ACID-SENSING ION CHANNELS
: REGULATION AND PHYSIOLOGIC FUNCTION
DISSERTATION
Presented in Partial Fulfillment of the Requirements for
the Degree Doctor of Philosophy in the Graduate
School of The Ohio State University
By
Jun-Hyeong Cho, M.D.
*****
The Ohio State University
2008
Dissertation Committee:
Professor Candice Askwith, Advisor
Professor Georgia Bishop Approved by Professor Jack Enyeart
Professor David Saffen ______Advisor Integrated Biomedical Sciences Graduate Program
ABSTRACT
Acid-sensing ion channels (ASICs) are proton (H+)-gated ion channels that
produce transient cation currents in response to extracellular acid. ASICs are expressed
widely in the brain, and contribute to learning, memory, and fear-related behaviors.
Inappropriate activation of ASICs during prolonged acidosis also induces neuronal
damage during brain ischemia. However, little is known about how ASICs contribute to
neuronal function or how ASICs are regulated by endogenous modulators.
For my thesis, I began by investigating the role of sulfhydryl compounds on ASIC
activity. I determined that glutathione, of which concentration increases in ischemic
brains, potentiated ASIC1a-mediated H+-gated currents by increasing apparent proton sensitivity and slowing channel desensitization. This potentiation of ASIC1a is due to the relief of tonic inhibition by transition metal ions, and the reduction of redox-sensitive residues. These results suggest that endogenous sulfhydryl compounds such as glutathione potentiate ASICs in neurons, and may exacerbate acidotoxic neuronal death during ischemia.
ASIC1 knockout mice display defects in learning and memory, suggesting a role of ASICs in synaptic function. I investigated how ASICs contribute to synaptic transmission by comparing synaptic responses between wild-type and ASIC knockout
ii hippocampal neurons in microisland culture. I determined that neurons from ASIC1
knockout mice have an increased probability of neurotransmitter release and altered
short-term plasticity. Further, transfection of ASIC1a into ASIC1 knockout neurons
restored release probability. These results suggest that ASIC1a regulates basal synaptic
transmission and short-term plasticity by modulating neurotransmitter release at
glutamatergic synapses, and that ASIC1a contributes to normal learning and memory
through its presynaptic function.
I also investigated the physiologic role of ASICs in the cerebellum. Both ASIC1 and ASIC2 are expressed in the cerebellum including Purkinje cells, which have intrinsic firing property required for normal cerebellar function. Neither ASIC1 knockout nor
ASIC2 knockout mice showed any difference in tonic firing of Purkinje cell in acute cerebellar slices compared to wild-type mice. However, the tonic firing rate of Purkinje cells was increased significantly in ASIC1/ASIC2 double knockout mice. These results suggest that both ASIC1 and ASIC2 can contribute to normal cerebellar function by regulating Purkinje cell firing independently.
Together, these studies describe, for the first time, how ASICs are regulated by
glutathione, one of the ischemia-related signals, and how ASICs regulate the
glutamatergic synaptic transmission of hippocampal neurons and the tonic firing of
cerebellar Purkinje cells. These results discussed herein provide significant insights on
the pathologic mechanism of ischemic neuronal death, and the physiologic roles of
ASICs in synaptic function and neuronal excitability.
iii
Dedicated to my parents
iv
ACKNOWLEDGMENTS
I would like to thank my advisor, Dr. Askwith for her support and encouragement. I also greatly appreciate my dissertation committee members, Drs. Bishop, Enyeart, and Saffen for their invaluable advice.
v
VITA
July 9th, 1974………………………Born in Seoul, South Korea
1993 – 1999……….....………….… M.D. College of Medicine The Catholic University of Korea, Seoul, South Korea
2003 – Present …………….….……Graduate Research Associate Department of Neuroscience The Ohio State University, Columbus, Ohio
PUBLICATIONS
J.-H. Cho and C.C. Askwith: Potentiation of acid-sensing ion channels by sulfhydryl compounds. (2007) Am J Physiol Cell Physiol, 292, C2161-74
J.-H. Cho and C.C. Askwith: Presynaptic release probability is increased in hippocampal neurons from ASIC1 knockout mice. (2008) J Neurophysiol, 99, 426-441
FIELDS OF STUDY
Major Field: Integrated Biomedical Science, Biology of Neurological Disorders
vi
TABLE OF CONTENTS
Page Abstract ...... ii Dedication ...... iv Acknowledgments ...... v Vita ...... vi List of Tables ...... ix List of Figures ...... x
Chapter 1 General introduction………………………..………..……………..……..1 1.1. Genes, protein structure, and the expression of ASICs 1.2. Biophysical properties of ASICs 1.3 Modulation of ASICs 1.4. Pathophysiologic roles of ASICs in brain ischemia, and modulation of ASICs 1.5. Physiologic role of ASICs in synaptic function, learning, and memory 1.6 Physiologic role of ASICs in cerebellar function
Chapter 2 Potentiation of acid-sensing ion channels by sulfhydryl compounds….20 2.1. Introduction 2.2. Materials and Methods 2.3. Results 2.4. Discussion
vii
Chapter 3 Presynaptic release probability is increased in hippocampal neurons from ASIC1 knockout mice………………...………..………………….64 3.1. Introduction 3.2. Materials and Methods 3.3. Results 3.4. Discussion
Chapter 4 Tonic firing of cerebellar Purkinje cell is altered in acid-sensing ion channel 1 and 2 double knockout mice ……...………………………..118 4.1. Introduction 4.2. Materials and Methods 4.3. Results 4.4. Discussion
Chapter 5 General discussion and future directions………….………….....…….139
Bibliography……………………………...……………….……….…………………..155
viii
LIST OF TABLES
Chapter 1
Table 1.1 Features of ASICs…………………………………………………....……….14
Chapter 4
Table 4.1 Firing rates of cerebellar Purkinje cells….………...…….…………….…....130
Table 4.2 Coefficient of variation of interspike intervals of Purkinje cell firing…...... 131
ix
LIST OF FIGURES
Chapter 1
Figure 1.1 Structure of ASIC subunit……………………………………………....…..15
Figure 1.2 Activation of ASIC by extracellular protons…………..…………. ………..16
Figure 1.3 Homo- and Hetero-multimerization of ASIC1a and ASIC2a…. …….……..17
Figure 1.4 Pathophysiologic mechanism of ASIC-dependent neuronal damage
in ischemic brains…………………………………..………………...……..18
Figure 1.5 A previous prevailing hypothesis on ASIC function in synaptic transmission
………………………………………………………………………………19
Chapter 2
Figure 2.1 DTT potentiates H+-gated currents in CHO cells expressing human ASIC1a
……………………………………………………………………………….47
Figure 2.2 The long-lasting effect of DTT on hASIC1a is reversed by DTNB…..……..49
Figure 2.3 Glutathione potentiates hASIC1a currents in a concentration-dependent
manner……………………………………………………………………….51
Figure 2.4 DTT slows the desensitization and shifts the pH dose-response of hASIC1a
……………………………………………………………………………….53
Figure 2.5 DTT and glutathione potentiate H+-gated currents in hippocampal neurons
…………………………………………………………………………….....55
x Figure 2.6 DTT-induced potentiation of H+-gated currents of CHO cells expressing
mouseASIC1a-containing channels..…….………………………………….57
Figure 2.7 DTT potentiates mouse ASIC2a, ASIC1b, and ASIC3 H+-gated
currents………………………………………………………………...... ….58
Figure 2.8 The metal chelator TPEN mimics the transient effect of DTT……….……..59
Figure 2.9 Lysine 133 mutation reduces both TPEN and DTT-induced potentiation of
hASIC1a…………………………..………………………………………...61
Figure 2.10 Effect of TPEN and DTT on mouse ASIC1a, ASIC1b, and ASIC3…....….62
Chapter 3
Figure 3.1 H+-gated currents of autaptic neurons in microisland cultures……………...94
Figure 3.2 Whole-cell action potential-evoked postsynaptic currents in GABAergic and
glutamatergic autaptic neurons……..………………………………..……...96
Figure 3.3 ASIC currents do not contribute to the peak amplitude of EPSC in
glutamatergic autaptic neurons………………..………………………….....98
Figure 3.4 The effects of acute ASIC activation by exogenously applied acid on
postsynaptic currents…………………………………….…………………101
Figure 3.5 Paired-pulse ratio and AMPAR EPSCs during short trains of repetitive stimuli
in autaptic neurons…..………………………………………………..……103
Figure 3.6 The effects of HEPES concentration and amiloride on AP-evoked
postsynaptic currents and the paired-pulse ratio.…………....…….……….105
Figure 3.7 Quantal analyses of AMPAR-mediated miniature EPSC (mEPSC) of
hippocampal neurons in mass culture.……………...… ………..…………108
xi Figure 3.8 Readily-releasable pool (RRP) size and RRP refilling of wild-type and ASIC1
knockout neurons.…….…………...……………………………………….110
Figure 3.9 The progressive block of NMDAR by MK-801 in glutamatergic autaptic
neurons.………...…………………………………………………….…….112
Figure 3.10 Release probability measured in glutamatergic autaptic neurons on
microislands..………………………...………………………...…………114
Figure 3.11 Rescue of ASIC1 knockout neurons by ASIC1a expression....….……….116
Chapter 4
Figure 4.1 Firing rates of cerebellar Purkinje cells from wild-type and ASIC knockout
mice………….………………………………………………..…………....132
Figure 4.2 Spontaneous firing rates of cerebellar Purkinje cells under the inhibition of
fast synaptic transmission…………………...…………………..………....134
Figure 4.3 Regularity of tonic firing of Purkinje cells from wild-type and ASIC knockout
mice……………….…………………………..…………………………....136
Figure 4.4 The effects of acute activation and inhibition of ASICs on tonic firing of
wild-type Purkinje cells…...... …....138
Chapter 5
Figure 5.1 Potentiation of ASICs by sulfhydryl compounds through two different
mechanisms………..…………………………………………………..…...152
Figure 5.2 Regulation of presynaptic release probability by ASIC1a: working model..153
Figure 5.3 Potential mechanisms of altered tonic firing in ASIC1/2 DKO Purkinje
cells…………………………………………………………..…….………154
xii
CHAPTER 1
GENERAL INTRODUCTION
Acid-sensing ion channels (ASICs) are proton (H+)-gated ion channels activated
by extracellular acid. ASICs are expressed in neurons of both the central and peripheral
nervous systems, and contribute to the physiologic function of neurons and to neuronal
damage in pathologic conditions. H+-gated currents induced by rapid decrease in
extracellular pH were first described from sensory neurons of dorsal root ganglia in 1980
(Krishtal and Pidoplichko 1981a, b, 1980, 1981c). The first ASIC gene (ASIC2a) was cloned in 1996 (Price et al. 1996; Waldmann et al. 1996), however the ligand for this putative ion channel was not known until 1997, when Waldman et al. cloned ASIC1a gene from rat brain cDNA, and found that ASIC1a was activated by extracellular protons
(Waldmann et al. 1997b).
1 1.1. Genes, protein structure, and the expression of ASICs
1.1.1. ASIC genes
ASICs belong to the degenerin/epithelial sodium channel (DEG/ENaC)
superfamily (Bianchi and Driscoll 2002; Kellenberger and Schild 2002). In mammals,
there are four ASIC genes, which encode six ASIC subunits: ASIC1a, ASIC1b, ASIC2a,
ASIC2b, ASIC3, and ASIC4. ASIC1a/ASIC1b and ASIC 2a/ASIC2b are splice variants
(Krishtal 2003). The cDNAs of ASIC genes have been cloned in many species including
human, rats, mice, chicken, toadfish, and zebra fish (Bassler et al. 2001; Chen et al. 1998;
Coric et al. 2005; de Weille et al. 1998; Garcia-Anoveros et al. 1997; Grunder et al. 2000;
Kellenberger and Schild 2002; Lingueglia et al. 1997; Price et al. 1996; Waldmann et al.
1997a; Waldmann et al. 1997b; Waldmann et al. 1996). The fact that ASIC1 from early vertebrates (lamprey and shark) do not display proton sensitivity although they are highly homologous to mammalian ASIC1 (66-67%) suggests that the proton sensitivity of
ASIC1 arose early in the evolution of vertebrates (Coric et al. 2005).
1.1.2. ASIC structure
Each ASIC subunit has two transmembrane domains, an extracellular loop, and two intracellular domains (Krishtal 2003; Wemmie et al. 2006) (Fig. 1). The second transmembrane domain lines the pore of ASIC, and contributes to ion permeation (Jasti et al. 2007). The extracellular domain has 14 conserved cysteines forming 7 disulfide bonds (Jasti et al. 2007), and multiple sites for glycosylation (Saugstad et al. 2004) (Fig.
1). The extracellular domain binds to protons and modulators of ASICs. Two
2 intracellular domains contain target residues for phosphorylation by protein kinases (Gao
et al. 2005; Leonard et al. 2003), and are also involved in the interaction with other
proteins through a PDZ domain at C-terminus (Anzai et al. 2002; Baron et al. 2002; Chai
et al. 2007; Deval et al. 2004; Duggan et al. 2002; Hruska-Hageman et al. 2002) (Fig. 1).
The crystal structure of chicken ASIC1 in its desensitized state has recently been
reported (Jasti et al. 2007). Three ASIC subunits associate to form functional H+-gated ion channels (Jasti et al. 2007). ASIC1a, ASIC1b, ASIC2a, and ASIC3 form functional homotrimeric channels whereas ASIC2b and ASIC4 do not form functional channels by themselves. ASIC subunits also form heteromultimeric channels (Babinski et al. 2000;
Bassilana et al. 1997; Lingueglia et al. 1997). Subunit composition determines the biophysical properties of heteromeric ASIC channels (Hesselager et al. 2004). Thus, the heteromultimerization of ASIC subunits produces a great diversity of H+-gated channels
with different biophysical characteristics such as proton sensitivity, desensitization kinetics, and responses to ASIC modulators.
1.1.3. ASIC expression in the nervous system
ASIC1a, ASIC2a and ASIC2b are expressed in both the central and peripheral
nervous systems (Lingueglia et al. 1997; Price et al. 1996; Waldmann et al. 1997b)
whereas ASIC1b and ASIC3 are primarily expressed in peripheral sensory neurons (Chen
et al. 1998; Waldmann et al. 1997a) (Table 1). ASICs are expressed throughout the brain
including the cerebral cortex, hippocampus, amygdala, olfactory bulb, and the cerebellum. ASICs are localized predominantly in the soma and dendrites of central neurons (Wemmie et al. 2002; Zha et al. 2006). ASICs are also expressed in retinal
3 neurons (Ettaiche et al. 2006; Ettaiche et al. 2004), and dorsal horn neurons of the spinal
cord (Wu et al. 2004). In the peripheral nervous system, ASICs are expressed in sensory
ganglia including the dorsal root ganglia (DRG) (Alvarez de la Rosa et al. 2002; Benson
et al. 2002; Poirot et al. 2006), the spiral and vestibular ganglia (Hildebrand et al. 2004;
Mercado et al. 2006; Peng et al. 2004). The expression of ASICs in neurons throughout the nervous system suggests fundamental roles of ASICs in neuronal functions. Indeed,
ASICs have been implicated in peripheral sensation, synaptic plasticity, learning and memory (Table 1). However, it remains to be determined how ASICs contribute to these functions.
1.2. Biophysical properties of ASICs
1.2.1. Ion permeability
ASICs are cation-selective ion channels. The ion selectivity of ASICs is Na+ >>
K+ (Waldmann et al. 1997b). Due to the relative selectivity of Na+ to K+ permeation, the
reversal potential of ASICs is close to the equilibrium potential of Na+ (Waldmann et al.
1997b). Thus, the activation of ASICs by protons induces the influx of Na+, and membrane depolarization at resting membrane potential (Vukicevic and Kellenberger
2004). ASIC1a homomultimers are also permeable to Ca2+, and the permeability ratio
(PNa+ / PCa2+) ranges from 2.5 to 18.5 (Bassler et al. 2001; Waldmann et al. 1997b). The
Ca2+ permeability of ASIC1a homomultimers suggests that ASIC1a can activate intracellular signal transduction mechanisms involving Ca2+ in physiologic and
pathologic conditions.
4
1.2.2. Proton-gating mechanism
Protons are the only known ligand of ASICs. ASICs generate transient cation
currents in response to extracellular acid, and are desensitized rapidly with a time
constant (τD) of 0.5 to 2.5 seconds (Fig. 2). ASIC3 is activated by multiple protons, which displace pore-blocking Ca2+ from a high-affinity binding site (Immke and
McCleskey 2003). However, the gating mechanism of ASIC1a homomultimer is more
complex, and seems to involve proton-induced changes in allosteric conformation rather
than unblocking of the pore (Zhang et al. 2006). Although several extracellular residues have been proposed as proton-binding sites of ASIC1a (Jasti et al. 2007; Paukert et al.
2008), and ASIC2a (Baron et al. 2001; Smith et al. 2007b), the exact proton-binding residues still remain to be determined.
1.2.3. Proton sensitivity of ASICs
The pH sensitivity of ASICs is dependent upon subunit composition (Hesselager et al. 2004). The pH that gives half-maximal activation (pH0.5) is ~ 6.5, 5.9, and 6.7 for
ASIC1a, ASIC1b, and ASIC3 homomultimers respectively (Chen et al. 1998; Cho and
Askwith 2007; Immke and McCleskey 2003) (Table 1). Proton sensitivity of ASICs is
also changed by ASIC modulators such as extracellular Ca2+ (Babini et al. 2002; de
Weille and Bassilana 2001; Paukert et al. 2004).
Low pH0.5 of ASIC2a homomultimers (~ 4.1) suggests that they are unlikely to be
activated in physiologic conditions (Bassilana et al. 1997). However, ASIC2a modulates
the biophysical properties of ASIC1a by forming heteromultimeric channels in
5 hippocampal neurons (Askwith et al. 2004). Compared to ASIC1a homomultimers,
ASIC1a/ASIC2a heteromultimers are less proton-sensitive (pH0.5 ~ 4.8), and relatively
2+ impermeable to Ca (PNa+ / PCa2+ ~ 36) (Askwith et al. 2004; Bassilana et al. 1997;
Yermolaieva et al. 2004) (Fig. 3). Hippocampal neurons express both ASIC1a homomultimers and ASIC1a/ASIC2a heteromultimers. Thus, the level of ASIC2a expression affects H+-gated currents in neurons by regulating the relative ratio of
ASIC1a/ASIC2a heteromultimer to ASIC1a homomultimer.
1.2.4. Acute desensitization, and steady-state desensitization of ASICs
A rapid shift of the extracellular pH from neutral to acidic activates ASICs, which
generate H+-gated currents that desensitize acutely with a time constant of 0.5 to 2.5 seconds. Because most ASICs are desensitized nearly completely, there is little sustained current even in the presence of acid (Waldmann et al. 1997b). However, ASIC3- containing channels generate sustained currents when acid is applied continuously
(Waldmann et al. 1997a). Therefore, ASIC3 can induce a long-lasting response to
extracellular acidosis whereas ASIC1a and ASIC2a induce a transient response in
neurons. ASICs also undergo steady-state desensitization at mildly acidic pH. Mild
extracellular acidosis (pH > 7.0), which does not induce the activation of ASICs by itself,
prevents the activation of ASICs when pH decreases further (Chen et al. 2006a;
Sherwood and Askwith 2008). Therefore, unlike rapid acidic shifts of the extracellular
pH, slow and gradual decreases in pH do not activate ASICs.
6 1.3. Modulation of ASICs
1.3.1. Ca2+, Mg2+, and lactate
Ca2+ and Mg2+ , divalent ions abundant in the extracellular space, inhibit ASIC1
and ASIC3-mediated H+-gated currents through direct interaction with ASICs (Babini et al. 2002; de Weille and Bassilana 2001; Paukert et al. 2004). Extracellular concentration of Ca2+ and Mg2+ decreases locally by lactate, which accumulates in ischemic conditions due to anaerobic metabolism. Lactate removes inhibitory Ca2+ and Mg2+ from ASICs, and potentiates H+-gated currents (Immke and McCleskey 2001). Thus, the accumulation
of lactate in the extracellular space can exacerbate ASIC-mediated neuronal death in
brain ischemia, and cardiac pain in myocardial ischemia (Benson et al. 1999; Benson and
Sutherland 2001; Sutherland et al. 2001; Yagi et al. 2006).
1.3.2. Transition metal ions
Zn2+ potentiates ASIC2a homomultimers in micromolar concentrations (Baron et al. 2001) whereas ASIC1a-containing channels are inhibited by Zn2+ in a nanomolar range (Chu et al. 2004). Zn2+ is released from synaptic vesicles during synaptic
transmission, and may potentiate postsynaptic ASIC2a by acting on ‘low’-affinity
binding site of Zn2+. However, the physiologic significance of ‘high’-affinity Zn2+ inhibition of ASIC1a remains to be determined. Extracellular fluid normally contains
Zn2+ in concentrations sufficient to saturate the high-affinity binding site, and such ambient Zn2+ would inhibit ASIC1a tonically. Endogenous Zn2+ chelator such as
7 glutathione can remove Zn2+ from the high-affinity binding site of ASIC1a, and increase
H+-gated currents (Cho and Askwith 2007) (Chapter 2).
1.3.3. ASIC inhibitors
Amiloride, a clinically used diuretic drug, blocks ASIC1a homomultimers with
IC50 ~10 µM (Waldmann et al. 1997b). Amiloride is neither potent nor specific ASIC
blocker because 100 ~ 300 µM amiloride are required to inhibit H+-gated currents
sufficiently in neurons, and amiloride at these concentrations also inhibits other ion channels and enzymes. Tarantula psalmotoxin 1 (PcTX1) is a more specific antagonist of
ASICs, and prevents the activation of ASIC1a homomultimers (Chen et al. 2006a, 2005;
Escoubas et al. 2000). These ASIC inhibitors are used to study ASIC function, and may also be applied as potential therapeutics for neuroprotection in brain ischemia.
1.3.4. Other modulators
Tissue inflammation and ischemia accompany the release of proinflammatory mediators including NGF, serotonin, arachidonic acids, and nitric oxide, which potentiate
H+-gated currents (Cadiou et al. 2007; Mamet et al. 2002; Poirot et al. 2004; Smith et al.
2007a). Potentiation of ASICs by these modulators may exacerbate inflammation- induced pain. The expression, subcellular localization, and biophysical properties of
ASICs are also regulated by physical interaction with other intracellular proteins, and phosphorylation by protein kinases (Deval et al. 2004; Duggan et al. 2002; Hruska-
Hageman et al. 2002); (Gao et al. 2005).
8 1.4. Pathophysiologic roles of ASICs in brain ischemia, and modulation of ASICs
During brain ischemia, extracellular pH decreases to a level sufficient to activate
ASICs (Pignataro et al. 2007). Activation of Ca2+-permeable ASIC1a homomultimers by extracellular acidosis induces the pathologic accumulation of intracellular Ca2+, which causes neuronal death in culture (Xiong et al. 2004; Yermolaieva et al. 2004) (Fig. 4).
Further, ASIC1 knockout mice displayed smaller infarct volume in stroke models, and a similar neuroprotective effect was observed when ASIC inhibitors (amiloride or PcTX1) were administered in wild-type mice (Xiong et al. 2007, 2006) (Fig. 4). These results indicate that Ca2+-permeable ASIC1a contributes to neuronal damage following brain ischemia, and drugs targeting ASIC1a or preventing extracellular acidosis may have therapeutic potential for stroke patients. In rodent model of stroke, ASIC blockade by amiloride or PcTX1 reduced stroke volume more than MK-801, a NMDA receptor blocker (Xiong et al. 2004). Attenuating brain acidosis by NaHCO3 was also similarly neuroprotective (Pignataro et al. 2007). Further, the therapeutic window of ASIC blockade is longer up to 4 hours after onset of stroke compared to NMDA receptor blockade (Pignataro et al. 2007). Considering the failure of NMDA receptor blockade by
MK-801 to display clinical benefits in acute stoke patients, ASIC is a promising therapeutic target for neuroprotection in stroke.
Oxygen/glucose-deprivation in ischemic brains does not only induce H+-gated activation of ASICs, but it also potentiates H+-gated currents, and slows the desensitization of ASICs (Xiong et al. 2004) (Fig. 4). This suggests that there could be endogenous ASIC modulators, which may accumulate in ischemic condition and
9 potentiate ASIC activity. Indeed, several mechanisms related to ischemia have been
reported to potentiate ASIC activity. For example, anaerobic metabolism during
ischemia increases the extracellular concentration of lactate, which potentiates H+-gated
currents by removing inhibitory Ca2+ from ASICs (Immke and McCleskey 2001) (Fig. 4).
Arachidonic acid and nitric oxide, which accumulate in ischemic brains, also potentiate
H+-gated currents in neurons through direct interaction with ASICs (Allen and Attwell
2002; Cadiou et al. 2007; Smith et al. 2007a). Thus, these ASIC modulators could exacerbate acid-induced neuronal damage mediated by ASICs. However, it has not been thoroughly examined how other ischemia-related signals affect ASIC activity as potential endogenous modulators of ASICs.
In Chapter 2 of my thesis, I focused my research on sulfhydryl compounds as potential modulators of ASICs. Endogenous sulfhydryl compounds such as glutathione and L-cysteines accumulate in extracellular space in ischemic conditions. Due to the chemical nature of their sulfhydryl group(s), these compounds bind to metal ions with high affinity, and reduce redox-sensitive residues including disulfide bonds of proteins.
Interestingly, ASICs are modulated by metal ion including Zn2+, and have multiple
disulfide bonds in the extracellular domain (Chu et al. 2004; Jasti et al. 2007). Therefore,
I hypothesized that endogenous sulfhydryl compounds modulate ASIC activity by
chelating metal ions from ASICs, and/or reducing disulfide bond(s).
ASICs have drawn much interest as a potential target for reducing neuronal
damage in stroke patients. However, the important roles of ASICs in neuronal function
(see below) suggest that complete blockade or inhibition of ASICs might cause
intolerable adverse effects when applied in stroke patients. Thus, it is important to inhibit
10 only pathologic activation of ASICs while preserving endogenous ASIC activity. The
development of this type of therapeutic strategy relies on detailed information of how
ASICs are inappropriately activated by endogenous modulators related to ischemia.
Therefore, my research on how ASICs are affected by sulfhydryl compounds could
provide insights on understanding the pathophysiologic mechanism of ASIC-mediated
neuronal damage and developing novel neuroprotective therapeutics.
