rhythmic
nociceptive and non-nociceptive spinal dorsal horn neurons in the rat.
Neuroscience 61, 991 - l 006.

124

Santos SFA, Rebelo S, Derkach VA & Safronov BV. (2007). Excitatory intemeurons

dominate sensory processing in the spinal substantia gelatinosa of rat. J Physiol
581, 241-254.

Schneider S. (1992). Functional properties and axon terminations of intemeurons in
laminae III-V of the mammalian spinal dorsal horn in vitro. J Neurophysiol 68,
1 746-1759.

Schneider S, Sandiford D, Kavookjian A & Johnson B. (1995). Synaptic connectivity of

local circuit neurons in laminae III and IV of hamster spinal cord. J Comp Neurol
355, 380-391.

Schneider SP. (2003). Spike frequency adaptation and signaling properties of identified
neurons in rodent deep spinal dorsal horn. J Neurophysiol 90, 245-258.

Schneider SP. (2005). Mechanosensory afferent input and neuronal firing properties in
rodent spinal laminae III-V: re-exarnination of relationships with analysis of
responses to static and time-varying stimuli. Brain Res 1034, 71-89.

Schneider SP. (2008). Local circuit connections between hamster laminae III and IV
dorsal horn neurons. J Neurophysiol 99, 1306-1318.

Shao Y, Alicknavitch M & Farach-Carson MC. (2005). Expression of voltage sensitive
calcium channel (VSCC) L-type Cavl .2 (am) and T-type Cav3.2 (1113) subunits
during mouse bone development. Dev Dyn 234, 54-62.

Shin H-S, Cheong E-J, Choi S, Lee J & Na HS. (2008). T-type Ca2+ channels as
therapeutic targets in the nervous system. Curr Opin Pharmacol 8, 33-41.

Shin J-B, Martinez-Salgado C, Heppenstall PA & Lewin GR. (2003). A T-type calcium
channel required for normal function of a mammalian mechanoreceptor. Nat
Neurosci 6, 724-730.

SoRelle R. (1998). Withdrawal of Posicor from market. Circulation 98, 831-832.

Sucher NJ & Deitcher DL. (1995). PCR and patch-clamp analysis of single neurons.
Neuron 14, 1095-1100.

125

Sun H & Wu SH. (2008). Physiological characteristics of postinhibitory rebound
depolarization in neurons of the rat's dorsal cortex of the inferior colliculus
studied in vitro. Brain Res 1226, 70-81.

Talavera K2+ & Nilius B. (2006). Biophysics and structure- function relationship of T- -type
Ca channels. Cell Calcium 40, 97-114.

Talley EM, Cribbs LL, Lee JH, Daud A, Perez-Reyes E & Bayliss DA. (1999).
Differential distribution of three members of a gene family encoding low voltage-
activated (T-type) calcium channels. J Neurosci 19, 1895-1911.

Tarasenko AN, Ko +uk PG, Eremin AV & Isaev DS. (1997). Two types of low-voltage-
activated Ca +'channels 1n neurones of rat laterodorsal thalamic nucleus. J
Physio] 499, 77-86.

Thomson AM, West DC & Headley PM. (1989). Membrane characteristics and synaptic

responsiveness of superficial dorsal horn neurons in a slice preparation of adult rat
spinal cord. Eur J Neurosci 1, 479-488.

Todorovic SM & Jevtovic-Todorovic V. (2006). The role of T-type calcium channels in

peripheral and central pain processing. CNS Neurol Disord Drug Targets 5, 639-
653.

Treviito CL, Felix R, Castellano LE, Gutiérrez C, Rodriguez D, Pacheco J, Lopez-
Gonzalez I Gomora JC, Tsutsumi V, Hemandez-Cruz A, F iordelisio T, Scaling
AL & Darszon A. (2004). Expression and differential cell distribution of low-
threshold Ca +'channels 1n mammalian male germ cells and sperm. FEBS Lett
563, 87-92.

Uebachs M, Schaub C, Perez-Reyes E & Beck H. (2006). T-type Ca2+ channels encode
prior neuronal activity as modulated recovery rates. J Physiol 571, 519-536.

Urban NN, Henze DA & Barrionuevo G. (1998). Amplification of perforant-path EPSPs
in CA3 pyramidal cells by LVA calcium and sodium channels. J Neur0physio] 80,
1558-1561.

van Welie I, Remme MW, van Hooft JA & Wadman WJ. (2006). Different levels of Ih

determine distinct temporal integration in bursting and regular-spiking neurons in
rat subiculum. J Physiol 576, 203-214.

126

Vémai P, Osipenko ON, Vizi ES & Spat A. (1995). Activation of calcium current in
voltage-clamped rat glomerulosa cells by potassium ions. J Physiol 483, 67-78.

Viana F, Van Den Bosch L, Missiaen L, Vandenberghe W, Droogmans G, Nilius B &
Robberecht W. (1997). Mibefradil (Ro 40-5967) blocks multiple types of voltage-
gated calcium channels in cultured rat spinal motoneurones. Cell Calcium 22,
299-311.

Willis WD & Coggeshall RE. (2004). Sensory mechanisms of the spinal cord. Kluwer
Academic/Plenum, New York.

Yasaka T, Kato G, Furue H, Rashid MH, Sonohata M, Tamae A, Murata Y, Masuko S &
Yoshimura M. (2007). Cell-type-specific excitatory and inhibitory circuits

involving primary afferents in the substantia gelatinosa of the rat spinal dorsal
horn in vitro. J Physiol 581, 603-618.

Ye J, Zhang J, Xiao C & Kong J. (2006). Patch-clamp studies in the CNS illustrate a
simple new method for obtaining viable neurons in rat brain slices: glycerol
replacement of NaCl protects CNS neurons. J Neurosci Methods 158, 251-259.

Yoshimura M & Jessell TM. (1989). Membrane properties of rat substantia gelatinosa
neurons in vitro. J Neurophysiol 62, 109-118.

Yunker AM. (2003). Modulation and pharmacology of low voltage-activated ("T-type")
calcium channels. J Bioenerg Biomembr 35, 577-598.

Yunker AM & McEnery MW. (2003). Low-voltage-activated ("T-type") calcium
channels in review. J Bioenerg Biomembr 35, 533-575.

Yunker AMR, Sharp AH, Sundarraj S, Ranganathan V, Copeland TD & McEnery MW.
(2003). Immunological characterization of T-type voltage-dependent calcium
channel Cav3.1 (alphal G) and Cav3.3 (alphalI) isoforms reveal differences in
their localization, expression, and neural development. Neuroscience 117, 321-
335.

Zhang D, Chen J, Saraf A, Cassar S, Han P, Rogers JC, Brioni JD, Sullivan JP &

Gopalakrishnan M. (2006). Association of CatSperl or -2 with Cav3.3 leads to
suppression of T-type calcium channel activity. J Biol Chem 281, 22332-22341.

127

 

Zhuravleva SO, Kostyuk PG & Shuba YM. 1+999). Divalent cation selectivity of the
subtypes of low voltage-activated Ca channels in thalamic neurons.
Neuroreport 10, 651-657.

Zhuravleva SO, Kostyuk PG & Shuba YM. (2001). Subtypes of low voltage-activated

Ca + channels in laterodorsal thalamic neurons: possible localization and
physiological roles. Pflugers Arch 441, 832-839.

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