1.5. Physiologic role of ASICs in synaptic function, learning, and memory
In the brain, ASIC1 contributes to learning and memory. ASIC1 knockout mice
display defects in multiple aspects of learning and memory including spatial learning,
fear conditioning, and eyeblink conditioning, which require normal synaptic plasticity in
the hippocampus, amygdala, and the cerebellum (Wemmie et al. 2002). This indicates
that ASIC1 contributes to synaptic plasticity (Wemmie et al. 2002). In consistence with
the defect in spatial learning, hippocampal long-term potentiation at CA1 to CA3
synapses is impaired in ASIC1 knockout mice (Wemmie et al. 2002). Hippocampal LTP
was normally induced in ASIC1 knockout mice when the activation of NMDA receptor
was facilitated by applying Mg2+-free solution (Wemmie et al. 2002). This suggests that the activation of NMDA receptors is reduced during high-frequency firing in ASIC1 knockout neurons. Indeed, short-term facilitation of NMDA receptor-mediated EPSP was reduced in ASIC1 knockout mice (Wemmie et al. 2002).
In 2002, Wemmie et al. proposed a popular hypothesis that the activation of postsynaptic ASIC1 by protons released from synaptic vesicles might depolarize the
11 postsynaptic membrane, and facilitate the activation of NMDA receptors by relieving
Mg2+ block, thus contribute to synaptic plasticity, learning and memory (Bianchi and
Driscoll 2002; Wemmie et al. 2002) (Fig. 5). However, there is controversy regarding this hypothesis. Although the synaptic cleft can be acidified during high-frequency stimulation (Krishtal et al. 1987), ASIC-dependent postsynaptic current evoked by synaptic activity has not yet been described. Further, the inhibition of ASICs by amiloride (a nonspecific blocker of ASICs) or mild acidic pH (pH 6.7) did not affect postsynaptic currents in cultured hippocampal neurons (Alvarez de la Rosa et al. 2003).
In chapter 3, I tried to reconcile this controversy on how ASICs affect synaptic
transmission and plasticity using microisland cultures of hippocampal neurons. I tested
the hypotheses that postsynaptic ASICs may be activated by protons released from
synaptic vesicles during synaptic transmission, and that synaptically-activated ASICs
contribute to basal synaptic transmission. I also investigated the presynaptic function of
ASICs by comparing synaptic responses between wild-type and ASIC knockout neurons.
These studies are the first to suggest that ASIC1a affects presynaptic function (Cho and
Askwith 2008), and lend fundamental insights into understanding the molecular
mechanism how ASICs contribute to normal learning and memory.
1.6. Physiologic role of ASICs in cerebellar function
ASIC1 and ASIC2 are highly expressed in the cerebellar cortex (Garcia-Anoveros
et al. 1997; Lingueglia et al. 1997; Waldmann et al. 1997b), including the dendrites and cell bodies of Purkinje cells (Duggan et al. 2002; Jovov et al. 2003; Wemmie et al. 2003).
12 Further, Purkinje cells and granule neurons from cerebellar cortex display H+-gated currents upon extracellular acid application (Allen and Attwell 2002; Bolshakov et al.
2002), indicating functional expression of ASICs in these neurons. These suggest that
ASICs may contribute to cerebellar function. Indeed, ASIC1 knockout mice display
impaired eyeblink conditioning (Wemmie et al. 2002), which requires normal cerebellar
function. However, little is known on how ASIC1 contributes to cerebellar function.
To bridge the gap between the loss of ASIC1 and the behavioral deficit observed
in ASIC1 knockout mice, I investigated the role of ASICs in cerebellar function, and
focused my study on Purkinje cell firing. Purkinje cells are the sole output neurons of
cerebellar cortex to deep cerebellar nuclei, and have intrinsic firing property (Raman et
al. 1997). Normal cerebellar function requires proper intrinsic firing of Purkinje cells
(Raman et al. 1997). Therefore, I hypothesized that ASICs contribute to cerebellar
function by regulating the intrinsic firing of Purkinje cells, and compared firing pattern of
Purkinje cells between wild-type and ASIC knockout mice in acute cerebellar slice
preparation (chapter 4). Considering the lack of research on the function of ASICs in the
cerebellum, the studies presented in chapter 4 would provide significant insights to
understand the mechanism how ASICs contribute to cerebellar function.
13 H+ sensitivity Name Expression Physiologic function pH0.5
ASIC1a 6.5 Multiple brain regi ons including Synaptic plasticity cerebral cortex, hippocampus, Learning and memory cerebellum, olfactory bulb, I n n a te and acquired fear and am ygdala Spinal cord (dorsal horn neuron) Inflammation -induced hyperalgesia D RG Unknown
ASIC1b 5.9 Cochlear hair cell Hearing? Tongue Sour taste? D RG Nociception?
14 ASIC2a 4.1 Multiple brain regions Unknown Retina Retinal function Spiral and vestibular ganglia Hearing? Tongue Sour taste? D RG (mechanosensory neuron) Mechanosensation?
ASIC2b not gated by H Multiple brain regions Unknown Retina Retinal function D RG Unknown
ASIC3 6.7 DRG (mechanosensory neuron) Mechanosensation DRG (nociceptor neuron) Nociception S piral and vestibular ganglia Hearing?
ASIC4 not gated by H Brain (pituitary gland) Unknown
Table 1.1: Features of ASICs Conserved Cysteine
Glycosylation
PKA/CaMKII phosphorylation site
out
TM1 TM2
in PDZ domain N C
Figure 1.1: Structure of ASIC subunit
15 A. Proton-dependent gating of ASICs
pH 7.4 / closed out
ASIC in
pH 6.0 / open Proton(H+)
Na+/ Ca2+
pH 6.0 / desensitized Proton(H+)
B. Proton-gated current in a hippocampal neuron pH 7.4 pH 6.0
Figure 1.2: Activation of ASICs by extracellular protons
16 A. ASIC1a homotrimer pH 6.0
ASIC1a .2 nA 0 subunit 5 s
More proton-sensitive Slower desensitization Slower recovery Ca2+-permeable
B. ASIC1a/ASIC2a heterotrimer pH 6.0
ASIC1a .2 nA subunit 0 5 s
ASIC2a Less proton-sensitivive subunit Faster desensitization Faster recovery Ca2+-impermeable
Fig. 1.3: Homo- and hetero-multimerization of ASIC1a and ASIC2a
17 Occlusion of cerebral artery
Brain ischemia
Oxygen/glucose deprivation
Local extracellular acidosis Lactate Arachidonic acid H+-dependent activation of ASICs + Nitric oxide (NO) Glutathione (GSH) ? Pathologic accumulation of Amiloride intracellular Ca2+ - PcTX1
Acidotoxic neuronal damage
Figure 1.4: Pathophysiologic mechanism of ASIC-dependent neuronal damage in ischemic brains
18 ~ pH 5.7 Ca2+
Glutamate Proton (H+)
ASIC AMPA NMDA receptor receptor
Na+ / Ca2+ infux
?
Figure 1.5: A previous prevailing hypothesis on ASIC function in synaptic transmission
19
CHAPTER 2
POTENTIATION OF ACID-SENSING ION CHANNELS BY SULFHYDRYL COMPOUNDS
The acid-sensing ion channels (ASICs) are voltage-independent ion channels activated by acidic extracellular pH. ASICs play a role in sensory transduction, behavior,
and acidotoxic neuronal death which occurs during stroke and ischemia. During these
conditions, the extracellular concentration of sulfhydryl reducing agents increases. We
used perforated patch-clamp technique to analyze the impact of sulfhydryls on H+-gated currents from CHO cells expressing human ASIC1a (hASIC1a). We found that hASIC1a currents activated by pH 6.5 were increased almost two-fold by the sulfhydryl-containing reducing agents, dithiothreitol (DTT) and glutathione. DTT shifted the pH dose-response of hASIC1a toward more neutral pH (pH0.5 from 6.54 to 6.69) and slowed channel desensitization. The effect of reducing agents on native mouse hippocampal neurons and transfected mouse ASIC1a was similar. We found that the effect of DTT on hASIC1a was mimicked by the metal chelator, TPEN, and mutant hASIC1a channels with reduced
TPEN potentiation showed reduced DTT potentiation. Furthermore, the addition of DTT
in the presence of TPEN did not result in further increases in current amplitude. These
20 results suggest that the effect of DTT on human ASIC1a is due to relief of tonic
inhibition by transition metal ions. We found that all ASICs examined remained potentiated following the removal of DTT. This effect was reversed by the oxidizing agent DTNB in hASIC1a supporting the hypothesis that DTT also impacts ASICs via a redox sensitive site. Thus, sulfhydryl compounds potentiate H+-gated currents via two
mechanisms, metal chelation and redox modulation of target amino acids.
2.1. INTRODUCTION
Acid-sensing ion channels (ASICs) are members of the DEG/ENaC ion channel family. These voltage-independent ion channels are activated by extracellular acidosis and are expressed in neurons throughout the central and peripheral nervous system
(Waldmann 2001; Waldmann et al. 1997b). There are four ASIC genes which encode at least six ASIC subunits (ASIC1a, 1b, 2a, 2b, 3, and 4). ASIC subunits share common structural features with all DEG/ENaC ion channels including an intracellular N- and C- terminus, two transmembrane domains, and a large cysteine-rich extracellular region
(Kellenberger and Schild 2002). Individual subunits associate to form homomultimeric or heteromultimeric channels with different characteristics (Askwith et al. 2004; Benson et al. 2002; Hesselager et al. 2004). ASICs are cation permeable and modulate action
potential generation in neurons upon extracellular acidification (Poirot et al. 2006;
Vukicevic and Kellenberger 2004). In the peripheral nervous system, disruption of
individual ASIC genes affects multiple aspects of sensory transduction including
mechanotransduction and nociception (Chen et al. 2002; Jones et al. 2005; Mogil et al.
21 2005; Price et al. 2000; Price et al. 2001). In the central nervous system, disruption of
ASIC1 impacts fear-related behaviors as well as learning and memory (Wemmie et al.
2003; Wemmie et al. 2002; Wemmie et al. 2004).
ASICs contribute to neuronal death following stroke and ischemia (Xiong et al.
2004). During these conditions, the extracellular environment in affected tissue becomes acidic (Siesjo 1988). This increase in proton concentration is thought to activate ASICs.
In the brain, many neurons express both ASIC1a homomultimeric and ASIC1a/2a heteromultimeric channels (Askwith et al. 2004; Chu et al. 2004). Homomultimeric
ASIC1a channels are calcium permeable and require less acidic pH to be activated than
ASIC1a/2a channels (Askwith et al. 2004; Bassilana et al. 1997; Chu et al. 2002;
Yermolaieva et al. 2004). Activation of ASIC1a channels causes an acute increase in intracellular calcium. Excess activation of ASIC1a channels during stroke and ischemia causes neuronal death (Xiong et al. 2004; Yermolaieva et al. 2004). Injection of venom containing PcTX1, a peptide known to prevent ASIC1a activation, reduces neuronal damage in mouse models of ischemia (Xiong et al. 2004). Additionally, mice with a genetic disruption of the ASIC1 gene show less damage following ischemia (Xiong et al.
2004). These results indicate that preventing ASIC activation limits damage and, therefore, agents which alter ASIC activity may impact acidosis-induced neuronal death following stroke. ASICs also impact migration of malignant glioma cells suggesting
ASIC activity may also play a role in the pathogenesis of brain tumors (Bubien et al.
2004; Vila-Carriles et al. 2006).
ASICs are modulated by several compounds in a subunit specific manner. For example, RFamide-related peptides slow or prevent desensitization of ASIC1 or ASIC3-
22 containing channels (Askwith et al. 2000; Catarsi et al. 2001; Chen et al. 2006b; Deval et
al. 2003; Ostrovskaya et al. 2004; Xie et al. 2003). Low concentrations of zinc limit
ASIC1a and ASIC1a/2a channel activation, while high concentrations potentiate ASIC2a
homomultimeric channels (Baron et al. 2001; Chu et al. 2004; Gao et al. 2004). Recent
work indicates that rodent ASIC1a activity is also affected by redox reagents (Andrey et
al. 2005; Chu et al. 2006). These publications reported different effects of the redox
reagents and suggested distinct mechanisms of sulfhydryl-induced modulation. In order
to gain insight into the mechanism of modulation of human and mouse ASICs, we
undertook a study of glutathione and DTT, an exogenous sulfhydryl reagent that mimics
the effects of endogenous sulfhydryls. Our results suggest that sulfhydryls potentiate
ASIC1a through two mechanisms: reducing amino acid residues within the ASIC protein
and chelating transition metals which tonically inhibit ASIC activity. Furthermore, we
found that sulfhydryl compounds induce potentiation of most ASIC channels. These
results lend significant insight into sulfhydryl modulation of ASICs and suggest that
sulfhydryl-induced potentiation may impact neuronal activities linked to ASIC function.
2.2. MATERIALS AND METHODS
2.2.1. DNA constructs and CHO cell transfection
Human and mouse ASIC cDNA constructs were cloned in pMT3 expression vector as described (Askwith et al. 2004). The K133R mutant of human ASIC1a was
made using the QuickChange site-directed mutagenesis kit (Stratagene, La Jolla, CA).
Chinese hamster ovary (CHO) cells were transiently transfected with ASIC constructs
23 using electroporation. Briefly, trypsinized CHO cells (~ 107 cells) were suspended in 0.4 mL of electroporation solution (120 mM KCl, 25 mM HEPES, 10 mM K2HPO4, 10 mM
KH2PO4, 2 mM MgCl2, 0.15 mM CaCl2, 5 mM EGTA, and 2 mM MgATP, pH 7.6), and mixed with 2 ~ 4 µg of pEGFP-C1 (Clontech, Mountain View, CA) as well as 10 ~ 20 µg of ASIC constructs. Cells were electroporated with the Gene Pulser Xcell system (Bio-
Rad Laboratories, Hercules, CA) and plated at a density of 35 cells/mm2 onto 10 mm covelslips in a 35 mm culture dish. Cells were used for patch-clamping 2 to 3 days after transfection. Transfected cells were identified by green fluorescent protein (GFP) fluorescence.
2.2.2. Primary neuronal culture
Primary hippocampal neuron cultures were prepared using previously published methods (Askwith et al. 2004; Wemmie et al. 2002). Briefly, hippocampi were dissected from postnatal day 0 ~ 2 pups, freed from extraneous tissue, and cut into approximately 8 pieces. The hippocampal tissue was transferred into Leibovitz’s L-15 medium
(Invitrogen, Carlsbad, CA) containing 0.25 mg/ml bovine serum albumin and 0.375 mg/ml papain and incubated for 15 min at 37°C with 95% O2 / 5% CO2 gently blown over the surface of the medium. After incubation, the hippocampal tissue were washed three times with mouse M5-5 medium (Earle’s minimal essential medium with 5% fetal bovine serum, 5% horse serum, 0.4 mM L-glutamine, 22 mM glucose, penicillin/streptomycin, and insulin/selenite/transferrin) and triturated. Hippocampal cells (5 x 104 cells per well) were plated onto collagen-coated 10 mm coverslips in 24
well culture dishes. After 72 hours, cytosine β-D-arabinofuranoside was added to inhibit
24 glial proliferation. After 10 days in vitro, half of the culture medium was replaced with
fresh M5-5 medium every three days. Neurons were used from 12 to 20 days in culture.
2.2.3. Electrophysiology
We used the nystatin-based perforated patch-clamp technique to record H+-gated currents. This method proved very stable and ASIC current rundown was significantly smaller in perforated patch recording compared to conventional whole cell patch- clamping. The extracellular solution contained 140 mM NaCl, 5.4 mM KCl, 10 mM
HEPES, 10 mM MES, 2 mM CaCl2, 1 mM MgCl2, and 5.55 mM glucose.
Tetramethylammonium hydroxide was used to adjust the pH of the extracellular solution
to pH 4.0 ~ pH 7.4. Intracellular pipette solution contained 130 mM K-gluconate, 20
mM KCl, 10 mM HEPES, and 0.1 mM EGTA (pH 7.3). The pipette tip was filled with
the intracellular solution, and then back-filled with the solution containing 150 µg/mL of
nystatin. Nystatin stock solution (30 mg/mL) was made fresh in DMSO before patch-
clamping. Patch electrodes were pulled with a P-97 micopipette puller (Sutter Instrument
Co., Novato, CA) and fire-polished with a microforge (Narishige, East Meadow, NY).
Micropipettes with 3 ~ 7 MΩ were used for experiments. Large pyramidal neurons were
chosen for patch-clamp analysis. After attaining a Giga-Ohm seal, we monitored whole-
cell membrane capacitance and series resistance until the patch was fully perforated by
nystatin. This usually occurred within 5 minutes. Cells were continuously superfused
with the extracellular solution from gravity-fed perfusion pipes at a flow rate of
approximately 1 mL/min. Perfusion pipes were placed 250 to 300 µm away from cells
and flow was directed toward the recorded cells to ensure fast solution exchange. For 25 hippocampal neurons, we added ion channel inhibitors to the extracellular solution to
inhibit synaptic currents (10 µM cyano-7-nitroquinoxaline-2,3-dione, 50 µM D-2-amino-
5-phosphonovaleric acid, 30 µM bicuculline, and 500 nM tetrodotoxin). The addition of
dithiothreitol (DTT), 5,5’-dithio-bis 2-nitrobenzoic acid (DTNB), or N,N,N′,N′-
tetrakis(2-pyridylmethyl)ethylenediamine (TPEN) did not alter the pH of the extracellular
solution. Unless otherwise indicated, all reagents were purchased from Sigma-Aldrich
(St. Louis, MO) or Fisher Scientific (Waltham, MA).
The membrane potential was held constant at -70 mV. Data were collected at 5
kHz using an Axopatch 200B amplifier, Digidata 1322A and Clampex 9 (Molecular
Devices, Sunnyvale, CA). In most experiments, H+-gated currents were evoked by the
exogenous application of pH 6.5 or pH 6.0 extracellular solutions at 2 ~ 2.5 minute
intervals. Because of current rundown, we evoked 4 to 5 H+-gated currents to ensure a stable pretreatment control value before the incubation of reagents. Data were analyzed using Clampfit 9 software (Molecular Devices). Maximal current amplitudes were normalized to the pretreatment control values and are expressed as the mean ± the standard error of the mean (SEM). For determining the pH dose-response curves, peak
current amplitudes at different test pHs were normalized to the average amplitude of pH
5.0 currents evoked just before and after the test pH application. The average of
normalized currents at different pHs was fitted to the equation
+ n n (pH – pH0.5) I / I pH 5.0 = 1 / {1 + (EC50 / [H ]) } = 1 / {1 + 10 },
26 where n is Hill coefficient, and EC50 and pH0.5 are the proton concentration and pH yielding half of the pH 5.0 currents (I pH 5.0). To ensure that ASIC1a/2a heteromultimers were generated by expression of both ASIC1a and ASIC2a in CHO cells, we measured the rate of recovery from desensitization. Briefly, pH 6.0 was applied for 3 ~ 5 seconds, the pH was returned to 7.4 for approximately 2.5 seconds, and then a second pH 6.0 application was made. We calculated the percent recovery by comparing the peak amplitude of the first pH 6.0 application to that of the second pH 6.0 application.
Consistent with the previous report (Askwith et al. 2004; Benson et al. 2002), the recovery rate was faster in cells expressing both mASIC1a and mASIC2a (74 ± 4 %, n =
13) than in cells expressing mASIC1a alone (41 ± 3 %, n = 13, p < 0.0001, unpaired t test). We used unpaired Student’s t test for the comparison of H+-gated currents from different groups of cells and the paired Student’s t test when comparing H+-gated currents with the pretreatment control in the same cell. We used one-way ANOVA to compare currents between more than two different groups. Statistical analyses were performed with Minitab14 software (Minitab Inc, State College, PA).
2.3. RESULTS
2.3.1. DTT potentiates homomeric hASIC1a currents in CHO cells.
To better understand the mechanism of sulfhydryl modulation of ASIC currents, we expressed human ASIC1a (hASIC1a) in CHO cells and used perforated patch-clamp to analyze whole-cell currents. We evoked H+-gated currents by changing the extracellular pH from 7.4 to 6.5 for 5 seconds. Once stable pH 6.5-evoked currents were
27 established, we exposed cells to 1 mM DTT at pH 7.4. One minute after the addition of
DTT, we activated hASIC1a by changing the pH from 7.4 to 6.5 in the presence of DTT
(Fig. 1A). We found that the peak amplitude of H+-gated currents was increased almost two fold in the presence of 1 mM DTT compared to currents recorded before DTT addition (182 ± 17 % of preDTT control, n = 18, p = 0.0009, Fig. 1B ~ C). ASIC desensitization was also dramatically slowed in the presence of DTT (Fig. 1A). Similar results were obtained with 10 µM DTT (peak amplitude 180 ± 20 % of preDTT control, n
= 10, p = 0.003, Fig. 1C) indicating that lower concentrations of DTT have similar effects. Both pH 6.5 and DTT had no effect on CHO cells expressing only GFP (Fig. 1D) indicating that the potentiation observed was dependent on hASIC1a expression. Thus,
DTT potentiates human ASIC1a currents.
The potentiation by DTT could be due to two mechanisms, chelation of transition metals and/or reduction of amino acid residues (Paoletti et al. 1997; Sullivan et al. 1994).
The first effect would be transient and persist only as long as DTT is present. The latter effect could endure after the removal of DTT due to the stable nature of covalent modification. In order to investigate the involvement of these two mechanisms in sulfhydryl modulation of hASIC1a, we assessed whether currents remained potentiated after the removal of DTT. Cells were incubated with DTT solution at pH 7.4 for three minutes and then superfused with DTT-free pH 7.4 solution for one minute. Following this wash step, the pH was changed to pH 6.5 in the absence of DTT to activate ASIC1a currents (Fig. 1A). The amplitude of pH 6.5 currents after DTT incubation and removal was 134 ± 4 % of the preDTT control (n = 37, p < 0.0001, Fig. 1A ~ C). This potentiation was significantly smaller than the potentiation observed in the presence of
28 DTT (n = 13, p = 0.005). We continued to superfuse the cells with pH 7.4 solution and
activate ASIC1a currents with pH 6.5 every two minutes. We found that the increase in
peak current amplitude persisted for at least 5 minutes following DTT removal (Fig. 1A ~
B). In cells which afforded analysis at later times after DTT incubation, we found that
the effect was maintained for as long as 15 minutes (n = 2, data not shown). Together,
these results indicate that DTT has both transient (the increase in current in the presence
of DTT) and long-lasting (the potentiation after DTT removal) effects on hASIC1a.
We hypothesized that the long-lasting potentiation of hASIC1a currents after DTT
removal may be due to reduction of specific amino acids by DTT. To determine whether
oxidation of those residues could reverse the effect of DTT, we incubated the cells with
DTNB, a compound that covalently modifies proteins and forms mixed disulfides with
free sulfhydryl groups (Fig. 2). Following DTT incubation and removal, hASIC1a
currents were activated by pH 6.5 solution. Then, DTNB was applied for 3 minutes and
washed away for an additional 3 minutes (Fig. 2A). After DTNB incubation and
removal, hASIC1a currents were again activated by changing the pH from 7.4 to 6.5. We
found that DTNB exposure reversed the long-lasting potentiation of hASIC1a currents
following DTT incubation and washout (130 ± 5 % after DTT/wash, and 108 ± 5 % after
DTNB/wash, n = 12, Fig. 2B). Without previous DTT exposure, the application and
removal of DTNB did not alter the amplitude of H+-gated currents (107 ± 5% of previous preDTNB control, n = 9, p = 0.24 Fig. 2C ~ D). Therefore, the effect of DTNB was dependent on previous exposure to DTT. These results indicate that DTT-induced reduction of residues within hASIC1a potentiates H+-gated currents and oxidation of these reduced residues by DTNB reversed this potentiation.
29 Together, our results indicate that DTT has two effects on hASIC1a. First, the
presence of DTT during ASIC activation induces large potentiation of pH 6.5-activated
hASIC1a currents. Once DTT is removed, potentiation is reduced, but current amplitude
remains increased compared to preDTT controls. This effect is reversed by DTNB
indicating that reduction of amino acid residues by DTT is required for this long-lasting effect.
2.3.2. Glutathione potentiates hASIC1a
DTT is an exogenous sulfhydryl compound. In order to test whether endogenous
sulfhydryl compounds could potentiate hASIC1a, we treated transfected CHO cells with
reduced glutathione. In the presence of 100 µM glutathione, the amplitude of H+-gated currents increased (165 ± 16 %, n = 9, p = 0.003, Fig. 3A ~ B). Similar to DTT treatment, the amplitude of H+-gated currents decreased partly after a five minute
washout, but still remained elevated compared with pre-glutathione control (129 ± 9 %, n
= 7, p = 0.02, Fig. 3A ~ B). We also tested the effects of glutathione at lower
concentrations (1 µM and 10 nM, Fig. 3B) and observed that potentiation in the presence of glutathione was concentration-dependent (Fig. 3C). Persistent potentiation that remained after the removal of glutathione was only observed with 100 µM and 1 µM
glutathione suggesting that the long-lasting potentiation requires higher concentrations of glutathione (micromolar range) whereas transient potentiation in the presence of glutathione requires much lower concentrations of glutathione (nanomolar range).
30 2.3.3. DTT affects hASIC1a pH dose-response and channel desensitization
Our results indicate that DTT impacts the peak current amplitude of pH 6.5-
activated hASIC1a currents and channel desensitization (Fig. 1A). Human ASIC1a
channels desensitize in the presence of acidic solution, and desensitization controls the duration of ASIC currents. The presence of DTT caused an obvious slowing of channel desensitization (Fig. 1A and 4A). Desensitization is usually quantified by fitting the desensitization phase of the ASIC current to a single exponential equation and calculating
the tau of desensitization (τd). This point represents the time required for the current to
reduce to 37 % of its maximal value (1/e). Since hASIC1a desensitization in the presence
of DTT did not fit to an exponential equation, we quantified desensitization by measuring
+ the time required for H -gated currents to decrease to 37 % of the peak amplitude (Td .37).
Using this method to quantify desensitization, we found that the Td.37 of hASIC1a currents at pH 6.5 was larger in the presence of DTT (Td .37 = 1.49 ± 0.21 seconds for the
preDTT control, 4.56 ± 0.86 seconds during DTT incubation, n = 6, p = 0.011, Fig. 4B).
After the removal of DTT, desensitization of H+-gated currents was still two fold larger
than preDTT control values (Td.37 = 2.97 ± 0.40 seconds after DTT washout, n = 6, p =
0.007, Fig. 4B). These results indicate that desensitization of hASIC1a was slowed in the presence of DTT and remained affected after washout of DTT.
Our data show that DTT potentiates hASIC1a currents activated by pH 6.5 solutions (Fig. 1). To determine whether DTT potentiation was dependent on the pH of the activating solutions, we analyzed DTT effects using different activating pHs. We found that currents activated by pH 5.0 did not change significantly during or after DTT incubation (105 ± 7 %, n = 6, p = 0.41 during DTT incubation, 106 ± 6%, n = 5, p = 0.91
31 after washout of DTT, Fig. 4C). These results suggested that DTT-induced potentiation
was pH-dependent. We performed a detailed pH dose-response analysis of hASIC1a
during DTT incubation, and after DTT removal. The pH which induced half maximal
peak current amplitude (pH0.5) of hASIC1a before DTT incubation was 6.54. During
DTT incubation, the pH dose-response curve shifted toward neutral physiologic pH with
+ pH0.5 = 6.69 (Fig. 4D). Following DTT incubation and washout, H -gated currents at pH
6.5 and pH 6.7 were still significantly elevated and the pH dose-response curve remained
shifted toward neutral pH with pH0.5 = 6.60 (Fig. 4D). These results indicate that DTT increases the apparent affinity of hASIC1a to protons, and that the effect of DTT is pH- dependent.
2.3.4. Potentiation of ASIC currents by sulfhydryl compounds in hippocampal neurons
Our data indicate that sulfhydryls potentiate hASIC1a currents. To examine whether neuronal H+-gated currents are similarly affected by sulfhydryl compounds, we
cultured hippocampal neurons from postnatal mice and tested the effects of DTT and
reduced glutathione. Homomeric ASIC1a and heteromeric ASIC1a/2a channels
contribute to H+-gated currents in central neurons (Askwith et al. 2004). Because these two types of ASIC channels have different pH sensitivities and the effect of DTT is pH- dependent, we used pH 6.0 to activate H+-gated currents from neurons (Fig. 5A). Similar to our observations in CHO cells transfected with hASIC1a, the amplitude of pH 6.0- activated currents increased in the presence of 1 mM DTT (158 ± 15%, n = 11, p = 0.004,
Fig. 5A). After removal of DTT, currents remained elevated compared to preDTT
control currents (126 ± 5 %, n = 11, p = 0.0007, Fig. 5A ~ B). The peak current
32 amplitude of H+-gated currents in hippocampal neurons consistently decrease with repeated applications of acidic solutions. Therefore, we compared DTT-treated neurons to mock-treated neurons, and observed that DTT-treated neurons showed larger currents for at least 6 minutes indicating that potentiation persisted even after the removal of DTT
+ (Fig. 5B). H -gated current desensitization was slowed in the presence of DTT (Td.37 =
1.45 ± 0.16 seconds for preDTT control, 1.94 ± 0.30 seconds during DTT incubation, n =
9, p = 0.033, Fig. 5C). As opposed to our results in CHO cells, however, current desensitization after the removal of DTT was not significantly different from preDTT control values (Td.37 = 1.42 ± 0.11 seconds, n = 9, p = 0.37, Fig. 5C). We also tested the effect of the endogenous sulfhydryl compound, glutathione. Like DTT, glutathione affected H+-gated current amplitude (Fig. 5D). Quantification revealed that glutathione and DTT induced similar potentiation (Fig. 5E).
To determine if the differences in DTT modulation between hASIC1a expressed in CHO cells and H+-gated currents in mouse neurons was due to the presence of ASIC2 subunits, we analyzed DTT effects in neurons from ASIC2 knockout mice. In hippocampal neurons, H+-gated currents are due to activation of both homomultimeric
ASIC1a channels and heteromultimeric ASIC1a/2a channels. In the absence of the
ASIC2, ASIC1a homomultimeric channels predominate (Askwith et al. 2004). We found
that H+-gated currents in ASIC2 knockout neurons were potentiated in the presence of
DTT and remained potentiated following the removal of the sulfhydryl reagent (184 ± 19
%, p = 0.008 during DTT incubation, 135 ± 11 %, p = 0.023 after DTT incubation and
washout, n = 6, Fig. 5F ~ G). The Td.37 was also increased in the presence of DTT, but returned to preDTT levels with the removal of DTT from solution (Fig. 5H). These
33 results indicate that ASIC2 subunits are not responsible for the difference in DTT effects
between CHO cells expressing human ASIC1a and hippocampal neurons. This suggests
that other factors such as species differences between mouse and human ASIC1a are
responsible.
2.3.5. DTT potentiates mouse ASICs in CHO cells
The presence of DTT affected H+-gated currents in mouse neurons differently from H+-gated currents in CHO cells expressing human ASIC1a. To determine whether
species differences between mouse and human ASIC1a impacts DTT-induced
potentiation, we analyzed the effects of DTT on mouse homomeric ASIC1a and
heteromeric ASIC1a/2a expressed in CHO cells. Whereas DTT increased pH 6.5-
activated hASIC1a currents two fold, mASIC1a pH 6.0-activated currents increased 7
fold in the presence of DTT (716 ± 137 % of preDTT control, n = 9, p = 0.002, Fig. 6A).
After a 3 minute DTT incubation and 1 minute washout, DTT-induced effects decreased
but currents remained potentiated compared to preDTT control (199 ± 33 % of preDTT
control, n = 9, p = 0.014, Fig. 6A). The presence of DTT also slowed desensitization of
mASIC1a (Td .37 = 1.19 ± 0.15 seconds for preDTT control, 1.95 ± 0.35 seconds during
DTT incubation, n = 5, p = 0.021, Fig. 6A). After removal of DTT, however, desensitization was not significantly different from preDTT control (1.36 ± 0.23 seconds, n = 5, p = 0.39). Therefore, species differences between mouse and human ASIC1a impact DTT-induced potentiation of ASIC1a currents.
We also analyzed the effects of DTT on mouse ASIC1a/2a heteromultimeric channels (Fig. 6B). When DTT was applied to CHO cells expressing both mASIC1a and
34 mASIC2a channels, H+-gated currents increased two fold (204 ± 32% of preDTT control, n = 12, p = 0.0007, Fig. 6B). Thus, the effect of DTT on mASIC1a/2a pH 6.0-activated currents is substantially smaller than the effect of DTT on homomultimeric mASIC1a pH
6.0-activated currents (p = 0.005). After the removal of DTT, H+-gated currents
remained increased albeit to a much lesser extent (128 ± 9 % after the washout of DTT, n
= 12, p = 0.012, Fig. 6B). Desensitization of mASIC1a/2a channels was also slowed
during DTT incubation and after washout (Td .37 = 1.20 ± 0.08 seconds for preDTT control, 1.88 ± 0.27 seconds during DTT incubation, p = 0.027, 1.51 ± 0.14 seconds after the washout, p = 0.020, n = 7, Fig. 6B).
DTT potentiation of H+-gated currents from other ASIC subunits was observed as
well. We analyzed the effect of DTT on mouse ASIC2a homomultimeric channels
(mASIC2a) and found that DTT incubation caused an increase in mASIC2a current
amplitude which persisted after the removal of DTT (125 ± 10 % of preDTT control, n =
8, p = 0.038, Fig. 7A). We also analyzed the effect of DTT on two ASIC subunits
prominently expressed in sensory neurons, ASIC1b (a splice variant of ASIC1a), and
ASIC3. DTT affected mouse ASIC1b currents similarly to mASIC1a. The presence of
DTT induced an 8 fold increase in current amplitude (826 ± 135% of preDTT control, n =
8, p = 0.0004, Fig. 7B). After the removal of DTT, currents remained potentiated (230 ±
34 %, n = 8, p = 0.007). H+-gated currents from cells expressed mouse ASIC3 (mASIC3)
were also impacted by DTT. The presence of DTT caused pH-6.0 activated mASIC3
currents to increase to 357 ± 80 % of preDTT control (n = 10, p = 0.009, Fig. 7C). After removal of DTT, these currents remained elevated (288 ± 68 %, n = 10, p = 0.017).
Therefore, DTT potentiated the peak current amplitude of all ASICs examined.
35
2.3.6. The metal chelator TPEN mimics the transient effect of DTT on hASIC1a.
Sulfhydryl compounds display two chemical properties which impact ion channel
function. They modify the structure and function of proteins by reducing amino acid
residues and disulfide bonds. They also chelate transition metal ions such as Cu2+, Fe2+,
Mn2+, Ni2+, and Zn2+. ASIC1a is inhibited by extracellular Zn2+ in the nanomolar range
(Chu et al. 2004). Zinc decreases the apparent sensitivity of ASIC1a for protons and ambient Zn2+ concentration in the extracellular solution is enough to tonically inhibit
ASIC1a (Chu et al. 2004). We hypothesized that the acute potentiation of ASIC1a that is
observed during DTT incubation and disappears after washing was due to Zn2+ chelation.
To test this hypothesis, we used TPEN, a compound which chelates transition metal ions including zinc (Fig. 8). In the presence of 10 µM TPEN, peak H+-gated current amplitude of hASIC1a increased (Fig. 8A). The potentiation of hASIC1a current amplitude in the presence of either TPEN or DTT was remarkably similar (198 ± 18 % for TPEN, 201 ± 13 % for DTT, n = 14, p = 0.99, Fig. 8A). Like DTT, TPEN also slowed the desensitization of hASIC1a, and the Td .37 in the presence of TPEN was not significantly different from the Td .37 in the presence of DTT (4.25 ± 0.38 for DTT, 4.44 ±
0.23 for TPEN, n = 5, p = 0.50, Fig. 8B). However, TPEN-induced potentiation did diverge from DTT potentiation in one respect. Potentiation disappeared completely after
the removal of TPEN whereas currents remained potentiated following the removal of
DTT (107 ± 5 % and 150 ± 12 % after the washout of TPEN or DTT respectively, p =
0.013, n = 7, Fig. 8A). These results are consistent with the proposed mechanism of action of these two compounds. While TPEN simply chelates transition metals, DTT can
36 both chelate transition metals and covalently modify amino acids residues to induce long-
lasting effects on channel function.
To determine whether the effect of TPEN and DTT are additive, we studied the
consequences of co-application of both compounds. We found that application of DTT in the presence of TPEN caused no additional increase in hASIC1a current activated by pH
6.5 solutions (231 ± 34 % for TPEN, 246 ± 22 % for TPEN + DTT, n = 6, p = 0.49, Fig.
8C). Similar results were attained when the activating pH was 6.7 (265 ± 55% for TPEN,
263 ± 49% for TPEN + DTT, n = 5, p = 0.82, Fig. 8C). These results indicate that
potentiation of hASIC1a currents in the presence of DTT is due to metal chelation.
2.3.7. Mutation K133R attenuates both DTT and TPEN-induced potentiation of human
ASIC1a
If potentiation of hASIC1a currents in the presence of DTT is due to zinc
chelation, then mutant channels insensitive to extracellular zinc should be unaffected
acutely by DTT. In mouse ASIC1a, a substitution of lysine 133 to an arginine eliminates
high-affinity Zn2+ inhibition (Chu et al. 2004). We made this substitution in human
ASIC1a (K113R), transfected it into CHO cells, and analyzed the effects of TPEN and
DTT on H+-gated currents (Fig. 9). Similar to previous reports on mouse ASIC1a with this substitution, the pH dose-response curve of K133R was similar to that of hASIC1a
(data not shown). TPEN-induced potentiation of H+-gated currents from K133R was dramatically reduced compared to wild-type hASIC1a (138 ± 6 % for K133R and 202 ±
23 % for hASIC1a, p = 0.019, Fig. 9A ~ C). Thus, K133R shows significantly reduced
tonic inhibition by zinc, but some transition metal inhibition still remains (138 ± 6 %, n =
37 9, p = 0.0004). DTT-induced potentiation was also reduced in this mutant channel (Fig.
9B). In the presence of DTT, the increase in H+-gated currents of K133R was only 131 ±
10 % compared to hASIC1a potentiation of 204 ± 15 % (n = 9, p = 0.0007, Fig. 9C).
After the removal of DTT, H+-gated currents of K133R were only 113 ± 4 % of the preDTT control whereas hASIC1a was 151 ± 10 % of preDTT control (p = 0.023, Fig.
9C). These results indicate that the long-lasting effect of DTT, which we ascribe to redox
modulation of the channel, is also compromised in K133R. Thus, both TPEN and DTT-
induced potentiation is attenuated in the K133R mutant of hASIC1a. These results
indicate that lysine 133 affects both TPEN and DTT potentiation and suggests that these
compounds have similar mechanisms of action.
2.3.8. ASIC1b and ASIC3 are potentiated by the metal chelator TPEN
Our studies suggest that the presence of DTT potentiates ASIC1a by chelating
transition metals ions which tonically inhibit H+-gated currents (Fig. 8 ~ 9). Consistent with this idea, zinc inhibits ASIC1a currents at concentrations which are normally present in extracellular solutions (Chu et al. 2004), and our results indicate that the metal chelator
TPEN mimics DTT-induced potentiation (Fig. 8A ~ B). The effects of both TPEN and
DTT are not additive (Fig. 8C) suggesting that they act through a common mechanism.
However, DTT potentiates ASIC1b and ASIC3 (Fig. 7B~C), which are not reported to be sensitive to extracellular zinc (Chu et al. 2004). To examine whether these channels are
affected by metals and whether transient potentiation by DTT is due to metal chelation,
we expressed mASIC1a, mASIC1b and mASIC3 in CHO cells and then analyzed the
effect of TPEN and DTT on H+-gated currents (Fig. 10). Like hASIC1a, 10 µM TPEN
38 potentiated mASIC1a current and the application of DTT in the presence of TPEN did
not result in an additional increase in mouse ASIC1a current (581 ± 122% for TPEN, 579
± 134% for TPEN + DTT, n = 5, p = 0.96, Fig. 10A). Mouse ASIC1b was also enhanced
by TPEN in concentration-dependent manner (593 ± 149 %, n = 6, p = 0.02 for 10 µM
TPEN, and 882 ± 213 %, n = 4, p = 0.03 for 100 µM TPEN, Fig. 10B). DTT further
increased ASIC1b currents in the presence of 10 µM TPEN, but not in 100 µM TPEN (n
= 5, p = 0.008 for 10 µM TPEN, and n = 4, p = 0.84 for 100 µM TPEN compared with
TPEN alone, Fig. 10B). These results suggest that mASIC1a and mASIC1b differ in
their sensitivity to metal chelators and that the metal regulatory site is altered between
these splice variants. TPEN (10 µM) also increased ASIC3 currents (164 ± 14 %, n = 8,
p = 0.003, Fig. 10C). When co-applied with 10 µM TPEN, there was no additive enhancement of ASIC3 currents (n = 4, p = 0.18 compared with TPEN alone, Fig. 10C).
These results suggest that, like ASIC1a, the chelation of transition metal ions may underlie potentiation of ASIC1b and ASIC3 by DTT.
2.4. DISCUSSION
The acid-sensing ion channels are important for normal behavior and sensory transduction (Chen et al. 2002; Jones et al. 2004; Mogil et al. 2005; Price et al. 2000;
Price et al. 2001; Wemmie et al. 2003; Wemmie et al. 2002; Wemmie et al. 2004). In addition, calcium-permeable ASIC1a contributes to neuronal death following ischemia in the brain (Xiong et al. 2004). Therefore, understanding the factors which influence
ASIC1a activity may lend insight into the modulators which control cell death following 39 ischemia and stroke. In this study, we show that the activity of ASICs, including human
ASIC1a, is enhanced by sulfhydryl compounds. Since the extracellular concentration of
sulfhydryls can change in physiologic and pathological states (Andine et al. 1991;
Landolt et al. 1992; Slivka and Cohen 1993; Zangerle et al. 1992), the modulation of
ASICs by these compounds may have a strong impact on ASIC function in the central
and peripheral nervous system.
Similar to previous studies in mouse sensory neurons and cortical neurons
(Andrey et al. 2005; Chu et al. 2006), we found that sulfhydryl agents enhanced ASIC
currents. DTT shifts the pH-dose response curve of both human and mouse ASIC1a such
that less acidic pHs are required to activate H+-gated currents (Chu et al. 2006). Thus, sulfhydryl-induced potentiation of ASIC1a is conserved between species although there are differences in DTT modulation of human and mouse ASICs. We observed extremely robust (7 fold) increase of mouse ASIC1a currents in the presence of DTT whereas human ASIC1a was potentiated only two fold. This difference could be ascribed to the different pH solutions used to activate currents (pH 6.5 for human ASIC1a and pH 6.0 for mouse ASIC1a). However, at no place on the hASIC1a pH dose-response curve did we observe a 7 fold potentiation of current by DTT. Human and mouse ASIC1a channels are highly conserved with only 11 amino acid differences. Our results suggest that at least one of these amino acid differences affects how sulfhydryl reagents impact channel characteristics.
We also observed differences in potentiation between H+-gated currents of mouse hippocampal neurons and heterologously expressed mouse ASICs. In particular, DTT- induced potentiation of H+-gated currents in ASIC2 knockout neurons (which have
40 predominantly mASIC1a-like currents) was much smaller than that of mASIC1a
expressed in CHO cells. This difference could be due to the state of neuronal ASIC
channels (some may already be reduced or free of metal), the presence of other ASIC subunits, or neuron-specific modification of ASIC1a.
Sulfhydryl compounds modulate many ion channels by reducing disulfide bonds and chelating transition metal ions (Lipton et al. 2002; Wilkins and Smart 2002). For example, DTT produces an increase in NMDA receptor current which persists even after removal of DTT. It is thought that this effect is produced by reduction of disulfide bonds between two cysteine residues (C744 and C798) in the NR1 subunit (Sullivan et al.
1994). DTT also increases NMDA receptor activity by chelating transition metals and relieving tonic Zn2+-inhibition (Paoletti et al. 1997). Our data suggest that DTT affects
ASIC1a in a similar manner. In the presence of DTT or glutathione, the peak current
amplitude of hASIC1a currents is potentiated and desensitization is slowed. This effect is
mimicked by the addition of the metal chelator TPEN, and the addition of DTT with
TPEN does not increase currents further. Furthermore, the increase in currents induced by
DTT was significantly attenuated in a mutant channel (K133R) that is less sensitive to
extracellular zinc. Together, these results indicate that DTT potentiates hASIC1a by
chelating transition metals. Sulfhydryls also potentiate ASIC1b and ASIC3 currents.
Like ASIC1a, ASIC1b and ASIC3 currents were not further potentiated by DTT in the
presence of TPEN suggesting that the presence of DTT increases currents by chelating
transition metal ions which tonically inhibit ASIC1b and ASIC3.
Human ASIC1a currents remain potentiated following DTT incubation and
removal. This long-lasting potentiation is reversed by the addition of DTNB which
41 covalently modifies free sulfhydryl groups. Additionally, this long-lasting effect is not induced by treatment with TPEN alone. These results suggest that DTT impacts peak current amplitude of hASIC1a by reducing amino acid residues within the ASIC1a
protein as well as metal chelation. The fact that all ASICs examined showed long-lasting
potentiation after the removal of DTT suggests that the residue(s) undergoing redox
modulation are conserved among ASIC subunits. Since glutathione is not readily
membrane permeable, residues within the extracellular domain are likely targets for
sulfhydryl-induced reduction. Free and disulfide-bonded cysteine residues are major sites for redox modulation by oxidizing and reducing agents (Lipton et al. 2002). ASICs have fourteen conserved cysteine residues within their extracellular domain
(Kellenberger and Schild 2002). These residues are thought to form disulfide bonds which impact the tertiary structure. Conserved cysteine residues are essential for cell- surface expression of the related DEG/ENaC channel, the epithelial sodium channel
(Firsov et al. 1999). The role of the conserved cysteine residues in the extracellular domain of ASICs has not yet been reported. Our results suggest that modification of these cysteine residues in ASICs impacts channel function.
Although all ASICs showed long-lasting potentiation after DTT incubation, only
ASIC1a, 1b and 3 displayed increased potentiation in the presence of DTT compared to the potentiation following DTT removal. These three channels were also potentiated by
TPEN, suggesting that they are tonically inhibited by basal concentrations of transition metals. Either TPEN or DTT chelates metals and frees the ion channel from metal inhibition. It is known that ASIC1a channels are inhibited by extracellular zinc at concentrations which are typically present in experimental solutions (Chu et al. 2006).
42 Like DTT, zinc chelation with TPEN is known increase the apparent proton sensitivity of
ASIC1a (Chu et al. 2004). The removal of zinc, however, has not been reported to impact channel desensitization and we observe robust changes in desensitization of human ASIC1a with the addition of TPEN or DTT. Recent work has shown that lead, cadmium, nickel, and copper inhibit ASIC1a currents (Staruschenko et al. 2006; Wang et al. 2006; Wang et al. 2007). Furthermore, we found that TPEN potentiated ASIC1b and
ASIC3 channels reported to be insensitive to zinc. It is well known that TPEN, DTT, and glutathione chelate other transition metal ions as well or even better than zinc. For
2+ example, the stability constants (log10 KC ) for the chelation by TPEN are 20.6 for Cu ,
14.6 for Fe2+, 10.3 for Mn2+, 18.0 for Ni2+, and 18.0 for Zn2+ (Smith and Martell 1974).
2+ 2+ 2+ 2+ 2+ For the chelation of Cd , Cu , Ni , Pb , and Zn , the stability constants log10 KC
(DTT) = 14.6, 15.3, 10.7, 13.7, 11.1, and log10 KC (glutathione) = 10.2, 15.5, 10.4, 10.6,
8.0 respectively (Krezel et al. 2001; Smith and Martell 1974). Our data suggests that
ASICs may be sensitive to one of these metals in addition to zinc. Further experiments
are necessary to determine the identity of the metal responsible for the basal inhibition in
our experiments. We cannot exclude the possibility that our solutions contain small
amounts of other transition metals which inhibit ASIC currents.
Sulfhydryl modification of ASICs has been described previously (Andrey et al.
2005; Chu et al. 2006). Although both papers reported potentiation of H+-gated currents
by DTT, the specific changes in H+-gated current characteristics and the proposed
mechanisms are different. For example, Chu et al. reported that DTT potentiates H+- gated currents two fold whereas Andey et al. observed only 46 % potentiation. Chu et al. focused on mouse ASIC1a and H+-gated currents in neurons from the cortex. Andrey et
43 al. worked in sensory neurons which express ASIC1b and ASIC3 in addition to ASIC1a
and ASIC2. Our results may explain the discrepancies between these two papers. We
found that mASIC1b, mASIC2a, and mASIC3 were all potentiated by DTT. The results
obtained from sensory neurons may represent DTT modification of heteromultimeric
channels composed of these other subunits which may have very different levels of
potentiation compared to mASIC1a. Chu et al. also found that DTT shifts the pH dose- response curve of mASIC1a-containing channels and does not impact peak current amplitude when the channel is activated by a maximal acidic stimulus (Chu et al. 2006).
However, Andrey et al. found that DTT did not affect the pH dose-response, and increased the peak current amplitude of pH 4.5-activated currents in sensory neurons
(Andrey et al. 2005). DTT-induced potentiation of H+-gated currents in sensory neurons
may be due to potentiation of ASIC1b, ASIC2a, or ASIC3-containing subunits which
may be impacted differently than ASIC1a and show a different pH sensitivity of
potentiation.
Our results also diverge from previous reports. We find that all ASICs tested
were potentiated by DTT. However, Chu et al. reported that only ASIC1a-containing channels are affected by DTT (Chu et al. 2006). In addition, we report that the largest
potentiation by DTT is due to metal chelation. Chu et al. reported that the effect of DTT
was not due to chelation of transition metals (Chu et al. 2006). The exact reason for these
discrepancies is unclear although we can offer several explanations. First, the majority of
our studies focused specifically on ASIC potentiation in the presence of sulfhydryls.
Second, we used the metal chelator TPEN which has a very high affinity for metals.
Previous publications used tricine, a relatively weak chelator of transition metals (Ahmed
44 2003; Chu et al. 2006). For the chelation of Cd2+, Ni2+, Pb2+, and Zn2+, the stability constants log10 KC (tricine) = 4.4, 5.5, 4.3, and 5.6 respectively (Ahmed 2003). We
observed that DTT further increased hASIC1a currents in the presence of 10 mM tricine
(121 ± 5 %, n = 5, p = 0.015), but not 10 µM TPEN. This suggests that the use of TPEN
rather than tricine is responsible, at least in part, for the discrepant results (Chu et al.
2006). One final possibility is that our solutions may contain a different basal
concentration of metals. As stated before, our experiments did not distinguish which
transition metal is inhibiting ASIC activity. For example, the concentration of
contaminating zinc is usually in the nanomolar range for experimental solutions (Amar et
al. 2001). We found that our solutions contain 150 nM zinc as well as nanomolar
concentrations of chromium, copper, and nickel. It is likely that additional metals inhibit
ASIC activity. It is also likely that the types and concentration of metals will vary in solution between different laboratories. Thus, our solutions may have contained basal concentration of a metal that inhibits ASIC activity which was not present within the solutions of other investigators.
The concentration of extracellular sulfhydryls is altered in both pathological and physiological conditions. The extracellular concentration of endogenous sulfhydryl compounds increases in the ischemic brain (Andine et al. 1991; Landolt et al. 1992;
Slivka and Cohen 1993). Glutathione is abundant in neurons and glial cells and is released during ischemia from cells with damaged membranes. In addition, oxygen/glucose-deprivation opens neuronal gap junction hemichannels resulting in the loss of ionic homeostasis and the efflux of cytosolic molecules less than 1 kD in size
(Thompson et al. 2006). This raises the possibility that reduced glutathione may be
45 released from neurons through hemichannels activated during ischemia, even when cell
membranes remains intact. It is known that the extracellular concentration of glutathione
and cysteine can increase to 1 µM and 5 µM respectively in rat models of focal ischemia
(Landolt et al. 1992). This increase in sulfhydryls has been shown to impact the activity
of multiple ion channels involved in ischemia-induced neuronal death. Interestingly,
glutathione potentiates both NMDA receptors and ASICs, two channels known to
enhance excitotoxic and acidotoxic neuronal death following stroke and ischemia.
Sulfhydryl modulation of ASICs could have profound effects on neuronal death
following stroke.
Neuronal activity also induces the release of endogenous sulfhydryl compounds,
which in turn regulate ion channels involved in synaptic transmission and plasticity
(Gozlan and Ben-Ari 1995; Zangerle et al. 1992). Low levels of glutathione have
previously been linked to impaired synaptic plasticity (Almaguer-Melian et al. 2000;
Cruz-Aguado et al. 2001; Steullet et al. 2006). ASIC1a localizes to synapses and ASIC1 knockout animals show impaired performance in a number of behavioral tests of learning and memory (Wemmie et al. 2003; Wemmie et al. 2002). ASIC1 also plays a prominent role in fear-related behaviors (Wemmie et al. 2004). Interestingly, the enzyme which maintains the levels of reduced glutathione in the brain has also been linked to fear and anxiety-related behaviors in mice (Hovatta et al. 2005). Thus, the potentiation of ASICs by sulfhydryl compounds may regulate synaptic plasticity, learning and memory, and possibly fear-related behaviors.
46 A 1 mM DTT Wash Wash Wash pH 6.5
1 nA 2 sec
B 220 n = 10 *
t 200 n e
r 180 r ) l u o c r
160 t d n **
o 140 * ze * i c l
a
% 120 ( m r
o 100
N DTT 80 Wash 60 0 2 4 6 8 10 12 14 Time (min)
p < 0.0001 p < 0.01 C 250 1 mM DTT p < 0.001 p < 0.01 10 µM DTT
t 200 n e r r ) l u o c r 150
t d n o ze i c l
a 100 % ( m r o
N 50
0 PreDTT DTT DTT PreDTT DTT / wash n = 18 n = 37 n = 10
D 1 mM DTT Wash pH 6.5
0.5 nA 2 sec
Figure 2.1: DTT potentiates H+-gated currents in CHO cells expressing human ASIC1a (continued)
47 Figure 2.1: continued
(A) Representative trace of H+-gated currents before and after DTT incubation. CHO cells expressing human ASIC1a (hASICa) were exposed to extracellular acid (pH 6.5, grey bars) and 1 mM DTT for 3 minutes (black bar). DTT-free solution exchange for 1 ~
5 minutes is indicated by ‘wash’ arrows. Dotted line represents the peak current amplitude before DTT addition. (B) Time course of DTT-induced potentiation of hASIC1a H+-gated current. Extracellular pH 6.5 solutions were added at 2 minute intervals. Peak current amplitude was normalized to that of the preDTT control (100 %,
indicated by arrow) evoked just before DTT incubation (n = 10, * p < 0.05, ** p < 0.01
compared with control, paired t test). (C) Quantification of the transient (DTT) and
long-lasting effects (DTT/wash) of 1 mM and 10 µM DTT on pH 6.5-activated currents.
Data are expressed as the mean of normalized peak current amplitude before addition of
DTT, in the presence of DTT, and following wash with DTT-free solution (1 minute). P
values were determined using a paired t test. (D) Representative trace of the effect of
DTT and pH 6.5 in CHO cells transfected with pEGFP alone. Note that H+-gated current
was not evoked by acid (pH 6.5), and DTT incubation had no effect. In all experiments,
error bars represent the standard error of the mean (SEM).
48 DTT / wash DTNB / wash A pH 6.5
1 nA 2 sec B n = 12 p < 0.001 p < 0.01 140
130 t n e r r )
l 120 u o c r
t d n
o 110 ze i c l
a % ( m
r 100 o N 90 DTT DTNB Wash Wash 80 0 2 4 6 8 10 12 14 16 18 Time (min)
n = 9 ns
C D
DTNB / wash t 120 n e
pH 6.5 r 100 ) r l u o r c 80 t
n d e o 60 z c i
l
a 40 % ( m r 20 o 0.5 nA N 0 2 sec PreDTNB DTNB / wash
Figure 2.2: The long-lasting effect of DTT on hASIC1a is reversed by DTNB (continued)
49 Figure 2.2: continued
(A) Representative trace of the effect of DTT and DTNB on human ASIC1a expressed in
CHO cells. H+-gated currents were activated by pH 6.5 solution before DTT addition, after DTT addition (1 mM DTT for 3 minutes and 3 minute wash), and after DTNB addition (1 mM DTNB for 3 minutes and 3 minute wash). Note that all pH 6.5 applications were performed in the absence of DTT or DTNB. (B) Time course of the
long-lasting effects of DTT and DTNB. Peak current amplitude was normalized to that
of the preDTT control (arrow). DTT and DTNB addition are indicated by black bars.
Washes with extracellular solution free of reagents are indicated by grey bars (n = 12,
paired t test). (C) Representative trace of the effect of DTNB (1 mM) on human ASIC1a
currents without previous DTT incubation. DTNB was added for 3 minutes and washed
out for 3 minutes. (D) Quantification of the effect of DTNB alone on human ASIC1a.
pH 6.5-activated current amplitude from hASIC1a was not significantly affected (ns) by
DTNB when cells were not previously treated with DTT (n = 9, p = 0.24, paired t test).
Error bars represent the SEM.
50 100 µM A Glutathione Wash pH 6.5
1 nA 2 sec B 200 100 µM 1 µM 10 nM t
n ** e r r ) l 150 u o c
r **
t * d n * *
o ** ze i c l * * a % (
m 100 r o
N Glutathione Wash 50 0 2 4 6 8 10 12 14 Time (min) 100 C GSH GSH/wash (9) 80 ** 60
(10) 40 * * (13) (7) ∆ I (% control)
pH 6.5 20 * * (7) 0 (10) 0.01 0.1 1 10 100 [Glutathione] (µM)
Figure 2.3: Glutathione potentiates hASIC1a currents in a concentration-dependent manner (continued)
51 Figure 2.3: continued
(A) A representative trace shows the potentiation of human ASIC1a current by reduced
glutathione. H+-gated currents were activated by pH 6.5 solutions (grey bar) during glutathione incubation (100 µM for 3 min as indicated by black bar) and following removal of glutathione (1 minute wash). Washout is indicated by arrow. (B) Time course of the potentiation by 100 µM, 1 µΜ, and 10 nM glutathione. Peak amplitudes were normalized to pre-glutathione control (arrow). Glutathione incubation is indicted by black bar. Solution exchange (wash) is indicated by grey bar. n = 7 ~ 13 for each group,
* p < 0.05, ** p < 0.01 compared to control, paired t test. (C) The concentration- response relationship of glutathione. The changes of current amplitudes (∆ I pH 6.5) during glutathione incubation (GSH) and after 3 minute washout (GSH/wash) are expressed as the percentage of pre-glutathione control (* p < 0.05, ** p < 0.01 compared with control, paired t test). A one-way ANOVA revealed a significant difference between ∆ I pH 6.5 during glutathione incubation and ∆ I pH 6.5 after the washout of glutathione (F (1, 54) =
4.49 and p = 0.04). Both transient and long-lasting effects of glutathione on ∆ I pH 6.5 were significantly different depending upon the concentration (one-way ANOVA, F (2, 29)
= 5.83 and p = 0.007 for the transient effect, F (2, 21) = 3.61 and p = 0.04 for the long- lasting effect). Numbers in the parentheses represents the numbers of cells examined.
52 A B n = 6 p < 0.01 p < 0.05 p < 0.05 6 1 mM DTT Wash pH 6.5 5
4
3 (sec)
d .37 2 1 nA T 2 sec 1 0 PreDTT DTT DTT/wash ) l 250 pH 6.5
C o D r * t (10) pH 5.0 1.2 PreDTT n
o 200 DTT c
1
% DTT/wash (
** *** t (10) 0.8
n 150 e r r (6) (5) * u pH 5 0.6 c
100 I / d
e 0.4 ** z i l
a 50 *
m 0.2 r o
N 0 0 PreDTT DTT DTT/wash 5.8 6 6.2 6.4 6.6 6.8 7 7.2 7.4 pH
Figure 2.4: DTT slows the desensitization and shifts the pH dose-response of hASIC1a (continued)
53 Figure 2.4: continued
(A) Representative recording showing desensitization kinetics before, during and after
DTT incubation. pH 6.5 addition is indicated by grey bars, and DTT incubation (1 mM)
is indicated by black bars. Removal of DTT-containing solution is indicated by wash
arrow. Dotted line represents the peak current amplitude before DTT application. (B)
Quantification of the DTT-induced change in desensitization. Desensitization was
quantified as the time for hASIC1a currents to decrease to 37 % of the peak amplitude (Td
.37). Statistical significance was determined using a paired t test. Error bars represent the
SEM. (C) Quantification of the peak current amplitude of pH 6.5 or pH 5.0-activated currents before, during, and after DTT addition. Peak current amplitudes were normalized to the preDTT control values. Numbers in the parenthesis represents the numbers of cells examined. * p < 0.05, ** p < 0.01 compared with preDTT control, paired t test. (D) Change in pH dose-response relationship of human ASIC1a by DTT.
H+-gated currents were evoked by pH 6.0, 6.5, 6.7, and 6.9 solutions without DTT incubation (preDTT), during DTT incubation (DTT), or after the washout of DTT
(DTT/wash). Peak current amplitudes were normalized to currents evoked by pH 5.0 at the indicated condition (n = 4 ~ 8 for each data point and presented as the mean ± SEM).
Normalized currents at pH 6.5 and pH 6.7 were significantly larger in DTT-treated cells
compared to cells without DTT incubation (* p < 0.05, ** p < 0.01, *** p < 0.001,
unpaired t test).
54 1 mM DTT Wash Wash Wash A pH 6.0
0.2 nA
200 2 sec B DTT (n=11) Mock (n=8) C 2.5 n = 9 *** * t n
e 2 r 150 r ) l u o c r
t 1.5
d *** n o ze (sec)
i *** c l
a % ** d .37 1 ( m T
r 100 o
N 0.5
DTT Wash 0 50 PreDTT DTT DTT/wash 0 5 10 15 20 Time (min) D 100 µM E Mock (n=10) Glutathione Wash 1 mM DTT (n=11) # pH 6.0 100 µM GSH (n=9) 200 # *** ***
t
n 150 e r
r *** ) u 0.1 nA x
c ** T
e 2 sec d
e 100 z Pr i l % a F. ASIC2 KO ( m
1 mM DTT Wash r pH 6.5 o 50 N
0 During DTT or GSH After washout
0.5 nA 2 sec G. ASIC2 KO H. ASIC2 KO 250 # n = 6 5 n = 5
t ** **
n 200
e 4 r r ) l u o c r * 150 t 3 d n o ze (sec) i c l 100 a 2 % d .37 ( m T r o
N 50 1
0 0 PreDTT DTT DTT/wash PreDTT DTT DTT/wash
Figure 2.5: DTT and glutathione potentiate H+-gated currents in hippocampal neurons (continued) 55 Figure 2.5: continued
(A) Representative traces show the potentiation of H+-gated currents activated at pH 6.0 by 1 mM DTT (4 minutes). (B) Time course of H+-gated currents at pH 6.0 in
hippocampal neurons in the presence of DTT and following DTT removal (filled circles)
or mock-treated neurons (open circles). Note the rundown of H+-gated currents in mock- treated neurons. Peak current amplitudes were normalized to that of the preDTT or pre- vehicle control (100 %, arrow). ** p < 0.01, *** p < 0.001 compared with mock-treated control, unpaired t test. (C) The effect of DTT on desensitization of H+-gated currents. *
p < 0.05, paired t test. (D) Representative trace of the effect of 100 µM glutathione on cultured hippocampal neurons (4 minute glutathione incubation and 1 minute washout).
(E) Quantification of transient and long-lasting effects of 1 mM DTT and 100 µΜ
glutathione on H+-gated currents in hippocampal neurons (** p < 0.01, *** p < 0.001
compared with mock-treated group, unpaired t test, # p < 0.05 transient versus long-
lasting effects, paired t test). The transient effect was measured during DTT or
glutathione incubation (4 minutes), and the long-lasting effect was measured after
washout (1 minute). (F) Representative trace of the effect of DTT on H+-gated currents from ASIC2 knockout neurons. (G) Quantification of the effect on pH 6.5-induced current amplitude. * p < 0.05, ** p < 0.01 compared with preDTT control, # p < 0.05
transient versus long-lasting effects, paired t test. (H) The effect of DTT on desensitization of H+-gated currents in ASIC2 knockout neurons. ** p < 0.01 compared
with preDTT control, paired t test.
56 A. mASIC1a #
1 mM t 1000 n = 9 2.5 n = 5 n *
DTT Wash e ** r ) r
pH 6.0 l 800 2 u o r c t
n d 600 1.5 e o z c i
(sec) l 400 1 a d .37 % ( T m
r * 200 0.5
2 nA o N 2 sec 0 0 PreDTT DTT DTT/wash PreDTT DTT DTT/wash
# B. mASIC1a/2a #
1 mM t 250 n = 12 2.5 n = 7 Wash n ** DTT e * r )
pH 6.0 r
l 200 2 u o
0 r c * t
n d 150 1.5
e * o z (sec) c i
l
100 d .37 1 a % T ( m r 50 0.5
1 nA o 2 sec N 0 0 PreDTT DTT DTT/wash PreDTT DTT DTT/wash
Figure 2.6: DTT-induced potentiation of H+-gated currents of CHO cells expressing mouse ASIC1a-containing channels
Representative traces and quantification of H+-gated currents in CHO cells expressing
(A) mouse ASIC1a homomultimers or (B) mouse ASIC1a/2a heteromultimers. DTT was applied for 3 minutes and washed out for one minute. pH 6.0 solution was applied every
2 ~ 2.5 minutes (in presence or absence of DTT as indicated) to activate ASIC currents.
Desensitization was quantified by measuring the Td.37 as in Fig. 4. * p < 0.05, ** p < 0.01 compared with preDTT control, # p < 0.05 comparing pH 6.0-activated currents in presence of DTT to pH 6.0 currents activated after the removal of DTT, paired t test.
57 1 mM t 150
n n = 8
A. mASIC2a DTT Wash e * pH 4.0 r r ) l u o c r
t 100 d n e o z c i
l a %
( 50 m r
1 nA o N 2 sec 0 PreDTT DTT DTT/wash
1200 ## t n = 8
1 mM n
B. mASIC1b e DTT Wash r 1000 r ) ***
pH 6.0 l u o c r 800
t d n e o 600 z c i
l a
% 400 ( m 0.5 nA r ** o 200 2 sec N 0 PreDTT DTT DTT/wash
#
C. mASIC3 1 mM t 500 n n = 10
DTT Wash e
r **
pH 6.0 r )
l 400 u o c
r *
t
d 300 n e o z c i
l 200 a % ( m r 100
0.5 nA o N 2 sec 0 PreDTT DTT DTT/wash
Figure 2.7: DTT potentiates mouse ASIC2a, ASIC1b and ASIC3 H+-gated currents
Representative trace and quantification of the effect of DTT on (A) mouse ASIC2a, (B) mouse ASIC1b, and (C) mouse ASIC3 expressed in CHO cells. ASIC currents were activated by pH 4.0 or pH 6.0 solutions every two minutes. DTT (1 mM) was applied for
3 minutes. During this 3 minute time period, ASIC currents were evoked by application of pH 4.0 or pH 6.0 solutions in the presence of DTT. DTT was removed by exchanging the bath solution and superfusing the cell with DTT-free solution for one minute. pH 4.0 or pH 6.0 solution (in the absence of DTT) was then applied. * p < 0.05, ** p < 0.01, *** p < 0.001 compared with preDTT control. # p < 0.05, ## p < 0.01 comparing pH 6.0 amplitude in the presence of DTT and after DTT removal, paired t test.
58 TPEN ns 250 A 10 µM TPEN Wash 1 mM DTT Wash DTT pH 6.5 ******
p < 0.05 t 200 n e r ) r l
u ** o r c 150 t
n d e o z c i
l 100 a % ( m
1 nA r o
2 sec N 50
0 PreTx TPEN or DTT Wash Control n = 14 n = 7 B ns 5 n = 5 10 µM TPEN 1 mM DTT Overlay *** *** pH 6.5 (TPEN / DTT) 4
3 (sec) 2 d .37 T 1 nA 1 2 sec
0 PreTx TPEN DTT Control ns C TPEN TPEN+DTT pH 6.5 pH 6.5 (n=6) ns 350 pH 6.7 (n=5) * * 300
t ** 1 nA n * e 250 r ) r l u 2 sec o r c t 200 n d e o z c i
TPEN TPEN+DTT l 150
pH 6.7 a % ( m r 100 o N 50
1 nA 0 PreTx TPEN TPEN+DTT 2 sec Control
Figure 2.8: The metal chelator TPEN mimics the transient effect of DTT (continued)
59 Figure 2.8: continued
(A) Representative trace of human ASIC1a H+-gated currents in the presence of TPEN
(10 µM) or DTT (1 mM). Quantification of the peak current amplitude of pH 6.5-
activated currents in the presence of TPEN or DTT and after washout (** p < 0.01, *** p
< 0.001, paired t test). There was no significant difference (ns) in the acute potentiation
by TPEN and by DTT. (B) Representative trace and quantification of the change in
desensitization when either TPEN or DTT is applied (*** p < 0.001, paired t test). (C)
Representative trace of the effect of applying both TPEN and DTT concurrently, and
quantification of the potentiation of hASIC1a by TPEN or TPEN + DTT. Peak amplitude
values were normalized to that of the pretreatment control (PreTx, 100 %). There was no significant difference (ns) in the potentiation by TPEN and by TPEN + DTT. * p < 0.05,
** p < 0.01, paired t test
60 10 µM 1 mM A. hASIC1a TPEN DTT pH 6.5 pH 6.5 Wash
0.5 nA 1 nA 2 sec 2 sec 10 µM 1 mM B. K133R TPEN DTT Wash -hASIC1a pH 6.5 pH 6.5
2 nA 2 sec
p < 0.05 hASIC1a p < 0.001 C 250 K133R (1*1*)
) *** l (11) o r
t p < 0.05
n 200 o c (4*) % (
*** t 150 (9) (9*) n e
r ** r (8) u c
d 100 e z i l a m
r 50 o N
0 PreTx TPEN DTT DTT / wash Control
Figure 2.9: Lysine 133 mutation reduces both TPEN and DTT-induced potentiation of hASIC1a
Representative traces of the potentiation of (A) hASIC1a and (B) K133R-hASIC1a by
TPEN (10 µM, 1 minute) and DTT (1 mM, 3 minutes). H+-gated currents at pH 6.5 were recorded before, during, and after incubation of TPEN or DTT. (C) Quantification of the effect of TPEN and DTT on hASIC1a and K133R. Numbers in the parentheses represents the numbers of cells examined (* p < 0.05, ** p < 0.01, *** p < 0.001 compared to pretreatment control, paired t test).
61 ns
10 µM TPEN t 700 n = 5 A. mASIC1a n * TPEN + DTT e *
r 600 )
pH 6.0 r l u o 500 r c t
n d 400 e o z c i
300 l a %
( 200 m r
0.5 nA o 100 N 0 2 sec PreTx TPEN TPEN Control /DTT
10 µM p < 0.05 TPEN
10 µM TPEN B. mASIC1b t 1400 n = 5 TPEN + DTT n ** pH 6.0 e
r 1200 ) r l u o 1000 r c t
n d 800
e * o z c i
600 l a %
( 400 m r
0.5 nA o 200 N 0 2 sec PreTx TPEN TPEN Control /DTT
100 µM TPEN ns
t 1400 100 µM
TPEN + DTT n TPEN pH 6.0 e 1200 r )
r n = 4
l * * u
o 1000 r c t
n d 800 e o z c 600 i
l a % 400 ( m 1 nA r 200 o 2 sec N 0 PreTx TPEN TPEN Control /DTT
ns 250 n = 4 10 µM TPEN t
C. mASIC3 TPEN + DTT n
pH 6.0 e *
r 200 ) r l ** u o r c t 150 n d e o z c
i 100
l a % (
m 50 r
0.5 nA o
N 0 2 sec PreTx TPEN TPEN Control /DTT
Figure 2.10: Effect of TPEN and DTT on mouse ASIC1a, ASIC1b and ASIC3 (continued)
62 Figure 2.10: continued
Representative trace and quantification of TPEN and DTT addition on (A) mASIC1a, (B) mASIC1b, and (C) mASIC3 expressed in CHO cells. H+-gated currents were activated by pH 6.0 solutions in the presence of the indicated compounds. TPEN (10 µM or 100
µM) was applied for 1 minute at pH 7.4 prior to activation of H+-gated currents. When
DTT and TPEN were both present during the acid stimulus (TPEN+DTT), TPEN was applied for 1 minute at pH 7.4, then DTT (1 mM) was added in the continued presence of
TPEN at pH 7.4 for an additional 1 minute before acid-dependent activation. * p < 0.05,
** p < 0.01 compared to pretreatment control values (no DTT or TPEN), paired t test.
63
CHAPTER 3
PRESYNAPTIC RELEASE PROBABILITY IS INCREASED IN HIPPOCAMPAL NEURONS FROM ASIC1 KNOCKOUT MICE
The acid-sensing ion channels (ASICs) are H+-gated channels that produce transient cation currents in response to extracellular acid. ASICs are expressed in neurons throughout the brain, and ASIC1 knockout mice show behavioral impairments in learning and memory. The role of ASICs in synaptic transmission, however, is not thoroughly understood. We analyzed the involvement of ASICs in synaptic transmission using microisland cultures of hippocampal neurons from wild-type and ASIC knockout mice. There was no significant difference in single action potential (AP)-evoked excitatory postsynaptic currents (EPSCs) between wild-type and ASIC knockout neurons.
However, paired-pulse ratios (PPR) were reduced, and spontaneous miniature EPSCs
(mEPSCs) occurred at a higher frequency in ASIC1 knockout neurons compared to wild- type neurons. The progressive block of NMDA receptors by an open channel blocker,
MK-801, was also faster in ASIC1 knockout neurons. The amplitude and decay time constant of mEPSCs, as well as the size and refilling of the readily-releasable pool, were similar in ASIC1 knockout and wild-type neurons. Finally, the release probability, which
64 was estimated directly as the ratio of AP-evoked to hypertonic sucrose-induced charge
transfer, was increased in ASIC1 knockout neurons. Transfection of ASIC1a into ASIC1
knockout neurons increased the PPRs, suggesting that alterations in release probability
were not the result of developmental compensation within the ASIC1 knockout mice.
Together, these findings demonstrate that neurons from ASIC1 knockout mice have an
increased probability of neurotransmitter release, and indicate that ASIC1a can affect
presynaptic mechanisms of synaptic transmission.
3.1. INTRODUCTION
Acid-sensing ion channels (ASIC) are voltage-independent cation-permeable ion
channels activated by extracellular acidosis (Krishtal 2003; Waldmann 2001; Wemmie et
al. 2006). These H+-gated channels are members of the degenerin/epithelial Na+ channels
(DEG/ENaC) family (Kellenberger and Schild 2002). Four ASIC genes encode at least six subunits (ASIC1a, 1b, 2a, 2b, 3, and 4) through alternative splicing. ASIC subunits are expressed in central and peripheral neurons, and the characteristics of H+-gated currents are determined by the ASIC subunits expressed within the cell. ASIC1a,
ASIC2a, and ASIC2b are expressed throughout the brain with particularly high abundance in the cerebral cortex, hippocampus, amygdala, olfactory bulb, and the cerebellum (Garcia-Anoveros et al. 1997; Lingueglia et al. 1997; Price et al. 1996;
Waldmann et al. 1997b; Waldmann et al. 1996). ASIC1a forms heteromeric channels with ASIC2a, as well as Ca2+-permeable homomeric channels (Askwith et al. 2004;
Benson et al. 2002; Gao et al. 2007; Hesselager et al. 2004; Yermolaieva et al. 2004).
65 ASIC1a is enriched in synaptosomal fractions and expressed in dendritic spines (Hruska-
Hageman et al. 2002; Wemmie et al. 2002). ASIC1 knockout mice display impaired
spatial learning, eye-blink conditioning, and fear conditioning (Wemmie et al. 2003;
Wemmie et al. 2002). In turn, transgenic mice over-expressing ASIC1a exhibit enhanced
fear conditioning (Wemmie et al. 2004). Together, these observations suggest that
ASIC1a plays a role in synaptic transmission.
Wemmie et al. performed extracellular field potential recordings of hippocampal
slices, and observed that hippocampal CA1 long-term potentiation (LTP) was impaired in
ASIC1 knockout mice (Wemmie et al. 2002). Specifically, NMDA receptor activation
during high-frequency stimulation was reduced in hippocampal CA3-CA1 synapses of
ASIC1 knockout mice. The authors hypothesized that protons released from synaptic
vesicles activate postsynaptic ASICs, which depolarize the postsynaptic membrane and facilitate activation of NMDA receptors by relieving Mg2+ block (Wemmie et al. 2002).
Depolarization-induced removal of Mg2+ block from the NMDA receptor is usually mediated by the activation of AMPA receptors. In specific situations, such as silent synapses which express few functional AMPA receptors in the postsynaptic membrane,
ASICs might induce NMDA receptor activation and thereby facilitate LTP. ASIC- mediated H+-gated current, however, has not yet been observed during high-frequency stimulation, and inhibition of ASIC channels does not appear to affect postsynaptic currents (Alvarez de la Rosa et al. 2003). These results suggest that ASICs may impact synaptic transmission through an alternate mechanism.
In this study, we examined the role of ASICs in synaptic transmission using
microisland cultures of hippocampal neurons from wild-type and ASIC knockout mice.
66 Solitary neurons in microisland culture are synaptically isolated from other neurons;
monosynaptic connections (autapses) form between the axon and dendrites of the same
neuron (Bekkers and Stevens 1991). This preparation allows a detailed analysis of
presynaptic and postsynaptic parameters of synaptic transmission in the absence of complex polysynaptic circuitry (Bekkers and Stevens 1991). Microisland cultures have
been widely used for electrophysiologic studies of synaptic transmission on a cellular and molecular level (Chavis and Westbrook 2001; Rhee et al. 2002; Rosenmund and Stevens
1996; Wierda et al. 2007). Using this method, we determined that the probability of neurotransmitter release is increased in neurons from ASIC1 knockout mice, suggesting that ASIC1a can influence glutamatergic synaptic transmission through presynaptic mechanisms.
3.2. MATERIALS AND METHODS
3.2.1. Microisland culture of hippocampal neurons
Primary hippocampal neuron cultures were prepared using previously published methods (Askwith et al. 2004; Cho and Askwith 2007). Briefly, hippocampi were dissected from postnatal day 0-1 pups, freed from extraneous tissue, and cut into pieces.
ASIC1 and ASIC2 knockout mice develop and breed normally, and there are no overt abnormalities in brain morphology (Price et al. 2000; Wemmie et al. 2002).
Hippocampal tissue was transferred into Leibovitz’s L-15 medium containing 0.25 mg/mL bovine serum albumin and 0.38 mg/mL papain, and incubated for 15 minutes at
37°C with 95% O2 and 5% CO2 gently blown over the surface of the medium. After
67 incubation, the hippocampal tissue was washed three times with mouse M5-5 medium
(Earle’s minimal essential medium with 5% fetal bovine serum, 5% horse serum, 0.4 mM
L-glutamine, 16.7 mM glucose, 5,000 U/L penicillin, 50 mg/L streptomycin, 2.5 mg/L
insulin, 16 nM selenite, and 1.4 mg/L transferrin) and triturated. For conventional mass
cultures, a collagen solution containing 0.5 mg/mL rat tail collagen in 1:1,000 acetic acid
was spread onto glass 10 mm coverslips. For microisland cultures, a microatomizer was
used to spray a fine mist of collagen solution onto the coverslips (Bekkers and Stevens
1991). The collagen was allowed to dry completely and then the coverslips were exposed
to UV light for one hour. Hippocampal cells were plated on coverslips in 24 well dishes
in M5-5 media at a density of 500,000 cells per well for mass culture and 250,000 cells per well for microisland culture. After 48-72 hours, 10 µM cytosine β-D-
arabinofuranoside was added to inhibit glial proliferation. Neurons in conventional mass
culture were used from 9 to 13 days in culture, and neurons in microisland culture were
used from 12 to 19 days in culture. Multiple breeding pairs of ASIC1 knockout (3 pairs),
ASIC2 knockout (3 pairs), and wild-type (composed of 3 pairs of wild-type siblings of
ASIC1, and 3 pairs of wild-type siblings of ASIC2 knockout) mice were used to produce
neonatal pups. No difference was observed in H+-gated currents or postsynaptic responses between wild-type neurons from the ASIC1 or ASIC2 knockout mouse lines.
3.2.2. Transfection of primary hippocampal neurons
Human ASIC1a cDNA (Genbank accession number NM_001095) was cloned
into the pcDNA3.1 expression vector (Invitrogen, Carlsbad, CA). Hippocampi were
dissected from ASIC1 knockout pups of postnatal day 0-1 as before except that
68 hippocampal cells were transfected with the ASIC1a or vector just prior to plating.
Briefly, dissociated hippocampal cells were split into two groups from the same
preparation (3-4 million cells each group), suspended in 100 µL of Nucleofector®
Solution from the Basic Nucleofector® Kit (Aamaxa Inc., Gaithersburg, MD), and mixed with 1 µg of pEGFP-C1 (Clontech, Mountain View, CA) as well as 2 µg of either the
ASIC1a construct or vector alone (pcDNA3.1). Hippocampal cells were electroporated with the NucleofectorTM II (program O-05, Amaxa Inc.), and plated on collagen-spritzed
coverslips in 24 well dishes at a density of 500,000-1,000,000 cells per well in M5-5 media. Culture medium was replaced with fresh M5-5 medium 24 hours after plating.
Cytosine β-D-arabinofuranoside (10 µM) was added 48-72 hours after plating. Single neurons on microislands were used for patch-clamping from 14 to 20 days in culture.
Transfected cells were identified by green fluorescent protein (GFP) fluorescence and selected for recording using a Nikon TE2000-S epifluorescent microscope. The whole- cell configuration was established on isolated neurons with green fluorescence, and acidic solution (pH 6.0) was applied to examine the functional expression of ASIC1a. Because the vast majority of wild-type neurons (51 out of 53 neurons) displayed H+-gated currents
in excess of 0.25 nA, we excluded ASIC1a-transfected neurons with green fluorescence
and pH 6.0-evoked response less than 0.25 nA. This method ensured that each ASIC1a-
transfected neuron included in the study expressed ASIC1a. In the ASIC1a-transfection
group, 36% of the neurons with green fluorescence (12 out of 33 neurons) displayed
typical H+-gated currents with a peak amplitude greater than 0.25 nA. In the vector-
transfection group, none of the neurons with green fluorescence displayed transient H+- gated currents. 69
3.2.3. Electrophysiology
To record H+-gated and postsynaptic currents, we used the whole-cell voltage
clamp technique. The extracellular solution contained 128 mM NaCl, 5.4 mM KCl, 2
mM CaCl2, 1 mM MgCl2, 5.55 mM glucose, 1 µM glycine, and 0.8 mM HEPES. Unless
otherwise indicated, this low concentration of pH buffer (HEPES) was used to facilitate
ASIC activation by endogenous acidic fluctuations in pH. Tetramethylammonium
hydroxide was used to adjust the pH of the extracellular solution to either pH 7.4 or pH
6.0. An extracellular pH 6.0 solution containing 10 mM HEPES and 10 mM MES as pH
buffers was used to evoke H+-gated currents. The intracellular pipette solution contained
121 mM KCl, 10 mM NaCl, 2 mM MgCl2, 5 mM EGTA, 10 mM HEPES, 2 mM Mg-
ATP, and 300 µM Na3GTP (pH 7.25). For perforated patch recording, we used the
intracellular pipette solution containing 130 mM K-gluconate, 20 mM KCl, 10 mM
HEPES, and 0.1 mM EGTA (pH 7.3). The pipette tip was filled with the intracellular solution and back-filled with the solution containing 150 µg/mL of nystatin. Patch electrodes were pulled with a P-97 micopipette puller (Sutter Instrument Co., Novato,
CA) and fire-polished with a microforge (Narishige, East Meadow, NY). Micropipettes with 2-4 MΩ were used for experiments. The membrane potential was held constant at -
70 mV. Data were collected at 5 kHz using an Axopatch 200B amplifier, Digidata
1322A and Clampex 9 (Molecular Devices, Sunnyvale, CA). Neurons were continuously superfused with the extracellular solution from gravity-fed perfusion pipes at a flow rate of approximately 1 mL per minute. Perfusion pipes were placed 250 to 300 µm away from cells, and flow was directed toward the recorded cells to ensure fast solution 70 exchange. There was no significant difference in whole-cell membrane capacitance of
neurons from the three genotypes (95 ± 10, 101 ± 17, 85 ± 10 pF, n = 24, 10, 22 for wild-
type, ASIC1 knockout, and ASIC2 knockout neurons respectively), suggesting neuron
size was not different between genotypes.
H+-gated currents were evoked by the exogenous application of pH 6.0
extracellular solutions. Desensitization time constant (τD) was calculated by fitting the
+ decay phase of H -gated currents to the single exponential equation, I(t) = Imax exp(-t/τD),
+ where Imax is the peak amplitude of H -gated current at pH 6.0. To record action potential
(AP)-evoked whole-cell postsynaptic currents, solitary autaptic neurons on microislands were selected, and stimulated every 10 seconds (0.1 Hz) with a transient depolarization of membrane potential from -70 mV to +10 mV for 2 ms. Extracellular solution containing
10 µM cyano-7-nitroquinoxaline-2,3-dione (CNQX) / 50 µM D-2-amino-5- phosphonovaleric acid (AP5) or 30 µM bicuculline was used to determine if the autaptic neuron was glutamatergic or GABAergic. To isolate α-amino-3-hydroxy-5-
methylisoxazole-4- propionate receptor (AMPAR) or N-methyl-D-aspartate receptor
(NMDAR)-mediated excitatory postsynaptic current (EPSC), we used 1 mM Mg2+ or 10
µM CNQX in Mg2+-free solution respectively. We attained similar results when the
AMPAR EPSC was isolated with 50 µM AP5. Decay time constants (τ) of AMPAR
EPSC or GABAA receptor-mediated postsynaptic current (GABAAR PSC) were calculated by fitting curves of postsynaptic currents to single exponential equation, I(t) =
Imax exp(-t/τ), where Imax is the peak amplitude of AMPAR EPSC or GABAAR PSC. To quantify the decay rate of NMDAR EPSCs, we calculated weighted mean decay time constant of NMDAR EPSC. NMDAR EPSC curves were fitted to the double exponential 71 equations, I(t) = If exp(-t/τf) + Is exp(-t/τs), where If /τf and Is /τs are fast and slow components of peak amplitudes and decay time constants of NMDAR EPSC, and then weighted mean decay time constants (τw) were calculated from the equation, τw= τf (If
/(If + Is)) + τs (Is /(If + Is)). Miniature EPSC (mEPSC) and hypertonic (500 mM) sucrose- induced currents were recorded from neurons in conventional mass culture using extracellular solution containing 1 mM Mg2+, 30 µM bicuculline, and 1 µΜ tetrodotoxin.
Unless otherwise indicated, all reagents were purchased from Invitrogen/Gibco
(Carlsbad, CA), Sigma-Aldrich (St. Louis, MO), or Fisher Scientific (Waltham, MA).
Data were analyzed using Clampfit 9 software (Molecular Devices). We analyzed mEPSC using the template search function of Clampfit 9. Data are presented as the mean
± the standard error of the mean (SEM). As appropriate, a two-tailed Student’s t test and one-way ANOVA with Bonferroni’s simultaneous multiple comparisons were used for statistical analyses and performed with Minitab15 software (Minitab Inc, State College,
PA).
3.3. RESULTS
3.3.1. H+-gated currents are altered in ASIC knockout neurons cultured on
microislands
In conventional mass-cultured hippocampal neurons, ASIC1a homomultimers and
ASIC1a/ASIC2a heteromultimers are responsible for the majority of H+-gated currents
(Askwith et al. 2004). The ASIC1a subunit plays a dominant role, and loss of ASIC1a
eliminates H+-gated currents evoked by pHs above 5 (Wemmie et al. 2002; Xiong et al. 72 2006). The ASIC2a subunit modulates H+-gated current by forming heteromultimeric channels with ASIC1a that desensitize more rapidly and recover from previous acid applications more quickly compared to homomultimeric ASIC1a channels (Askwith et al.
2004; Bassilana et al. 1997; Benson et al. 2002). To determine if ASIC1 and ASIC2 have similar roles in microisland culture, we analyzed H+-gated currents of solitary
hippocampal neurons from wild-type, ASIC1 knockout, and ASIC2 knockout mice.
Application of acidic extracellular solution (pH 6.0) did not induce a substantial transient
current in ASIC1 knockout neurons (Fig. 1A-B, peak amplitude = 39 ± 17 pA, n = 13, p
< 0.01, wild-type versus ASIC1 knockout, one-way ANOVA). By comparison,
extracellular application of pH 6.0 solution induced transient inward currents in both
wild-type and ASIC2 knockout neurons (Fig. 1A-B, peak amplitude = 796 ± 106 pA, n =
53 for wild-type, and 1595 ± 217 pA, n = 31 for ASIC2 knockout). The peak amplitude
+ and desensitization time constant (τD) of H -gated currents were significantly larger in
ASIC2 knockout neurons compared to wild-type neurons (Fig. 1B-C, p < 0.01 for peak
amplitude, one-way ANOVA; p < 0.05 for desensitization time constants, unpaired t
test). Recovery from desensitization was also substantially slower in ASIC2 knockout
neurons compared to wild-type neurons (Fig. 1A and D, 45 ± 3%, n = 37 for wild-type,
17 ± 3%, n = 31 for ASIC2 knockout, p < 0.0001, unpaired t test). These results are
consistent with the loss of ASIC1a/ASIC2a heteromeric channels, which desensitize and
recover faster, in the ASIC2 knockout neurons (Askwith et al. 2004; Bassilana et al.
1997; Benson et al. 2002). These results indicate that ASIC subunits play similar roles in
microisland culture and conventional mass culture (Askwith et al. 2004).
73 Whether a solitary neuron in microisland culture is glutamatergic or GABAergic
can be determined by analyzing the action potential-evoked postsynaptic current.
Extracellular solutions containing CNQX/AP5 or bicuculline were used to identify the
autaptic neuron as glutamatergic or GABAergic respectively. Both glutamatergic and
GABAergic autaptic neurons expressed H+-gated currents in wild-type and ASIC2 knockout neurons. The current density (peak current amplitude divided by whole-cell membrane capacitance) of H+-gated currents was significantly larger in GABAergic neurons than in glutamatergic neurons (Fig. 1E, 13.7 ± 2.2 pA/pF, n = 24 for glutamatergic neurons, and 22.3 ± 2.7 pA/pF, n = 18 for GABAergic neurons, p < 0.05, unpaired t test). The desensitization time constant and recovery from desensitization of
H+-gated currents were not different between these two populations of neurons (data not shown). Together, these data indicate that both glutamatergic and GABAergic neurons in microisland culture express H+-gated channels with similar biophysical characteristics to mass-cultured neurons.
3.3.2. Whole-cell postsynaptic currents in solitary neurons in microisland culture
To gain insight into the role of ASICs in synaptic transmission, action potential
(AP)-evoked postsynaptic currents of solitary neurons in microisland culture were analyzed using the whole-cell voltage-clamp technique. In GABAergic autaptic neurons there was no significant difference in the peak amplitude or decay time constant of
GABAA receptor-mediated postsynaptic current between wild-type and ASIC knockout
neurons (Fig. 2A-B, peak amplitude = 4.19 ± 0.43, 3.48 ± 0.34, 4.52 ± 0.56 nA, n = 61,
48, 32 for wild-type, ASIC1 knockout, and ASIC2 knockout respectively, p = 0.27; decay
74 time constant = 119 ± 24, 132 ± 27, 115 ± 22 ms, n = 12, 14, 13 for wild-type, ASIC1
knockout, and ASIC2 knockout respectively, p = 0.87, one-way ANOVA). In
glutamatergic autaptic neurons, AMPA receptor and NMDA receptor-mediated
excitatory postsynaptic currents (AMPAR EPSC and NMDAR EPSC) were individually
isolated using either 1 mM Mg2+ or 10 µM CNQX respectively (Fig. 2C). Although there was a slight increase in the average peak amplitudes in ASIC1 knockout neurons, there was no significant difference between wild-type and ASIC knockout neurons in the peak amplitudes of either the AMPAR EPSC or NMDAR EPSC (Fig. 2D-E, peak amplitude of
AMPAR EPSC = 3.65 ± 0.50, 4.29 ± 0.58, 3.87 ± 0.60 nA, n = 40, 42, 28 for wild-type,
ASIC1 knockout, and ASIC2 knockout respectively, p = 0.69; peak amplitude of
NMDAR EPSC = 1.10 ± 0.17, 1.58 ± 0.26, 1.08 ± 0.18 nA for wild-type, ASIC1 knockout, and ASIC2 knockout respectively, p = 0.17, one-way ANOVA). There was also no significant difference in the decay time constant of AMPAR or NMDAR EPSC
(decay time constant of AMPAR EPSC = 11.4 ± 0.8, 13.3 ± 1.3, 10.6 ± 0.9 ms, n = 36,
40, 27 for wild-type, ASIC1 knockout, and ASIC2 knockout respectively, p = 0.18; weighted mean decay time constant of NMDAR EPSC = 198 ± 14, 182 ± 9, 155 ± 11 ms, n = 45, 48, 28 for wild-type, ASIC1 knockout, and ASIC2 knockout respectively, p =
0.07, one-way ANOVA). To compare the relative contribution of AMPA receptors and
NMDA receptors to the total EPSC, we recorded both AMPAR and NMDAR EPSCs in the same neurons, and calculated the ratio of AMPAR EPSC to NMDAR EPSC. The
AMPAR / NMDAR EPSC ratio was significantly smaller in ASIC1 knockout neurons compared to wild-type and ASIC2 knockout neurons (Fig. 2C and F, AMPAR EPSC /
NMDAR EPSC ratio = 4.41 ± 0.33, 2.82 ± 0.15, 4.52 ± 0.57, n = 40, 42, 28 for wild-type,
75 ASIC1 knockout, and ASIC2 knockout respectively, p < 0.05, ASIC1 knockout versus wild-type or ASIC2 knockout, one-way ANOVA). The fact that ASIC1 knockout
neurons displayed a decrease in the AMPAR / NMDAR EPSC ratio could be due to a
relative decrease in the peak amplitude of AMPAR EPSC or a relative increase in peak
amplitude of the NMDAR EPSC in ASIC1 knockout neurons. Although variability in
AMPAR and NMDAR EPSCs between neurons prevented the detection of a statistically
significant change in the peak amplitude of AMPAR or NMDAR EPSC, this result
suggests that there is a fundamental difference in glutamatergic synaptic transmission in
ASIC1 knockout neurons.
3.3.3. ASIC currents are not components of EPSCs
If ASIC currents contribute directly to the postsynaptic current, then loss of H+- gated currents could explain the altered AMPAR / NMDAR EPSC ratio in the ASIC1 knockout neurons. ASIC1a is enriched in synaptosomal fractions and expressed in dendritic spines suggesting ASICs play a role at postsynaptic sites (Hruska-Hageman et al. 2002; Wemmie et al. 2002; Wemmie et al. 2004). The pH inside synaptic vesicles is acidic (< pH 6.0) (Miesenbock et al. 1998), and protons are released from synaptic vesicles during synaptic transmission. Under certain circumstances, such as high frequency synaptic transmission, these released protons can lower the pH of the synaptic cleft (Krishtal et al. 1987) and affect pH-sensitive ion channel activity. This has been shown for voltage-gated Ca2+ channels in retinal cone photoreceptor and bipolar cells
(DeVries 2001; Hosoi et al. 2005; Palmer et al. 2003; Vessey et al. 2005). ASIC
activation evoked by proton released from synaptic vesicles has not been demonstrated,
76 although spontaneous autocrine release of protons activates endogenous ASIC currents in
HEK293 cells (Lalo et al. 2007). Protons released from synaptic vesicles may be rapidly
trapped by endogenous pH buffers, and the acute activation of ASICs by protons would
likely be for only a short time (Krishtal et al. 1987). Thus, protons released during
synaptic transmission may activate ASIC1a-containing channels generating postsynaptic
currents in wild-type neurons, which could contribute to the early component of EPSC
(AMPAR EPSC). ASIC1 knockout neurons would lack this H+-dependent current, and thus display a smaller relative AMPAR / NMDAR EPSC ratio. To test this hypothesis, we examined whether ASICs directly contribute to postsynaptic currents.
First, we determined whether ASIC current could be isolated during synaptic transmission. Both AMPAR and NMDAR EPSCs were simultaneously inhibited using
10 µM CNQX and 50 µM AP5 in wild-type glutamatergic neurons on microislands.
When both AMPA and NMDA receptors were blocked, a small transient residual current was observed in glutamatergic neurons that contributed to the peak AMPAR EPSC (Fig.
3A). This current was not significantly different between wild-type and ASIC1 knockout neurons (128 ± 31 pA, n = 7 for wild-type neurons; 124 ± 50 pA, n = 7 for ASIC1 knockout neurons, p = 0.96, unpaired t test). We also recorded both the AMPAR EPSC and the residual current in the same neuron, and calculated the ratio of the residual current amplitude to the peak amplitude of the AMPAR EPSC. The ratio was not different between wild-type and ASIC1 knockout neurons (Fig. 3B, 4.06 ± 0.77%, n = 7 for wild-type neurons; 5.18 ± 0.96%, n = 7 for ASIC1 knockout neurons, p = 0.38, unpaired t test), suggesting that the contribution of the residual current to peak AMPAR
EPSC was not affected by the loss of ASIC1.
77 We also examined the effect of amiloride (300 µM), a non-specific ASIC blocker,
on residual current and AMPAR EPSCs (Fig. 3C and E). H+-gated currents evoked by exogenous acid application were blocked nearly completely by 300 µM amiloride (Fig.
3C). However, the contribution of the residual current to AMPAR EPSCs in wild-type
glutamatergic neurons was not affected by amiloride (Fig. 3C-D, 3.62 ± 0.80% and 3.30
± 0.98% before and during amiloride application respectively, n = 5, p = 0.38, paired t
test). Furthermore, AMPAR EPSCs were not altered by amiloride in either wild-type or
ASIC1 knockout neurons (Fig. 3E-F, 92.1 ± 4.1% of pretreatment control, n = 8, p =
0.10, paired t test for wild-type; 99.7 ± 4.6%, n = 8, p = 0.35, paired t test for ASIC1 knockout; p = 0.23, wild-type versus ASIC1 knockout, unpaired t test). Together, these results indicate that ASIC currents are not components of the EPSC, and the altered
AMPAR / NMDAR EPSC ratio in ASIC1 knockout neurons cannot be explained by loss of postsynaptic ASIC currents evoked during synaptic transmission. Thus, another mechanism is responsible for the decrease in AMPAR / NMDAR EPSC ratio in the
ASIC1 knockout neurons.
3.3.4. Exogenous acid application did not alter whole-cell postsynaptic currents.
Activation of ASIC1a channels could induce changes in AMPAR, NMDAR, or
GABAAR-mediated postsynaptic currents through signal transduction cascades. To test
this, we measured whole-cell postsynaptic currents before and after activating ASICs
with exogenously applied acid. Postsynaptic currents were recorded every 10 seconds in
wild-type autaptic neurons. After the peak amplitude of baseline postsynaptic currents
stabilized, we briefly evoked H+-gated currents with the application of pH 6.0 78 extracellular solutions for 5 seconds. We then returned the pH to 7.4 and continued to
measure postsynaptic currents every 10 seconds for several minutes (Fig. 4). Peak
amplitudes of AMPAR, NMDAR, and GABAAR postsynaptic currents were unchanged
following acid application (Fig. 4A-C, n = 4-5). AMPAR EPSCs were increased robustly
when we applied 2 µM phorbol 12,13-dibutyrate (PDBu), a diacylglycerol analog that
potentiates synaptic transmission through protein kinase C and other mechanisms (Fig.
4D, n = 6) (Wierda et al. 2007). This indicates that our experimental configuration could
support signal transduction dependent changes in synaptic transmission. To further
conserve intracellular signaling mechanisms, we performed the same experiment using
nystatin-based perforated patch recording. Again, we did not observe changes in
postsynaptic current amplitudes following acid application (n = 4-5, data not shown).
These results suggest that acute activation of ASICs does not rapidly alter postsynaptic currents.
3.3.5. Paired-pulse ratio is reduced in ASIC1 knockout neurons
Our results suggest that the decrease in the AMPAR / NMDAR EPSC ratio in
ASIC1 knockout neurons is not due to activation of postsynaptic ASIC currents during the recording interval. To determine if other aspects of synaptic transmission are altered
in ASIC knockout neurons, we assessed the synaptic response to repetitive and paired-
pulse stimulation (Mennerick and Zorumski 1995; Zucker and Regehr 2002). A pair of
stimuli with short intervals from 20 ms to 400 ms was applied, and the paired-pulse ratio
(PPR) was determined by dividing the peak amplitude of the second AMPAR EPSC by
the peak amplitude of the first EPSC (Fig. 5A). Compared to wild-type neurons, the
79 PPRs with interpulse intervals of 50, 100, and 200 ms were significantly reduced in
ASIC1 knockout neurons (Fig. 5A-B, p < 0.05, wild-type versus ASIC1 knockout, one- way ANOVA). Furthermore, during a short train of repetitive stimuli with 50 ms intervals, the amplitudes of the second through the fifth AMPAR EPSC normalized to the first EPSC amplitude were reduced in ASIC1 knockout neurons compared to wild-type neurons (Fig. 5C-D).
The fact that the PPR was altered was particularly surprising. Previous studies analyzing extracellular field potentials of Schaffer collateral-CA1 synapses in hippocampal slices did not observe changes in paired-pulse facilitation between ASIC1 knockout and wild-type mice. In contrast, we found that the PPR of ASIC1 knockout neurons in microisland cultures was reduced. The reason for this discrepancy is unclear.
However, more variables such as polysynaptic activity contribute to the PPR in hippocampal slices compared to the PPR in microisland cultures (Mennerick and
Zorumski 1995; Zucker and Regehr 2002), and the simplicity and enhanced experimental control over these variables in the microisland culture system may have facilitated detection of alterations in PPR. However, we also tested whether our pH buffer conditions may have allowed observation altered PPR. Our experiments utilize a lower
HEPES concentration (0.8 mM) compared to others (10 mM) to facilitate ASIC activation by endogenous acidic fluctuations in pH. To determine if the difference in
PPR was due to the low concentration of pH buffer used in our studies, we analyzed postsynaptic currents and PPR when the external buffer was increased to 10 mM HEPES, a concentration of HEPES that supports a buffering capacity close to that of
- - CO2/bicarbonate and phosphate buffers in vivo (24 mM HCO3 and 1 mM H2PO4 under
80 5% CO2). We found that increasing the concentration of HEPES from 0.8 mM to 10 mM
slightly reduced the peak amplitude of AMPAR EPSCs, NMDAR EPSCs, and GABAAR
PSCs. However, this effect was not different between wild-type, ASIC1 knockout, and
ASIC2 knockout neurons (Fig. 6A-B, n = 6-24, p = 0.48, 0.83, and 0.60 for AMPAR
EPSC, NMDAR EPSC, and GABAAR PSC respectively, one-way ANOVA).
Furthermore, PPR with 50 ms interval was not altered significantly by increasing HEPES
concentration from 0.8 mM to 10 mM in wild-type neurons (Fig. 6C-D, n = 15, p = 0.81,
paired t test). This indicates that the difference in the PPR between wild-type and ASIC1
knockout neurons is not due to the low concentration of pH buffer (0.8 mM HEPES).
We also analyzed whether the reduction in the PPR was due to the loss of ASIC currents within the second EPSC. To test this, we measured residual currents in the presence of CNQX and AP5 in a paired-pulse protocol with a 50 ms interval (Fig. 6E).
We did not observe a significant difference in the relative contribution of residual current to peak amplitude of AMPAR EPSC between the first and the second stimulation in either wild-type or ASIC1 knockout neurons (Fig. 6F, residual current after the first and the second stimulation = 4.7 ± 0.8% and 4.7 ± 0.6% of peak AMPAR EPSC, n = 5, p =
0.94 for wild-type; 5.5 ± 5.8% and 5.8 ± 0.78%, n = 4, p = 0.73 for ASIC1 knockout, paired t test). Furthermore, inhibition of ASICs with amiloride did not affect the PPR with a 50 ms interval in either wild-type or ASIC1 knockout neurons (Fig. 6G-H, 102 ±
2.6% of pre-amiloride control, n = 4, p = 0.51 for wild-type; 95 ± 2.7%, n = 4, p = 0.17
for ASIC1 knockout, paired t test). These results suggest that the difference in PPR
between wild-type and ASIC1 knockout neurons is not due to the direct contribution of
ASIC current to AMPAR EPSC.
81
3.3.6. The frequency of spontaneous neurotransmitter release is increased in ASIC1
knockout neurons
Alterations in the paired pulse ratio can be due to multiple factors. To further
define the synaptic alterations in ASIC1 knockout neurons, we performed quantal
analyses on spontaneous miniature EPSC (mEPSC) of wild-type and ASIC1 knockout
neurons in conventional mass culture (Mennerick et al. 1995). We recorded AMPA
receptor-mediated mEPSC in extracellular solution containing 1 mM Mg2+, 30 µM
bicuculline, and 1 µM tetrodotoxin to prevent action potential firing (Fig. 7A). The average peak amplitude of mEPSC was not different between wild-type and ASIC1 knockout neurons (Fig. 7B, 19.8 ± 2.2 pA, n = 11 neurons for wild-type, 21.1 ± 2.5 pA, n
= 8 neurons for ASIC1 knockout, p = 0.70, unpaired t test). In addition, neither the decay time constant nor the charge transfer of mEPSCs was different between wild-type and
ASIC1 knockout neurons (decay time constant of mEPSC = 6.9 ± 0.1 and 7.1 ± 0.1 ms, for wild-type and ASIC1 knockout, p = 0.10, unpaired t test; charge transfer of mEPSC =
102 ± 11 and 105 ± 12 fC for wild-type and ASIC1 knockout, p = 0.86, unpaired t test).
These results indicate that the postsynaptic response to fusion of a single synaptic vesicle is not different in ASIC1 knockout neurons. However, mEPSCs were more frequent in
ASIC1 knockout neurons than in wild-type neurons (Fig. 7C, 2.3 ± 0.2 Hz for wild-type;
5.1 ± 1.1 Hz for ASIC1 knockout, p < 0.05, unpaired t test). An increase in mEPSC frequency is commonly caused by an increase in the total number of synapses, an increase in the size of the readily-releasable pool of synaptic vesicles, or an increase in the probability of neurotransmitter release. Previous studies have determined that the 82 density of dendritic spines is equivalent in ASIC1 knockout and wild-type neurons in
hippocampal slices (Zha et al. 2006). This suggests that the increased frequency of
mEPSCs may be due to a larger number of synaptic vesicles in the readily-releasable pool
or a higher probability of neurotransmitter release in ASIC1 knockout neurons.
3.3.7. Readily-releasable pool size and refilling are not different in ASIC1 knockout
neurons
The readily-releasable pool (RRP) size and refilling were analyzed using
hypertonic sucrose solution. When hypertonic sucrose is applied, synaptic vesicles in the entire RRP release neurotransmitters in a Ca2+-independent manner and produce postsynaptic currents proportional to the size of the RRP (Rosenmund and Stevens 1996).
Extracellular solution containing 500 mM sucrose was applied to neurons in mass culture for 4-5 seconds and the sucrose-induced postsynaptic current response was recorded in 1 mM Mg2+, 30 µM bicuculline, and 1 µM tetrodotoxin (Fig. 8A). This sucrose-induced response was completely inhibited by 10 µM CNQX (Fig. 8A) indicating that it was mediated by AMPA receptors (Rosenmund and Stevens 1996). RRP size was estimated from the sucrose-induced charge transfer calculated by integrating the transient component of the sucrose-evoked current. We determined that the RRP size was not significantly different between wild-type and ASIC1 knockout neurons (Fig. 8B, 0.86 ±
0.09 nC, n = 20 for wild-type, 0.95 ± 0.11 nC, n = 24 for ASIC1 knockout, p = 0.52, unpaired t test). The RRP refilling was also analyzed by comparing the charge transfer in response to a second sucrose application 3-4 seconds after the first (Fig. 8A) (Priller et al.
2006). RRP refilling, estimated as the ratio of the charge transfer by the second sucrose 83 application to the charge transfer induced by the first sucrose application, was not
significantly different between wild-type and ASIC1 knockout neurons (Fig. 8C, 0.53 ±
0.02, n = 17 for wild-type, 0.52 ± 0.03, n = 20 for ASIC1 knockout, p = 0.93, unpaired t test). These results indicate that the size and refilling of the RRP are not different between wild-type and ASIC1 knockout neurons in conventional mass culture.
3.3.8. The progressive block of NMDA receptors by MK-801 was faster in ASIC1
knockout neurons
An increase in the release probability could account for both the more frequent
mEPSCs and reduced PPR observed in ASIC1 knockout neurons (Zucker and Regehr
2002). The release probability of ASIC1 knockout neurons was analyzed using
progressive block of the NMDA receptors (NMDAR) by MK-801 (Rosenmund et al.
1993). Because MK-801 blocks NMDAR irreversibly while they are open, the rate of
NMDAR EPSC decrease correlates with the probability of neurotransmitter release (Futai
et al. 2007; Rosenmund et al. 1993). Thus, the higher the release probability is, the faster
the rate of MK801-induced decrease of NMDAR EPSCs. The rate of progressive block
of NMDAR by MK801 could also be affected by the duration of NMDAR opening.
Because NR2B-containing NMDARs open longer than NR2A-containing NMDARs, a
difference in the ratio of NR2B to NR2A-containing NMDARs can hamper interpretation
of NMDAR progressive block by MK-801 (Monyer et al. 1992). However, we find that
the weighted mean decay time constant of NMDAR EPSCs was not different between
wild-type and ASIC1 knockout neurons (198 ± 14, n = 45 for wild-type, and 182 ± 9, n =
84 48 for ASIC1 knockout, p = 0.37, unpaired t test), suggesting that NMDAR remain open
for similar periods of time in response to glutamate.
Under normal conditions, the NMDAR EPSCs of autaptic neurons stimulated
every 10 seconds did not change significantly over time (Fig. 9A). In the presence of 10
µM MK-801, the NMDAR EPSCs gradually decreased in a stimulus-dependent manner
(Fig. 9A). NMDAR EPSC amplitudes were normalized to the first NMDAR EPSC in the
presence of MK-801 and plotted against stimulus number. The data were fit to a single
exponential equation, and the rate of the decrease of the NMDAR EPSCs was quantified
by calculating the tau (τ) in stimulus number (Fig. 9B-D). The rate of NMDAR EPSC
decrease in the presence of MK-801 was faster, and the tau was significantly reduced in
ASIC1 knockout neurons compared to wild-type neurons (Fig. 9C-D, τ = 22.4 ± 1.9 stimuli n = 9 for wild-type, 16.1 ± 1.5 stimuli, n = 8 for ASIC1 knockout, p < 0.05, unpaired t test). These results suggest that the release probability is higher in ASIC1 knockout glutamatergic neurons.
3.3.9. Neurotransmitter release probability is increased in ASIC1 knockout neurons
A second experiment was used to assess the release probability of wild-type and
ASIC knockout neurons. Both the action potential (AP)-evoked EPSC and hypertonic sucrose-induced current were recorded in the same glutamatergic autaptic neurons (Fig.
10A). The probability of neurotransmitter release was calculated by dividing the charge transfer of AP-evoked EPSC by the charge transfer of the response following depletion of the readily-releasable pool of synaptic vesicles with hypertonic sucrose (Moulder et al.
2004). The release probability of wild-type and ASIC2 knockout neurons was 6-7% (Fig. 85 10D, 7.0 ± 0.8%, n = 27 for wild-type, and 6.2 ± 0.5%, n = 12 for ASIC2 knockout), which is close to the value obtained in other studies (Rhee et al. 2002). The release probability of ASIC1 knockout neurons was significantly higher compared to wild-type and ASIC2 knockout neurons (Fig. 10D, 10.9 ± 1.7%, n = 19 for ASIC1 knockout, p <
0.05, ASIC1 knockout versus wild-type or ASIC2 knockout, one-way ANOVA).
Together, this experiment and the experiment assessing progressive block of NMDAR by
MK-801 (Fig. 9) indicate that ASIC1 knockout neurons have a higher probability of neurotransmitter release than wild-type neurons.
3.3.10. Rescue of ASIC1 knockout neurons by ASIC1a expression
We find that the release probability is higher in neurons from ASIC1 knockout mice compared to neurons from wild-type mice. To determine if this effect is the result of developmental compensation in response to ASIC1 gene disruption, we performed rescue experiments in cultured neurons. Hippocampal neurons from ASIC1 knockout mice were transfected during isolation with either vector alone or vector expressing
ASIC1a. Neurons were plated to foster microisland conditions, and whole-cell patch clamping was used to measure acid-evoked currents and glutamatergic synaptic transmission after 14-20 days in culture. We found that extracellular acid (pH 6.0) failed to evoke H+-gated currents in ASIC1 knockout neurons transfected with vector alone
(Fig. 11A, peak amplitude = 74 ± 17 pA, n = 9). In ASIC1a-transfected ASIC1 knockout
neurons, extracellular acid induced typical H+-gated currents with characteristics similar
to wild-type neurons (Fig. 11A, peak amplitude = 1489 ± 458 pA, n = 6, p < 0.05,
ASIC1a-transfected versus vector-transfected neurons, unpaired t test ). Thus,
86 transfection of ASIC1 knockout neurons with ASIC1a restored H+-gated currents. We next examined synaptic transmission in transfected ASIC1 knockout. GABAA receptor- mediated postsynaptic currents were not significantly different in ASIC1 knockout neurons transfected with ASIC1a or vector alone (peak amplitude = 5.5 ± 1.3 nA, n = 6 for ASIC1a-transfected neurons, and 3.5 ± 0.8 nA, n = 8 for vector-transfected neurons, p
= 0.23, unpaired t test). However, the peak amplitude of AMPA receptor-mediated
EPSCs was profoundly smaller in ASIC1a-transfected neurons compared to vector- transfected neurons (Fig. 11B, 1.60 ± 0.27 nA, n = 6 for ASIC1a-transfected neurons, and
4.42 ± 0.66 nA, n = 9 for vector-transfected neurons, p < 0.01, unpaired t test).
Furthermore, the paired-pulse ratios at interpulse intervals of 40 and 50 ms were increased in ASIC1a-transfected neurons (Fig. 11C, p < 0.05, unpaired t test). Because the paired-pulse ratio is inversely correlated with release probability (Zucker and Regehr
2002), these results suggest that release probability of ASIC1 knockout neurons transfected with ASIC1a is decreased. Further, we suggest that the decreased release probability also resulted in smaller amplitude of AMPAR EPSCs in ASIC1a-transfected
ASIC1 knockout neurons. These experiments show that the paired-pulse ratio of ASIC1 knockout neurons can be restored by in vitro expression of ASIC1a, and suggest that the increase in release probability observed in ASIC1 knockout neurons is not due to developmental compensation.
87 3.4. DISCUSSION
Given that ASICs are activated by extracellular protons, a role for ASICs in
biological processes involving extracellular acidosis is not surprising. For example,
ASICs mediate acid-induced nociception during inflammation (Price et al. 2001;
Sutherland et al. 2001). ASICs are involved in the retinal response to light where acidic
pH changes are known to impact neuronal signaling (Ettaiche et al. 2006; Ettaiche et al.
2004). In the brain, ASIC1a activation causes neuronal death during prolonged acidosis
following ischemia (Xiong et al. 2004). However, ASICs are also involved in neuronal
processes where the role of extracellular acidosis is not well defined. For example,
ASIC1 is required for normal fear-related behaviors as well as learning and memory
(Wemmie et al. 2003; Wemmie et al. 2002; Wemmie et al. 2004). Although rapid
extracellular pH transients have been observed during synaptic transmission (Krishtal et
al. 1987), how ASICs impact neurotransmission is not clear.
In this study, we investigated the role of ASICs in synaptic transmission by
comparing postsynaptic currents of cultured hippocampal neurons from wild-type, ASIC1
and ASIC2 knockout mice. We observed alterations in synaptic transmission in
glutamatergic neurons from ASIC1 knockout animals using multiple experimental
paradigms. First, the AMPAR / NMDAR EPSC ratio was reduced in ASIC1 knockout neurons (Fig. 2F). Second, the paired-pulse ratio (PPR) of AMPAR EPSC was reduced and the depression of AMPAR EPSCs during a short train of stimuli was greater in
ASIC1 knockout neurons (Fig. 5). Alterations in the PPR can be due to both presynaptic and postsynaptic mechanisms. In an effort to identify the nature of the PPR changes in
88 ASIC1 knockout neurons, we assessed other aspects of synaptic transmission. We determined that the quantal size of mEPSCs, the size of readily-releasable pool (RRP) of
synaptic vesicles, and RRP refilling were not significantly different in ASIC1 knockout
neurons compared to wild-type neurons (Fig. 7-8). In contrast, the frequency of mEPSCs
was increased (Fig. 7C), and the progressive block of NMDAR by MK-801 was faster in
ASIC1 knockout neurons (Fig. 9). These results suggest that the reduced PPR in ASIC1
knockout neurons is due to the increased probability of neurotransmitter release. This
was confirmed by the observation that the release probability, as measured by the ratio of
AP-evoked to hypertonic sucrose-evoked charge transfer, was increased in ASIC1
knockout neurons (Fig. 10). Together, these results support the conclusion that cultured
ASIC1 knockout neurons have a higher release probability compared to wild-type
neurons. Further, the fact that mEPSC frequency increased and the release probability
was augmented without a change in the readily-reliable pool size suggests an enhanced
fusion propensity of individual vesicles in ASIC1 knockout neurons. Transfection of
ASIC1a increased the PPR of ASIC1 knockout neurons and reduced AMPAR EPSCs
(Fig. 11). These results are consistent with an ASIC1a-dependent rescue of the release
probability of ASIC1 knockout neurons, and suggest that the increased release probability
observed in ASIC1 knockout neurons is not likely due to developmental compensation.
Interestingly, there were no significant differences in synaptic transmission in ASIC2
knockout neurons (Fig. 2F, 5B, and 10D), indicating that these alterations are specific to
disruption of ASIC1.
As the release probability is increased in ASIC1 knockout neurons, AP-evoked
whole-cell postsynaptic currents should be larger compared to wild-type neurons.
89 Although the average peak amplitudes of single AP-evoked AMPAR EPSCs or NMDAR
EPSCs were larger in ASIC1 knockout neurons (Fig. 2D-E, and Fig. 10B), the difference
did not reach statistical significance, and we expect a larger increase in EPSC amplitude
with such an increase in release probability. We suggest that the postsynaptic response of
ASIC1 knockout neurons is altered such that the EPSC amplitude is unchanged even with
the increased release probability. In support of this idea, we did observe that AMPAR /
NMDAR EPSC ratio was reduced in ASIC1 knockout neurons (Fig. 2F). Because the
quantal size of AMPAR-mediated mEPSC was not changed in ASIC1 knockout neurons
(Fig. 7B), it is not likely that the number of AMPAR per synapse or the conductance of
individual postsynaptic AMPAR was different in the ASIC1 knockout neurons. In
contrast, disruption of ASIC1 may have affected the ratio of AMPAR-expressing
functional synapses to silent synapses which contain NMDAR but not AMPAR. Thus,
the number of AMPAR-expressing functional synapses may be reduced in ASIC1
knockout neurons. Such a reduction would mask the effect of enhanced release
probability on AMPAR EPSCs. In rescue experiments, we did observe alterations in
postsynaptic current consistent with altered release probability. ASIC1 knockout neurons
transfected with ASIC1a displayed dramatically reduced AMPAR EPSCs compared to
vector-transfected neurons consistent with a decrease in release probability (Fig. 11B).
This result suggests that presynaptic alterations in ASIC1 knockout neurons can be
rescued by expression of ASIC1a in culture.
It is hypothesized that postsynaptic ASICs are activated by protons released from
synaptic vesicles during neurotransmission. ASIC activation, in turn, contributes to depolarization and calcium-induced signaling cascades. This model has been difficult to
90 prove and, like others before, we did not detect ASIC-mediated postsynaptic currents
during synaptic transmission (Fig. 3A-D). Even multiple stimulations failed to induce
measurable ASIC-dependent currents (Fig. 6E-F). Furthermore, we did not observe any
ASIC-dependent effect of amiloride on AMPAR EPSC or PPR (Fig. 3E-F, and 6G-H).
Given the large postsynaptic currents recorded, the sensitivity of the method, and the
presence of H+-gated currents evoked by exogenous acid application in these neurons, we
conclude that specific conditions must exist for ASICs to be activated in this manner and
make a substantial contribution to the postsynaptic currents. However, definitive changes
in synaptic transmission were observed in ASIC1 knockout neurons even though ASIC
currents were not detected during synaptic transmission. Our data suggests that the
higher release probability in ASIC1 knockout neurons is due to either (1) disruption of
ASIC1a-specific signaling which has long-term effects on presynaptic mechanisms, or
(2) loss of an unconventional non-ionotropic ASIC function which is not dependent on ion conduction of ASIC1a.
First, disruption of ASIC1a-specific signaling could have long-term effects on presynaptic mechanisms. Under normal conditions, ASIC1a activation could lead to long-term changes on synaptic transmission through activation of signal transduction cascades. ASIC1a homomultimers are Ca2+-permeable (Yermolaieva et al. 2004), and may activate Ca2+/calmodulin-dependent protein kinase II (CaMKII). Phosphorylated
CaMKIIα is reduced in brains from ASIC1 knockout mice (Zha et al. 2006) indicating that disruption of ASIC1 can impact activation status of CaMKII, which is essential for the recruitment of AMPAR to synapses and for the LTP induction in the CA1 region of hippocampus (Lisman et al. 2002). CaMKII may also affect presynaptic mechanisms to
91 alter neurotransmitter release (Chi et al. 2001; Llinas et al. 1991; Sanhueza et al. 2007;
Waxham et al. 1993). To investigate the possibility that previous activation of ASIC1a
causes changes in synaptic transmission, we assessed postsynaptic current following
activation of ASICs by exogenous acid application. However, we did not observe
changes in postsynaptic currents in the time frame assessed (2-5 minutes, Fig. 4). Thus,
ASIC1a-induced changes in signal transduction may occur under other specific
conditions or required a longer period of time to manifest.
Second, changes in synaptic transmission of ASIC1 knockout neurons could be
due to the loss of an unconventional non-ionotropic ASIC function. Such ‘non-
conducting’ functions have been reported for other ion channels such as NMDA receptors
(Alvarez et al. 2007) and HCN (Ih) channels (Beaumont et al. 2002; Beaumont and
Zucker 2000; Zhong et al. 2004). ASIC1a could regulate release probability by direct interaction with neurotransmitter release machinery. Previous studies indicate that
ASICs are localized postsynaptically (Hruska-Hageman et al. 2002; Wemmie et al.
2002), but do not exclude the possibility of ASIC1a at presynaptic sites (Alvarez de la
Rosa et al. 2003). Ionotropic neurotransmitter receptors such as NMDA receptors, kainite receptor, and GABAA receptor are expressed presynaptically as well as
postsynaptically and regulate neurotransmitter release (MacDermott et al. 1999).
Furthermore, ASIC1a, ASIC2a, and γ-ENaC form heteromeric channels in malignant
glioma cells, and this complex interacts with syntaxin 1A, a component of SNARE
complex involved in exocytosis of synaptic vesicles (Berdiev et al. 2003). Although it is
not known whether this interaction also occurs in neurons, this raises a possibility that
ASICs might regulate neurotransmitter release directly. Alternatively, postsynaptic
92 ASIC1a could affect synaptic transmission in a retrograde manner like postsynaptic PSD-
95, which regulates neurotransmitter release by physical interaction with presynaptic
neurexin (Futai et al. 2007). ASICs have a large extracellular loop of largely unknown
function, and the extracellular loop of postsynaptic ASIC1a could physically interact with
a presynaptic counterpart to regulate neurotransmitter release directly.
ASIC1 knockout mice exhibited defects in multiple aspects of learning and
memory such as spatial learning, fear conditioning, and eye-blink conditioning (Wemmie
et al. 2003; Wemmie et al. 2002). This suggests a fundamental role of ASIC1a in
synaptic transmission and plasticity. Our results indicate that disruption of ASIC1 increases presynaptic neurotransmitter release. In the brain, the greater basal release probability might prevent additional increases in synaptic transmission, and therefore limit long-term potentiation in ASIC1 knockout mice (Hruska-Hageman et al. 2002;
Wemmie et al. 2002). We observed altered paired-pulse modulation and depression during short trains of stimulation in ASIC1 knockout neurons. These types of short-term plasticity are important for information processing in neural networks (e.g. producing reliable response to repetitive activation) and contribute to normal behavior (Blitz et al.
2004). Thus, dysregulation of short-term plasticity may also cause defects in multiple aspects of cognitive function in ASIC1 knockout mice. Our results indicate that the consequences of ASIC1 disruption are complex and impact both presynaptic and postsynaptic mechanisms. In addition, ASIC1a may play a fundamental role in synaptic transmission by regulating presynaptic release probability.
93 A Wild-type ASIC1 KO ASIC2 KO pH 6.0 pH 6.0 pH 6.0 .2 nA 0 5 s
B p < 0.01 C D p < 0.0001 E p < 0.05 ) 2 3 p < 0.05 50 30 A n ( 40 e 1.5 p < 0.01 y d r u e y (pA/pF)
t 2 20 t i v i l 30 s) o s p c
1 n D m e e τ ( a
20 d
t k 1 10 n % R a 0.5 e e
10 r r P u 0 0 0 C 0 Wild-type ASIC1 KO ASIC2 KO Wild-type ASIC2 KO Wild-type ASIC2 KO Glutamatergic GABAergic (53) (13) (31) (52) (30) (37) (31) (24) (18)
Figure 3.1: H+-gated currents of autaptic neurons in microisland cultures (continued)
94 Figure 3.1: continued
(A) Representative traces of H+-gated currents of wild-type, ASIC1 knockout (ASIC1
KO), and ASIC2 knockout (ASIC2 KO) neurons in microisland culture. To evoke H+- gated currents, we exchanged the extracellular solution of pH 7.4 to pH 6.0 for 6-7 seconds (grey bars). To estimate recovery from desensitization, the pH of the extracellular solution was returned to 7.4 for approximately 2.5 seconds, and then a second pH 6.0 application was made in wild-type and ASIC2 knockout neurons. (B)
Peak amplitude of pH 6.0-induced H+-gated currents. (C) The desensitization time
+ constants (τD) of H -gated current at pH 6.0. (D) Recovery from desensitization was determined by applying two pulses of pH 6.0 with an interval of 2.5 seconds. Recovery was assessed by comparing the peak current amplitude in response to the first pH 6.0 applications to the peak current amplitude of the second pH 6.0 application. (E) The
current density of H+-gated currents in glutamatergic and GABAergic autaptic neurons
from wild-type mice. Current density was calculated by dividing the peak amplitude of
pH 6.0-induced current by whole-cell membrane capacitance. Numbers in the
parentheses indicates the numbers of neurons examined. All data in this and later figures
are expressed as the mean ± the standard error of the mean. One-way ANOVA (B) and unpaired t test (C-E) were used to assess statistical significance.
95 A GABAergic neuron B GABA A R PSC 6 )
Wild-type ASIC1 KO ASIC2 KO A n
( 5
e
d 4 u t i l
p 3 m a 2 k a e
1 nA 1 P 50 ms 0 Wild-type ASIC1 KO ASIC2 KO
C Glutamatergic neuron Wild-type ASIC1 KO ASIC2 KO
NMDAR EPSC 1 nA 50 ms AMPAR EPSC
D AMPAR EPSC E NMDAR EPSC F AMPAR EPSC / NMDAR EPSC ratio p < 0.05 p < 0.05 5 2.0 6 ) ) A A n 4 n 5 ratio
1.5 A 4 3 D M
N 3 1.0 2 / A 2 P
1 0.5 M A 1 Peak amplitude ( Peak amplitude ( 0 0.0 0 Wild-type ASIC1 KO ASIC2 KO Wild-type ASIC1 KO ASIC2 KO Wild-type ASIC1 KO ASIC2 KO
Figure 3.2: Whole-cell action potential-evoked postsynaptic currents in GABAergic and glutamatergic autaptic neurons. (continued)
96 Figure 3.2: continued
(A) Representative traces of whole-cell GABAA receptor-mediated postsynaptic current
(GABAAR PSC) of wild-type, ASIC1 knockout, and ASIC2 knockout neurons. A two- millisecond depolarization to +10 mV from a -70 mV holding potential was used to evoke neurotransmitter release and postsynaptic currents. (B) Quantification of peak amplitude of GABAAR PSC. n = 62, 48, and 32 for wild-type, ASIC1 knockout, and
ASIC2 knockout. (C) Representative traces of whole-cell AMPA receptor-mediated
EPSC (black), and NMDA receptor-mediated EPSC (grey) of wild-type, ASIC1 knockout, and ASIC2 knockout neurons. AMPAR EPSCs were recorded in the presence of 1 mM Mg2+. NMDAR EPSCs were isolated in Mg2+-free extracellular solution using
10 µM CNQX, an AMPA receptor antagonist. Membrane potential was held at -70 mV.
(D-F) Average peak amplitude of the AMPAR EPSCs (D), NMDAR EPSCs (E), and the
average AMPAR EPSC / NMDAR EPSC ratio from individual neurons (F). n = 40, 42, and 28 for wild-type, ASIC1 knockout, and ASIC2 knockout. One-way ANOVA was used to assess statistical significance.
97 10 mV A B n.s. -70 mV 7 + CNQX / AP5
) 6 t C n S e
r 5 P r u E
c
4 l R a A u P 3 d
NMDAR EPSC i M s A e (+ CNQX) 2 R % A ( 1 1 n 0 AMPAR EPSC 10 ms Wild-type ASIC1 KO (+ AP5) n.s. D 5 C b )
c t
b. CNQX / AP5 C
n 4 S e r P r u E
c 3
l R a A A
c. CNQX / AP5 u P d
i 2
+ amiloride M s A e
0.5 n R
% 1 a. AP5 5 ms ( 0 + Amiloride Wash - Amiloride + Amiloride pH 6.0 pH 6.0 pH 6.0 A 0.2 n 2 s F - Amiloride E + Amiloride Wild-type AMPAR EPSC ) 120
ASIC1 KO AMPAR EPSC % (
- Amiloride + Amiloride - Amiloride + Amiloride C 100 S P E
80 R A
P 60 M A
40 e v i t a 0.5 nA
1 nA 20 l 50 ms e
50 ms R 0 Wild-type ASIC1 KO
Figure 3.3: ASIC currents do not contribute to the peak amplitude of EPSC in glutamatergic autaptic neurons (continued)
98 Figure 3.3: continued
(A) Representative traces of AMPAR and NMDAR EPSC (grey traces) of a wild-type
autaptic neuron. In the presence of 10 µM CNQX and 50 µM AP5, small residual current was observed (black trace). Vertical dotted lines indicate the time of the peak amplitude of AMPAR and NMDAR EPSCs. A voltage step above the traces indicates transient membrane depolarization from -70 mV to 10 mV for 2 ms used to evoke an action potential. (B) The ratio of the residual current measured in the presence of CNQX and
AP5 to the peak AMPAR EPSC in wild-type (n = 7) and ASIC1 knockout (n = 7) neurons. There is no significant difference in the ratio of residual current to AMPAR
EPSC between wild-type and ASIC1 knockout neurons (unpaired t test). (C)
Representative traces of AMPAR EPSCs and residual current in the presence of CNQX and AP5 in a wild-type autaptic neuron (upper traces). AMPAR EPSCs (a, grey trace) were recorded in the presence of AP5. Residual current (b, black trace) was isolated using CNQX and AP5 and was not affected by 300 µM amiloride (c, grey trace). Lowers traces show amiloride (300 µM) inhibition of H+-gated currents evoked by extracellular acid application (pH 6.0, grey bars). (D) The ratio of the residual current to the AMPAR
EPSC before and during amiloride application. Amiloride did not change the ratio significantly (n = 5, paired t test). (E) Representative traces of AMPAR EPSCs before (- amiloride, black) and during 300 µM amiloride application (+ amiloride, gray) in wild- type and ASIC1 knockout neurons. (continued)
99 Figure 3.3: continued
(F) Quantification of the effect of amiloride on AMPAR EPSCs. Current amplitude
during amiloride application (+ amiloride) was normalized to current amplitude before
amiloride treatment (- amiloride). Amiloride did not change the peak amplitude of
AMPAR EPSC significantly (n = 8, p = 0.10 for wild-type, and n = 8, p = 0.65 for ASIC1 knockout, paired t test).
100 A n = 5 B n = 4 1.2 1.2 C C S S P P 1 1 E E 0.8 AR AR 0.8 pH 6.0 D P pH 6.0 M M 0.6 N A 0.6
d d e e z z 0.4 0.4 i i l l a a m m r r 0.2 0.2 o o N N 0 0 0 50 100 150 200 250 300 350 400 0 50 100 150 200 Time (s) Time (s) C D n = 5 n = 6 2.5 1.2 C C S S 1 P 2
A 0.8 A R P 1.5
B pH 6.0 A AMPAR E
G 0.6
d d e 1 e z i z l
i 0.4 l a a m r
m 0.5 r
0.2 o
o PDBu N N 0 0 0 50 100 150 200 0 50 100 150 200 Time (s) Time (s)
Figure 3.4: The effects of acute ASIC activation by exogenously applied acid on postsynaptic currents (continued)
101 Figure 3.4: continued
(A-C) Time courses of the changes of AMPAR EPSC (A), NMDAR EPSC (B), and
GABAAR PSC (C) following ASIC activation in wild-type autaptic neurons.
Postsynaptic currents with stable peak amplitudes were recorded before and after a 6-7 second application of pH 6.0 extracellular solution (arrows and grey bars), which induced typical H+-gated currents in wild-type neurons (inset, grey traces). Postsynaptic currents were normalized to the currents preceding the acid application. Representative traces of
postsynaptic currents before and after acid application are shown (inset, black traces).
Scale bars: 0.5 nA / 50 ms for postsynaptic currents, and 0.5 nA / 2 s for H+-gated
currents. (D) The effect of PDBu, a diacylglycerol analog on AMPAR EPSC. PDBu (2
µM) was applied continuously for 2 minutes as indicated by the black bar in the graph.
AMPAR EPSCs were normalized to the EPSCs preceding PDBu application. AMPAR
EPSC traces before and during PDBu application are shown (inset). Scale bars: 0.5 nA
and 50 ms.
102 A Wild-type ASIC1 KO ASIC2 KO 1 nA 50 ms B 1.2 1 * * * 0.8 Wild-type R
P 0.6
P ASIC1 KO 0.4 ASIC2 KO
0.2
0 20 50 100 200 400 Interpulse interval (ms)
C Wild-type ASIC1 KO
50 ms
D C 1.1 S P
E 1
AR 0.9 Wild-type P n = 22 M
A 0.8 ASIC1 KO d
e ** * n = 21
z 0.7 i l **
a ** 0.6 m r o
N 0.5 0 50 100 150 200 250 Time (ms)
Figure 3.5: Paired-pulse ratio and AMPAR EPSCs during short trains of repetitive stimuli in autaptic neurons (continued)
103 Figure 3.5: continued
(A) Representative traces of AMPAR EPSCs evoked by paired-pulse in wild-type,
ASIC1 knockout, and ASIC2 knockout neurons. Glutamatergic autaptic neurons were
stimulated with a pair of stimuli with 50 ms interval in the presence of 2 mM Ca2+ and 1 mM Mg2+. (B) Quantification of paired-pulse ratio (PPR) at intervals from 20 ms to 400
ms in wild-type, ASIC1 knockout, and ASIC2 knockout neurons. PPR was calculated as
the ratio of the peak amplitude of the second AMPAR EPSC to the peak amplitude of the
first AMPAR EPSC. n = 11-58 for wild-type, n = 7-29 for ASIC1 knockout, and n = 6-
10 for ASIC2 knockout neurons. * p < 0.05, wild-type versus ASIC1 knockout neurons,
one-way ANOVA. (C) Representative trace of AMPAR EPSCs evoked by five
successive stimuli with 50 ms interval in the presence of 2 mM Ca2+ and 1 mM Mg2+ in wild-type and ASIC1 knockout neurons. (D) Quantification of AMPAR EPSCs during a short train of five repetitive stimuli in wild-type and ASIC1 knockout neurons. * p <
0.05, ** p < 0.01, unpaired t test.
104 AMPAR EPSC NMDAR EPSC A B AMPAR EPSC 0.8 mM 10 mM 0.8 mM 10 mM NMDAR EPSC
GABA A R PSC
) 120 % (
e 100 d u 0.5 nA t i 0.5 nA l 50 ms 50 ms p 80 m a
t
GABA A R PSC n 60 e r 0.8 mM 10 mM r u
c 40
e v i t
a 20 l e R 0
1 nA Wild-type ASIC1 KO ASIC2 KO 50 ms
0.8 mM HEPES 10 mM HEPES n = 15 C D 1.2 n.s. 1 0.8 R
P 0.6 P 0.4
2 nA 0.2 50 ms 0 0.8 mM HEPES 10 mM HEPES E 10 mV F -70 mV 1st stimulus 7 2nd stimulus ) t C 6 n
+ CNQX / AP5 S e r P
r 5 E u
c
R l 4 A a u P
d 3 i M s A e 2 R
AMPAR EPSC % ( 1 (+ AP5) 1 nA 20 ms 0 Wild-type ASIC1 KO
G Wild-type ASIC1 KO H - Amiloride - Amiloride + Amiloride - Amiloride + Amiloride 120 + Amiloride
) 100 % (
R 80 P P
e 60 v i t a
l 40 e R 20 1 nA
0.5 nA 0 50 ms 50 ms Wild-type ASIC1 KO
Figure 3.6: The effects of HEPES concentration and amiloride on AP-evoked postsynaptic currents and the paired-pulse ratio (continued)
105 Figure 3.6: continued
(A) Representative traces of AMPAR EPSC, NMDAR EPSC, and GABAAR PSC from
wild-type neurons. Postsynaptic currents were recorded from the same autaptic neuron in
0.8 mM (black) and 10 mM (grey) HEPES-containing extracellular solutions. (B) The
effect of the HEPES concentration on postsynaptic currents. Postsynaptic currents were
recorded in 0.8 mM and 10 mM HEPES-containing extracellular solutions from wild-
type, ASIC1 knockout, or ASIC2 knockout neurons. Within each neuron, current
amplitude in 10 mM HEPES solution was normalized to that in 0.8 mM HEPES solution.
These relative current amplitudes (%) were not significantly different between wild-type,
ASIC1 knockout, and ASIC2 knockout neurons (n = 6-24, p = 0.48, 0.83, and 0.60 for
AMPAR EPSC, NMDAR EPSC, and GABAAR PSC respectively, one-way ANOVA).
(C) Representative traces of AMPAR EPSC in a wild-type neuron evoked by the paired- pulse protocol of 50 ms interval in the extracellular solution containing 2 mM Ca2+ and 1 mM Mg2+ with 0.8 mM (black) and 10 mM (grey) HEPES. (D) Quantification of the
effect of the HEPES concentration on the PPR. The PPR was not significantly different
between 0.8 mM and 10 mM HEPES (n = 15, p = 0.81, paired t test). (E) Representative
traces of AMPAR EPSC and residual current in the presence of CNQX and AP5 in a
wild-type autaptic neuron. AMPAR EPSC was recorded in the presence of AP5 (grey
trace), and residual current was isolated using CNQX and AP5 (black trace). Voltage
steps above the traces indicate transient membrane depolarization from -70 mV to 10 mV
used to evoke action potentials with 50 ms interval. (continued)
106 Figure 3.6: continued
(F) Quantification of the ratio of the residual current in the presence of CNQX and AP5 to peak AMPAR EPSC in wild-type (n = 5) and ASIC1 knockout (n = 4) neurons. There
is no significant difference in the contribution of residual current to peak AMPAR EPSC
between the first and the second stimulation in either a wild-type or an ASIC1 knockout
neuron (n = 5, p = 0.94 for wild-type; n = 4, p = 0.73 for ASIC1 knockout, paired t test).
(G) Representative traces of AMPAR EPSC evoked by the paired-pulse protocol of 50
ms interval before (- amiloride, black) and during 300 µM amiloride application (+
amiloride, gray) in wild-type and ASIC1 knockout neurons. (H) Quantification of the
effect of the amiloride on PPR. PPR during amiloride application (+ amiloride) was
normalized to PPR before amiloride treatment (- amiloride). Amiloride did not change
PPR significantly in either wild-type or ASIC1 knockout neurons (n = 4, p = 0.51 for
wild-type; n = 4, p = 0.17 for ASIC1 knockout, paired t test).
107 A Wild-type ASIC1 KO 50 pA 50 pA Wild-type 1 sec ASIC1 KO 1 sec 5 pA 5 pA 5 ms 5 ms
1 30 n.s.
B ) 25 A e y
0.8 p t v ( i i
C l t 20 i e a S b l
0.6 d P a u u 15 t b E i m l o
r 0.4 u m p 10 p
C Wild-type m
0.2 ASIC1 KO a 5 0 0 0 20 40 60 80 Wild-type ASIC1 KO mEPSC amplitude (pA)
C 1 8 p < 0.05 ) z
e 0.8 y H
t 6 v ( i i C
l t i y S a b l
0.6 c P a u n 4 b E e m o
0.4 u r m u q
p Wild-type C e 2 r 0.2 ASIC1 KO f 0 0 0 20 40 60 80 Wild-type ASIC1 KO mEPSC frequency (Hz)
Figure 3.7: Quantal analyses of AMPAR-mediated miniature EPSC (mEPSC) of hippocampal neurons in mass culture (continued)
108 Figure 3.7: continued
(A) Representative traces of mEPSC in wild-type (left) and ASIC1 knockout (right)
neurons in mass culture. AMPAR-mediated mEPSC was recorded in extracellular solution containing 1 mM Mg2+, 30 µM bicuculline, and 1 µM tetrodotoxin. Averaged
mEPSCs in the same wild-type and ASIC1 knockout neurons are also shown (lower
traces). (B) Cumulative histogram of peak amplitude of mEPSC from 1179 and 2399
events from 11 wild-type and 8 ASIC1 knockout neurons respectively (left). Average
peak amplitude of mEPSC was calculated in each neuron, and quantified as a bar graph
(right, the mean ± the SEM). (C) Cumulative histogram of instantaneous frequency of mEPSC (left). Average frequency of mEPSC was calculated in each neuron, and quantified as bar graph (right). Statistical significance was assessed using unpaired t test.
109 A Wild-type Sucrose Sucrose + CNQX - CNQX 0.2 nA 2 s ASIC1KO Sucrose 0.2 nA 2 s B n.s. C n.s. 1.2 0.6
) 1 g C n n i ( 0.8 l l 0.4 i f e e z i r
0.6 s
P P R
R 0.4 0.2 R R 0.2 0 0 Wild-type ASIC1 KO Wild-type ASIC1 KO (20) (24) (17) (20)
Figure 3.8: Readily-releasable pool (RRP) size and RRP refilling of wild-type and ASIC1 knockout neurons (continued)
110 Figure 3.8: continued
(A) Representative recordings of sucrose response in wild-type (upper traces) and ASIC1 knockout (lower trace) neurons in mass culture. Hypertonic sucrose (500 mM) was applied for 4-5 seconds (grey bars), and the sucrose-induced current was recorded in extracellular solution containing 1 mM Mg2+, 30 µM bicuculline, and 1 µM tetrodotoxin.
The sucrose response was inhibited completely by 10 µM CNQX (+ CNQX, upper right trace). (B) Total charge transfer by the application of hypertonic sucrose. RRP size was estimated from hypertonic sucrose-induced charge transfer calculated by integrating the transient component of the sucrose response curve. (C) RRP refilling was estimated from analysis of a second sucrose response evoked 3-4 seconds after the first sucrose application. The ratio of the charge transfer by the second sucrose application to the charge transfer induced by the first sucrose application was calculated. Numbers in parentheses indicate the numbers of neurons examined. Statistical significance was assessed using unpaired t test.
111 A Mock b a b 1.2 C
a S
P 1 E 0.8 AR D
M 0.6 N b MK-801 d e 0.4 z i a l Mock a 0.2 m MK-801 r o
N 0 0 5 10 15 20 25 50 ms Stimulus number
B Wild-type NMDAR EPSC ASIC1 KO NMDAR EPSC
1st 5th 10th 20th
50 ms 50 ms C D C
S 1 Wild-type p < 0.05 P 30 E ASIC1 KO 0.8 25 AR
D 0.6 20 M N 15 d 0.4 e z
i 10 l
a 0.2
m 5 r τ (stimulus number) o
N 0 0 0 10 20 30 Wild-type ASIC1 KO Stimulus number
Figure 3.9: The progressive block of NMDAR by MK-801 in glutamatergic autaptic neurons (continued)
112 Figure 3.9: continued
(A) Representative traces of the first (a, black) and 22nd (b, grey) NMDAR EPSC (left),
and the time course of NMDAR EPSC change (right) in the presence or absence of 10
µM MK-801 in a wild-type autaptic neuron. EPSCs were evoked in the presence of MK-
801 or vehicle (mock) every 10 seconds for 4 minutes in Mg2+-free solution with 10 µM
CNQX. (B) Representative trace of NMDAR EPSCs (1st, 5th, 10th, and 20th) in the presence of MK-801 in wild-type (left) and ASIC1 knockout neurons (right). (C)
Stimulus-dependent decrease of NMDAR EPSC by MK-801. Peak amplitudes of
NMDAR EPSC were normalized to that of the first NMDAR EPSC in the presence of
MK-801. n = 8 for wild-type, and n = 8 for ASIC1 knockout neurons. (D)
Quantification of the rate of the NMDAR EPSC decrease by MK-801. Tau (τ ) in stimulus number was calculated by fitting the curve of NMDAR EPSC change to single exponential equations, I(n) = I1st exp(-n/τ), where n is stimulus number, I(n) is peak
amplitude of nth NMDAR EPSC, and I1st is the peak amplitude of the first NDMAR
EPSC in the presence of MK-801. Statistical significance was assessed using an unpaired t test.
113 A AP-evoked Hypertonic sucrose AMPAR EPSC response Wild-type Sucrose 1 nA
50 ms 0.2 nA 2 s ASIC1 KO Sucrose 1 nA 50 ms 0.2 nA 2 s
n.s. n.s. B 0.3 C 3 ) ) C C n n ( (
0.2 2 e s ro c u
0.1 s 1
AP-EPSC
Q Q
0 0 Wild-type ASIC1 KO Wild-type ASIC1 KO
D 15 p < 0.05 p < 0.05 ) % (
y t i l i 10 b a b o r p
e
s 5 a e l e R 0 Wild-type ASIC1 KO ASIC2 KO (27) (19) (12)
Figure 3.10: Release probability measured in glutamatergic autaptic neurons on microislands (continued)
114 Figure 3.10: continued
(A) Representative traces of AP-evoked AMPAR EPSC (left) and 500 mM sucrose-
induced response (right) of wild-type (upper traces) and ASIC1 knockout (lower traces)
neurons. Both AMPAR EPSC and sucrose response were recorded in the same neuron in
the presence of 1 mM Mg2+. (B-C) Quantification of AP-evoked (B) and sucrose- induced (C) charge transfers calculated by integrating AMPAR EPSC and hypertonic sucrose response curves. n = 27 for wild-type and n = 19 for ASIC1 knockout. (D)
Quantification of the release probability of wild-type, ASIC1 knockout, and ASIC2
knockout neurons. The probability of neurotransmitter release was calculated by dividing
total charge transfer of AP-evoked EPSC by hypertonic sucrose-induced charge transfer.
Numbers in the parentheses indicate the numbers of neurons examined. One-way
ANOVA was used to determined statistical significance.
115 A H +-gated current B H +-gated current Vector ASIC1a 2.5 p < 0.05 pH 6.0 pH 6.0 ) A n ( 2 e d u t i l 1.5 p m a 1 k a e
P 0.5
0.5 nA 0 2 s Vector ASIC1a C AMPAR EPSC D AMPAR EPSC Vector ASIC1a 6 p < 0.01 ) A
n 5 (
e
d 4 u t i l
p 3 m a 2 k a e 1 P 1 nA 0 50 ms Vector ASIC1a E F 1.2 Vector ASIC1a * * 1
0.8 R
P 0.6 P 0.4 ASIC1a 0.2 Vector
0 0.5 nA 0.5 nA 50 ms 50 ms 0 20 40 60 Interpulse interval (ms)
Figure 3.11: Rescue of ASIC1 knockout neurons by ASIC1a expression (continued)
116 Figure 3.11: continued
(A) Representative traces of H+-gated currents in ASIC1 knockout neurons transfected with vector (pcDNA3.1, left) or human ASIC1a (right). To evoke H+-gated currents, we
exchanged the extracellular solution of pH 7.4 to pH 6.0 for 6-7 seconds (grey bars). (B)
Peak amplitude of pH 6.0-induced H+-gated currents. n = 6 for ASIC1a-transfected neurons, and n = 9 for vector-transfected neurons. Unpaired t test. (C) Representative traces of whole-cell AMPA receptor (AMPAR)-mediated EPSC in ASIC1 knockout neurons transfected with vector or ASIC1a. AMPAR EPSCs were recorded in the presence of 1 mM Mg2+, and membrane potential was held at -70 mV. (D) Peak amplitude of the AMPAR EPSCs. n = 6 for ASIC1a-transfected neurons, and n = 9 for vector-transfected neurons. Unpaired t test. (E) Representative traces of AMPAR
EPSCs evoked by a paired-pulse with 50 ms interval in the presence of 2 mM Ca2+ and 1 mM Mg2+. (F) Quantification of the paired-pulse ratio (PPR) at intervals from 20 ms to
50 ms. The PPR was calculated as the ratio of the peak amplitude of the second AMPAR
EPSC to the peak amplitude of the first AMPAR EPSC. n = 4-6 for ASIC1a-transfected neurons, n = 5-9 for vector-transfected neurons. * p < 0.05, unpaired t test. Single glutamatergic neurons on microisland were selected for recording H+-gated currents and
AMPAR EPSC in these experiments.
117
CHAPTER 4
TONIC FIRING OF CEREBELLAR PURKINJE CELL IS ALTERED IN ACID-SENSING ION CHANNEL 1 AND 2 DOUBLE KNOCKOUT MICE
Acid-sensing ion channels (ASICs) are voltage-independent ion channels
activated by extracellular acid. Both ASIC1 and ASIC2 are expressed throughout the brain including the cerebellum. ASIC1 knockout mice display impaired eyeblink conditioning, suggesting the physiologic role of ASICs in normal cerebellar function.
Purkinje cells, the sole output neurons of the cerebellar cortex to deep cerebellar nuclei, express ASICs and have intrinsic firing property which is critical for cerebellar function.
To investigate how ASICs contribute to cerebellar function, we compared the tonic firing of Purkinje cells in acute cerebellar slices from wild-type and ASIC knockout mice.
Compared to wild-type mice, the rate or regularity of Purkinje cell firing was not different in either ASIC1 or ASIC2 knockout mice. However, the firing rate was increased significantly in ASIC1/ASIC2 double knockout mice (ASIC1/2 DKO) by
~15%. Increased firing rate of Purkinje cells from ASIC1/2 DKO mice was also observed in the presence of inhibitors of fast synaptic transmission. This suggests that disruption of ASIC1 and ASIC2 genes alters the intrinsic firing properties of Purkinje
118 cells. In addition, Purkinje cells from ASIC1/2 DKO mice displayed more regular firing, but only in the presence of endogenous synaptic transmission, suggesting a reduced synaptic activity to Purkinje cells in ASIC1/2 DKO mice. These results indicate that both
ASIC1 and ASIC2 may regulate Purkinje cell firing independently.
4.1. INTRODUCTION
Acid-sensing ion channels (ASICs) are H+-gated, voltage-independent cation channels activated by extracellular acid (Krishtal 2003; Waldmann 2001; Wemmie et al.
2006). There are four ASIC genes that encode six subunits (ASIC1a, ASIC1b, ASIC2a,
ASIC2b, ASIC3, and ASIC4) expressed in distinct and overlapping patterns throughout
the nervous system. Each subunit can form either homomultimeric or heteromultimeric
channels. Central neurons express ASIC1a, ASIC2a and ASIC2b (Garcia-Anoveros et al.
1997; Lingueglia et al. 1997; Price et al. 1996; Waldmann et al. 1997b; Waldmann et al.
1996). In central neurons, H+-gated currents activated by moderate acidosis (pH > 6) are
due to ASIC1a homomultimeric channels and ASIC1a/2a heteromultimeric channels.
Disruption of ASIC1 eliminates H+-gated currents activated by moderate acidosis whereas disruption of ASIC2 alters the characteristics of H+-gated currents (Askwith et al. 2004; Wemmie et al. 2002). During the prolonged acidosis that accompanies cerebral ischemia, ASIC1a homomultimeric channels contribute to neuronal death (Xiong et al.
2004). Under normal conditions, ASIC1 contributes to a variety of behaviors linked to learning and memory (Wemmie et al. 2003; Wemmie et al. 2002; Wemmie et al. 2004).
119 Mice with disruption of ASIC1 show deficiencies in spatial learning and fear
conditioning (Wemmie et al. 2003; Wemmie et al. 2002; Wemmie et al. 2004). Eyeblink
conditioning is also impaired in ASIC1 knockout mice (Wemmie et al. 2002), suggesting
ASIC1 is necessary for normal cerebellar function. ASIC1 and ASIC2 mRNAs are
highly expressed in the cerebellum (Garcia-Anoveros et al. 1997; Lingueglia et al. 1997;
Waldmann et al. 1997b). Immunohistochemistry of ASIC1a and ASIC2a indicate that
ASIC protein is expressed predominately on the cell body and dendrites of Purkinje cells
and granule cells in the cerebellum (Duggan et al. 2002; Jovov et al. 2003; Wemmie et al.
2003). Further, acutely dissociated Purkinje cells and granule cells display H+-gated currents with characteristics similar to ASIC1a and ASIC2a-containing channels (Allen and Attwell 2002; Bolshakov et al. 2002). These results suggest that ASIC1, and possibly ASIC2, impact Purkinje cell function in the cerebellum.
Cerebellar Purkinje cells are the sole efferent neurons of the cerebellar cortex, and
project to deep cerebellar nuclei (DCN). Purkinje cells have intrinsic pacemaker activity,
which allows rapid and regular spontaneous firing of action potentials (Raman and Bean
1999b). By firing action potentials spontaneously, they send tonic inhibitory signal to
DCN. Tonic spontaneous firing of Purkinje cells is regulated by many voltage-gated ion
channels (Raman and Bean 1999a), and fast synaptic transmission (Hausser and Clark
1997). Proper regulation of spontaneous firing of Purkinje cells is important for normal cerebellar control of motor function (Raman et al. 1997). To gain insight into how
ASICs contribute to normal cerebellar function, we tested the hypothesis that ASICs regulate Purkinje cell firing. We compared the tonic firing of Purkinje cells from wild- type, ASIC1 knockout, ASIC2 knockout, and ASIC1/ASIC2 double knockout mice using
120 extracellular recording of acute cerebellar slices. We find that the tonic firing of
cerebellar Purkinje cells from ASIC1/ASIC2 double knockout mice is altered.
4.2. MATERIALS AND METHODS
4.2.1. ASIC knockout mice
We performed extracellular recording using mice from 4 different groups of
breeding pairs; ASIC1+/-ASIC2+/+ x ASIC1+/-ASIC2+/+ (group A), ASIC1+/+ASIC2+/- x
ASIC1+/+ASIC2+/- (group B), ASIC1+/-ASIC2+/- x ASIC1+/-ASIC2+/- (group C), or ASIC1-/-
ASIC2-/- x ASIC1-/-ASIC2-/- (group D). Wild-type mice were from group A, B, or C;
ASIC1 knockout mice from group A or C; ASIC2 knockout mice from group B or C;
ASIC1/ASIC2 double knockout mice from group C or D. We observed no difference in
the tonic firing of cerebellar Purkinje cells between mice of the same genotype from
different groups of breeding pairs. Genotype of mice was determined using PCR and
confirmed after the harvest of cerebellar tissue.
4.2.2. Extracellular recording of cerebellar Purkinje cells
Adult mice of postnatal day 32-53 were anesthetized with isoflurane and decapitated. After the cerebellum was removed quickly, sagittal slices (300 µm) of the vermis and intermediate part of cerebellar hemisphere were prepared using a vibratome
(Campden Instruments) in chilled oxygenated artificial cerebrospinal fluid (aCSF), which contains 124 mM NaCl, 5 mM KCl, 1.2 mM MgSO4, 2.5 mM CaCl2, 28 mM NaHCO3,
1.3 mM KH2PO4, and 10 mM glucose. Slices were recovered in an interface chamber
121 (Harvard Apparatus) at 31°C for an hour before recording, and were continuously
perfused with aCSF bubbled with 95% O2 / 5% CO2 at the rate of 1 mL/minute.
Recording microelectrodes with 2-3 MΩ were pulled using a vertical micropipette
puller (Narishige), filled with 4 M NaCl, and placed into the Purkinje cell layer of
cerebellar slices. Stimulating micropipettes were filled with γ–aminobutyric acid (GABA
in regular aCSF), acidic solution (pH 6.0, aCSF without NaHCO3 and KH2PO4 replaced
with 10 mM HEPES), or amiloride (1 mM in aCSF), and connected to a Picospritzer
(General Valve). The pressure and duration of pneumatic pulse of the Picospritzer were
adjusted so that single pulse allows a droplet at the tip of stimulating micropipette.
Stimulating micropipette was placed ~100 µm away from the recording microelectrode.
In some recording, cyano-7-nitroquinoxaline-2,3-dione (CNQX, 10 µM), and picrotoxin
(100 µM) were added to aCSF to inhibit fast synaptic transmission. We recorded the
firing of Purkinje cells in lobule V and lobule VI, and the temperature was maintained
constant at 31 ± 1°C using TC-102 temperature controller (Medical Systems). We
focused on recording Purkinje cells with tonic firing pattern at constant rates for at least 2
minutes, and excluded cells with trimodal firing pattern. Data were collected using 8700
Cell Explorer preamplifier (Dagan), TM506 amplifier (Tektronix), CED 1401 plus A/D
converter, and Spike2 software (Cambridge Electronic Design). All reagents were
purchased from Sigma-Aldrich (St. Louis, MO), or Fisher Scientific (Waltham, MA).
Data were analyzed offline using Spike2 software. Average firing rates (Hz) of
Purkinje cells were calculated from two-minute recording. Coefficient of variation of interspike intervals (ISI) of a Purkinje cell was calculated by dividing the standard deviation of ISI with the average ISI of the Purkinje cell. Data are presented as the mean 122 ± the standard error of the mean (SEM). As appropriate, a two-tailed Student’s t test and one-way ANOVA with Bonferroni’s simultaneous multiple comparisons were used for statistical analyses, and performed with Minitab15 software (Minitab Inc, State College,
PA).
4.3. RESULTS
4.3.1. Tonic firing rate of cerebellar Purkinje cells
To examine how ASIC1 and ASIC2 contribute to the firing of cerebellar Purkinje
cells, we compared basal firing rates of wild-type, ASIC1 knockout (ASIC1 KO), and
ASIC2 knockout (ASIC2 KO) mice. For this study, we focused on recording Purkinje
cells with tonic firing pattern at constant rates, and excluded cells with trimodal firing
pattern. In wild-type mice, Purkinje cells in lobule V and VI displayed an average firing
rate of 84.7 ± 2.4 Hz (n = 103 cells, Table 1, Fig. 1). There was no significant difference
in firing rate between lobule V and lobule VI Purkinje cells (85.6 ± 3.1, n = 57 cells in
lobule V; 81.4 ± 3.8, n = 46 cells in lobule VI, p = 0.38, unpaired t test). Compared to
wild-type mice, basal firing rate was not different significantly in either ASIC1 KO or
ASIC2 KO mice (p = 1.00 for ASIC1 KO mice; p = 0.72 for ASIC2 KO mice, one-way
ANOVA, Table 1, Fig. 1). Because there was no significant effect of loss of either
ASIC1 or ASIC2 gene on basal firing rate, we considered the possibility that either
ASIC1 or ASIC2 might compensate the absence of each other. To test this hypothesis,
we examined the firing rate of Purkinje cells in ASIC1 and ASIC2 double knockout
(ASIC1/2 DKO) mice. Compared to wild-type mice, firing rate in ASIC1/2 DKO mice
123 was significantly higher (97.8 ± 2.3, n = 108 cells, p < 0.001 versus wild-type, one-way
ANOVA, Table 1, Fig. 1). This indicates that either ASIC1 or ASIC2 subunit may
decrease basal firing rate of Purkinje cells independently.
Purkinje cells receive glutamatergic (excitatory) synaptic input from climbing
fibers, and parallel fibers from cerebellar granule cells. Purkinje cells also receive
GABAergic (inhibitory) synaptic inputs from their own recurrent collateral axons, basket
cells and stellate cells. Since the extracellular solution did not include inhibitors of
synaptic transmission in our recordings, the altered firing rate observed in the ASIC1/2
DKO mice could be due to altered synaptic transmission or changes in the intrinsic firing
rate of Purkinje cells. To determine this, we applied inhibitors of fast synaptic
transmission (10 µM CNQX for AMPA/kainate receptors, and 100 µM picrotoxin for
GABAA receptors), and examined the effect on tonic firing of Purkinje cells. In the
presence of CNQX and picrotoxin, the firing rate was again significantly higher in
ASIC1/2 DKO mice compared to wild-type and ASIC1 KO mice (p < 0.01, one-way
ANOVA, Table 1, Fig. 2A-B). This suggests that intrinsic spontaneous firing rate of
Purkinje cells is increased in ASIC1/2 DKO mice. To test whether picrotoxin inhibit
GABAergic transmission effectively in our system, we compared the change of tonic firing rate of wild-type Purkinje cells in the absence or presence of picrotoxin. Without picrotoxin, GABA (100 µM in aCSF) applied using Picospritzer deceased the firing rate
(n = 3, Fig. 2C1). However, when slices were perfused with aCSF containing picrotoxin,
GABA did not induced significant change in the firing rate (n = 4, Fig. 2C2), suggesting
that picrotoxin applied in our system inhibit GABAA receptors-mediated signaling in
Purkinje cells effectively. 124
4.3.2. Regularity of tonic firing of cerebellar Purkinje cells
Regularity of Purkinje cell firing was also examined by calculating coefficient of
variation (CV) of interspike intervals (ISI) (Walter et al. 2006) (Fig. 3). Average CV of
firing of wild-type Purkinje cells was 0.278 ± 0.010 (n = 79 cells) in the absence of
inhibitors of fast synaptic transmission. Compared to wild-type mice, the average CV
was slightly reduced in ASIC1 or ASIC2 knockout mice although the difference did not reach statistical significance (p = 0.21 for ASIC1 KO; p = 0.054 for ASIC2 KO, one-way
ANOVA, Table 2 and Fig. 2B). However, CV was smaller significantly in ASIC1/2
DKO mice than in wild-type mice (0.208 ± 0.008, n = 100 cells, p < 0.0001, one-way
ANOVA, Fig. 2A-B). This indicates that Purkinje cell firing is more regular in ASIC1/2
DKO mice than in wild-type mice.
To determine whether the increased regularity of Purkinje cell firing is due to the
change in endogenous synaptic activity or the intrinsic property of Purkinje cells in
ASIC1/2 DKO mice, we compared the regularity of Purkinje cell firing in the presence of
CNQX and picrotoxin. When these inhibitors were applied, CV of ISI were decreased,
suggesting Purkinje cells fired action potentials more regularly without synaptic inputs
(Table 2), which is consistent with the previous report that fast GABAergic synaptic
transmission decreased the regularity of Purkinje cell firing (Hausser and Clark 1997).
Interestingly, in the presence of picrotoxin and CNQX, there were no significant
difference in the CV of ISI between wild-type and ASIC1/2 DKO (p = 1.00, one-way
ANOVA). These results indicate that the regularity of intrinsic firing of Purkinje cells is
not altered in ASIC1/2 DKO mice, but the endogenous synaptic activity to Purkinje cells
125 may be reduced in ASIC1/2 DKO mice, which displayed the increased regularity of
Purkinje cell firing only without CNQX and picrotoxin.
4.3.4. The effects of acute activation and inhibition of ASICs on Purkinje cell firing
Activation of ASIC channels induces depolarization and calcium influx in neurons (Vukicevic and Kellenberger 2004; Yermolaieva et al. 2004). Thus, the activation of ASIC channels by physiologic and microscopic pH fluctuations during the recording could cause the alteration of Purkinje cell firing rate in wild-type mice. Thus, we examined the effect of extracellular acid (pH 6.0), which would be expected to activate ASICs exogenously. Extracellular acid applied using Picospritzer (pH 6.0,
HEPES-buffered aCSF, n = 5) did not affect the firing rate of wild-type Purkinje cells
(Fig. 4A). We also analyzed the effect of amiloride, a nonspecific ASIC blocker. The application of amiloride (1 mM in aCSF, n = 5) did not alter firing rate of wild-type
Purkinje cells either (Fig. 4B). These results suggest that the difference in the rate of
Purkinje cell firing between wild-type and ASIC1/2 DKO mice may not be due to the acute activation of ASICs by physiologic extracellular acidification during recording.
4.4. DISCUSSION
We found that Purkinje cells from ASIC1/2 DKO mice fire action potentials more frequently compared to wild-type mice. Decreased inhibitory synaptic inputs onto
Purkinje cells can increase firing rate of Purkinje cells. However, the firing rate was still higher in ASIC1/2 DKO mice when fast synaptic transmission was inhibited
126 pharmacologically. These results suggest that more frequent firing in ASIC1/2 DKO
mice is due to the altered intrinsic firing property of Purkinje cells rather than change in
synaptic inputs onto Purkinje cells. We also examined the regularity of tonic firing of
Purkinje cells, which was estimated using coefficient of variation (CV) of interspike
intervals (ISI). The CV of ISI in ASIC1/2 DKO mice was smaller significantly compared
to wild-type mice, indicating more regular firing of Purkinje cells. However, this
difference in the regularity of tonic firing was lost when fast synaptic transmission was
inhibited by CNQX and picrotoxin. Because endogenous synaptic activity onto Purkinje cells decreases the regularity of tonic firing (Hausser and Clark 1997), this suggests that synaptic input onto Purkinje cells may also be reduced in ASIC1/2 DKO mice.
We observed increased tonic firing rate of Purkinje cells only in ASIC1/2 DKO
mice. We did not observe significant difference in Purkinje cell firing between wild-type
and either ASIC1 KO or ASIC2 KO mice. These results suggest that both ASIC1 and
ASIC2 regulate the tonic firing of Purkinje cells independently or that either subunit can
compensate for the loss of the other. A modest pH decrease (> pH 6.0), similar to that
thought to occur physiologically in vivo (Krishtal et al. 1987), does not induce H+-gated current in either ASIC1 KO or ASIC1/2 DKO neurons (Wemmie et al. 2002). Thus, if
ASICs were responding to modest acidification to induce changes in Purkinje cell firing, we would expect that the ASIC1 KO neurons would be affected as well as ASIC1/2 DKO neurons. The fact that Purkinje cells from ASIC1 KO mice fire normally suggests that the acute activation of ASICs by protons may not be the underlying mechanism of ASIC regulation of Purkinje cell firing. This is supported by our observation that either
127 exogenous application of acid (pH 6.0) or addition of the ASIC blocker, amiloride did not
change tonic firing of wild-type Purkinje cells.
There are several other possibilities regarding the mechanism of ASIC1 and
ASIC2 involvement in the tonic firing of Purkinje cells. First, there may be another
physiologic ligand of ASICs, other than protons, which binds and activates both ASIC1a
and ASIC2a channels with similar affinity. Second, the combined loss of ASIC1 and
ASIC2 could induce developmental changes which impact Purkinje cell activity
indirectly. Finally, ASIC1 and ASIC2 may affect cerebellar firing through a non-
conventional manner distinct from their proton-dependent gating. Interestingly, a recent report indicates that both ASIC1 and ASIC2 subunit interact directly with voltage-gated ion channels including large-conductance Ca2+-activated K+ channels, which regulate tonic spontaneous firing of Purkinje cells (Sausbier et al. 2004).
Through rapid spontaneous firing, Purkinje cells send tonic inhibitory signals to DCN, which is critical structure for eyeblink conditioning (McCormick and Thompson 1984).
Thus, more frequent intrinsic firing of Purkinje cells in ASIC1/2 DKO mice may prevent
DCN neurons from sending signals for the execution of conditioned response, and cause defects in eyeblink conditioning. Interestingly, we did not observe significant change in tonic firing of Purkinje cells from ASIC1 KO mice, which have been reported to display impaired eyeblink conditioning (Wemmie et al. 2002). This suggests that the defect in
ASIC1 KO mice may be due to a different mechanism such as altered synaptic transmission and plasticity (Cho and Askwith 2008; Wemmie et al. 2002). Although the physiologic consequence of altered tonic firing of cerebellar Purkinje cells in ASIC1/2
128 DKO mice need to be addressed with the detailed examination of cerebellar function, our study indicates that both ASIC1 and ASIC2 are required for the regulation of tonic
Purkinje cell firing and normal cerebellar function.
129 No inhibitor CNQX / Picrotoxin Genotype n Average ± SEM n Average ± SEM
Wild-type 103 (5) 83.7 ± 2.4 81 (5) 81.1 ± 2.5
ASIC1 KO 94 (4) 8 4.3 ± 2.7 80 (5) 80.7 ± 3.0
ASIC2 KO 102 (4) 8 9.2 ± 2.5 80 (4) 84.8 ± 2.7
ASIC1/2 DKO 108 (4) 9 7.8 ± 2.3 *** 80 (4) 92.8 ± 2.5 *
Table 4.1: Firing rates of cerebellar Purkinje cells
Numbers of Purkinje cells examined are indicated under ‘n’. Numbers in the parentheses indicate the number of mice used in each group. Average firing rate are expressed as mean ± the standard error of mean (SEM). ** p < 0.01, *** p < 0.001, compared with wild-type mice, one-way ANOVA.
130 No inhibitor CNQX / Picrotoxin Genotype n Average ± SEM n Average ± SEM
Wild-type 79 (5) 0.278 ± 0.010 81 (5) 0.129 ± 0.005
ASIC1 KO 86 (4) 0.250 ± 0.009 80 (5) 0.125 ± 0.005
ASIC2 KO 100 (4) 0.245 ± 0.008 80 (4) 0.131 ± 0.005
ASIC1/2 DKO 100 (4) 0.208 ± 0.008 *** 80 (4) 0.129 ± 0.008
Table 4.2: Coefficient of variation of interspike intervals of Purkinje cell firing
Numbers of Purkinje cells examined are indicated under ‘n’. Numbers in the parentheses indicate the number of mice used in each group. Coefficients of variation of interspike intervals are expressed as mean ± the standard error of mean (SEM). *** p < 0.0001, compared with wild-type mice, one-way ANOVA.
131 A 20 Wild-type Wild-type: lobule VI, 86 Hz 15
10
5
0 30 60 90 120 150 180 ASIC1 KO: lobule V, 84 Hz 20 ASIC1 KO
15
10
5
0 30 60 90 120 150 180 ASIC2 KO: lobule VI, 89 Hz 20 ASIC2 KO 15
10
5
0 30 60 90 120 150 180
ASIC1/2 DKO: lobule VI, 98 Hz 20 ASIC1/2 DKO 15
10
5 Frequency (%)
50 µV 0 30 60 90 120 150 180 50 ms Firing rate (Hz) B 110
100 *** )
z 90 H (
e t a
r 80
g n i r i
F 70
60
50 Wild-type ASIC1KO ASIC2KO ASIC1/2DKO
Figure 4.1: Firing rates of cerebellar Purkinje cells from wild-type and ASIC knockout mice (continued) 132 Figure 4.1: continued
(A) Representative traces (left, 0.5 second) of tonic firing of Purkinje cells in wild-type,
ASIC1 knockout (ASIC1 KO), ASIC2 knockout (ASIC2 KO), and ASIC1/ASIC2 double knockout (ASIC1/2 DKO). Histograms of firing rates with normal curves show the distribution of firing rates of individual Purkinje cells (right). (B) A bar graph shows the average firing rates with the SEM in wild-type and ASIC knockout mice. Average firing rate of ASIC1/2 DKO mice is higher significantly compared to wild-type. n = 94-108 cells from 4-5 mice for each genotype, *** p < 0.001, One-way ANOVA.
133 A. Picrotoxin/CNQX B. Picrotoxin/CNQX
110 100%
100 ) % ) (
80% z
* y H c (
90 n e e t
u 60% a r q
80 e g r f n
i r e i 40% v F i
70 t a l Wild-type u
m 20% 60 ASIC1/2 DKO u C 50 0% Wild-type ASIC1KO ASIC2KO ASIC1/2DKO 0 10 20 30 Interspike interval (ms)
C1. No inhibitor C2. + Picrotoxin
GABA 120 120
100 GABA 100 ) ) z z 80 80 H H ( (
e e t t a a r r 60 60
g g n n i i r r i i 40 40 F F
20 20
0 0 0 20 40 60 80 100 0 20 40 60 80 100 Time (s) Time (s)
Figure 4.2: Spontaneous firing rates of cerebellar Purkinje cells during inhibition of fast synaptic transmission (continued)
134 Figure 4.2: continued
(A) A bar graph shows the average spontaneous firing rates with the SEM in wild-type and ASIC knockout mice. ACSF containing CNQX (10 µM) and picrotoxin (100 µM) was perfused continuously. Average firing rate of ASIC1/2 DKO mice is higher significantly compared to wild-type. n = 80-81 cells from 4-5 mice for each genotype, ** p < 0.01, One-way ANOVA. (B) Cumulative histogram of interspike intervals of spontaneous firing of individual Purkinje cells from wild-type and ASIC1/2 DKO mice.
(C) The time courses of the change in firing rate of a wild-type Purkinje cells by GABA
(100 µM) applied using Picospritzer (arrows) in the absence (C1) or presence (C2) of picrotoxin (100 µM).
135 A Wild-type ASIC1/2 DKO
* * * * *
* * *
* * *
* * * * *
100 ms B 0.4 No inhibitor Picrotoxin/CNQX 0.3 I S I
f o 0.2 *** V C 0.1
0 Wild-type ASIC1KO ASIC2KO ASIC1/2 DKO
Figure 4.3: Regularity of tonic firing of Purkinje cells from wild-type and ASIC knockout mice (continued)
136 Figure 4.3: continued
(A) Each line indicates action potentials of a wild-type and an ASIC1/2 DKO Purkinje cell. Action potentials, of which interspike intervals (ISI) are not within 50-150% of average ISI, are indicated by asterisks (*). (B) The regularity of Purkinje cell firing was estimated as coefficient of variation (CV), which was calculated in each Purkinje cell by dividing the standard deviation (SD) of ISI with the mean of ISI, i.e. CV = SD / mean.
CV of ISI is smaller significantly in ASIC1/2 DKO mice compared with wild-type mice. n = 79-100 cells from 4-5 mice for each genotype, *** p < 0.0001, one-way ANOVA.
However, in the presence of inhibitors of fast synaptic transmission (picrotoxin and
CNQX), there was no significant difference in CV of ISI between wild-type and ASIC knockout mice. n = 80-81 cells from 4-5 mice for each genotype, p = 1.00, one-way
ANOVA).
137 A )
% n = 5 pH 6 ( 120 e t a r
g
n 100 i r i f
d
e 80 z i l a m r
o 60 N 0 20 40 60 80 100 Time (s)
B ) n = 5
% Amiloride ( 120 e t a r
g
n 100 i r i f
d
e 80 z i l a m r
o 60 N 0 20 40 60 80 100 Time (s)
Figure 4.4: The effects of acute activation and inhibition of ASICs on tonic firing of wild-type Purkinje cells
The time courses of the change of firing rate by exogenous acid application (A, HEPES- buffered aCSF, pH 6.0), and amiloride, a nonspecific ASIC blocker (B, 1 mM in regular aCSF) are presented. Amiloride and pH 6.0 acidic solutions were applied using
Picospritzer (arrows) in the presence of picrotoxin and CNQX.
138
CHAPTER 5
GENERAL DISCUSSION AND FUTURE DIRECTIONS
5.1. Potentiation of acid-sensing ion channels by sulfhydryl compounds
We found that ASIC-mediated H+-gated currents are potentiated by sulfhydryl compounds through two different mechanisms; chelation of metal ions and reducing amino acid residues (Cho and Askwith 2007) (Fig. 1). ASICs were also modulated by reduced glutathione (GSH), an endogenous sulfhydryl compound. Potentiation of ASIC currents by sulfhydryl compounds were observed in both CHO cells expressing ASICs heterologously, and cultured hippocampal neurons.
5.1.1. Physiologic significance
ASICs are tonically inhibited by transition metal ions including Zn2+ because extracellular fluid normally contains transition metal ions in concentrations high enough to inhibit ASICs. Therefore, if the high-affinity metal inhibition of ASICs is regulated
139 physiologically, there should be a mechanism through which contaminating metal ions
are removed from ASICs. However, such mechanism had not been described since the
report of high-affinity Zn2+ inhibition of ASIC1a-containing channels (Chu et al. 2004).
We observed that endogenous sulfhydryl compounds such as reduced glutathione
potentiate ASICs by chelating metal ions, which are already bound to and inhibit ASICs
(Fig. 1A). This finding suggests that the modulation of ASICs by endogenous sulfhydryl
compounds can be a mechanism, by which high-affinity Zn2+ inhibition of ASICs can be
regulated physiologically. Interestingly, endogenous sulfhydryl compounds including reduced glutathione and L-cysteine are released from neurons in synaptic activity- dependent manner (Zangerle et al. 1992). This raises a possibility that endogenous synaptic activity may regulate ASICs through synaptically-released sulfhydryl compounds, which relieve tonic inhibition of ASICs by transition metal ions.
5.1.2. Identifying residues for high-affinity metal inhibition and redox modulation
High-affinity Zn2+-binding residue of ASIC1a has not been identified yet.
Because the lysine residue at 133 probably bears a positive charge in ASIC1a, this
residue is not likely to coordinate metal ion in ASIC1a. Our observation indicates that
both ASIC1a and ASIC1b homomultimers were similarly potentiated in the presence of
TPEN and DTT. ASIC3-mediated H+-gated currents were also increased transiently by
TPEN and DTT whereas ASIC2a homomultimers did not display transient potentiation
by DTT. These suggest that target residue(s) for high-affinity metal inhibition should be
within the extracellular domain common to ASIC1a, ASIC1b, and ASIC3. Structure-
140 function study of ASICs with sequence comparison between ASIC1a, ASIC2a, and
ASIC3 may elucidate the residue(s) required for high-affinity metal inhibition.
Sulfhydryl compounds also induce sustained potentiation of H+-gated currents, which last even after the removal of the compounds (Fig. 1B). This sustained potentiation of ASIC current is reversed to pretreatment level when DTNB, an oxidizing agent is applied. These results suggest that sulfhydryl compounds also modulate ASICs by reducing amino acid residue(s) of the extracellular domain. In the extracellular domain of ASICs, there are 14 conserved cysteine residues, which form 7 disulfide bonds in ASIC1a (Jasti et al. 2007). Consistent with this, all ASIC homomultimers examined in our study displayed sustained potentiation by DTT. Therefore, targets for the reduction by sulfhydryl compounds are likely to be these disulfide bonds between conserved cysteine residues.
In the effort to identify target residue(s) for sulfhydryl-induced reduction, we individually mutated all conserved cysteine residues of ASIC1a, and examined the effect
of DTT on these 14 mutant channels. To our surprise, most of these mutants of ASIC1a
(11 out of 14) expressed in CHO cells did not display H+-gated current even at pH 5, which induces maximal currents in wild-type ASIC1a homomultimers. This suggests that conserved cysteine residues in the extracellular domain of ASIC1a are required for cell-surface expression or H+-dependent gating of ASIC1a by forming proper disulfide bonds. Interestingly, in all 3 mutant channels, which had H+-gated current, apparent proton (H+) sensitivity was significantly reduced suggesting that the conserved cysteine
residues are also important for normal pH sensitivity of ASIC1a. Further investigation is
required to identify target disulfide bond(s) for the reduction of ASIC1a by sulfhydryl
141 compounds using different expression systems such as Xenopus oocytes, which express
all 14 cysteine mutants of ASIC1a.
5.1.3. Pathophysiologic significance and therapeutic potentials of the sulfhydryl
regulation of ASICs
During brain ischemia, endogenous sulfhydryl compounds are released from
neurons or glial cells, and may exacerbate acid-induced neuronal death by potentiating
H+-gated currents. This is analogous to the increased extracellular concentration of
lactate during ischemic conditions, which potentiates H+-gated currents by chelating
Ca2+, which is normally present in the extracellular fluid and inhibits ASICs (Immke and
McCleskey 2001). It should be examined whether sulfhydryl modulation of ASICs
contribute to acid-induced neuronal death using both in vitro acidotoxicity assay and in
vivo animal model of stroke.
Sulfhydryl modulation of ASICs could be a potential therapeutic target for acidotoxic neuronal damage during stroke. Small allosteric molecules can be designed to bind metal inhibitory site of ASIC1a specifically and to mimic the inhibitory action of metal ions, which may be removed from ASICs by endogenous sulfhydryl compounds during brain ischemia. Considering physiologic roles of ASICs in synaptic transmission and plasticity, these small molecules may have less adverse effect when used clinically than ASIC blockers, which inhibit ASIC activation completely.
Interestingly, ebselen, an organoselenium compound, has been reported to show clinical benefits in a clinical trial for acute ischemic stroke (Ogawa et al. 1999;
Yamaguchi et al. 1998). This compound has been though to act as a scavenger of
142 oxygen-derived free radicals, which causes neuronal damage in brain ischemia (Green and Ashwood 2005). However, ebselen is also capable of oxidizing free cysteines, thus forming disulfide bonds (Herin et al. 2001). Although the mechanism of its neuroprotective effect is not completely understood, clinical benefits may come from the ability of ebselen to protect neurons from acidotoxicity by oxidizing free cysteine residues, which are already reduced by endogenous sulfhydryl compounds in brain ischemia. Therefore, sulfhydryl modulation of ASICs can be a promising target for neuroprotection in brain ischemia.
5.2. Regulation of presynaptic neurotransmitter release by ASIC1a
We observed increased neurotransmitter release probability in ASIC1 knockout mice (Cho and Askwith 2008). In rescue experiment, ASIC1a-transfected ASIC1 knockout neurons displayed smaller AMPAR EPSC and lower release probability than vector-transfected control. These results suggest that ASIC1a is necessary and sufficient to regulate synaptic transmission by modulating presynaptic release probability within the normal range. In this section, I discussed the physiologic significance of presynaptic
ASIC1a function in relation to synaptic plasticity, learning and memory. Working models on how ASIC1a can regulate presynaptic neurotransmitter release are also presented.
143 5.2.1. Presynaptic function of ASIC1a: relation to defects in learning and memory in
ASIC1 knockout mice
ASIC1 knockout mice display defects in multiple aspects of learning and memory
including impaired spatial learning, eyeblink conditioning, and fear-conditioning
(Wemmie et al. 2002). This suggests that ASIC1 is required in normal learning and
memory. However, it has not been clear how the loss of ASIC1 gene results in these
behavioral deficits. Our works provide some insights on this question.
First, we observed that release probability was higher in ASIC1 knockout neurons
compared to wild-type neurons, and that the expression of exogenous ASIC1a in ASIC1
knockout neurons reduced release probability to a level of wild-type neurons. Thus,
ASIC1a may be required for maintaining release probability within normal range in vivo.
Without ASIC1a, release probability increases abnormally, which may inhibit further
increase of synaptic strength during learning and memory process. Second, we also
observed altered short-term plasticity in ASIC1 knockout neurons, which displayed
reduced paired-pulse ratio, more frequent paired-pulse depression, and increased
depression of EPSCs during trains of high-frequency stimulation. Considering the
importance of short-term plasticity in normal behavior (Blitz et al. 2004), altered short-
term plasticity may underlie functional deficit in ASIC1 knockout mice. Finally, the
difference in release probability and short-term plasticity between wild-type and ASIC1
knockout neurons was observed only at glutamatergic synapses. ASIC1a expression in
ASIC1 knockout neurons reduced glutamatergic transmission, but did not affect
GABAergic transmission. These results suggest that presynaptic function of ASIC1a
may be specific to glutamatergic synapses. Thus, alteration of synaptic transmission only
144 at glutamatergic synapses may cause imbalance between glutamatergic (excitatory) and
GABAergic (inhibitory) synapses in ASIC 1 knockout neurons. This imbalance, which has been suggested as a potential pathophysiologic mechanism of neurodevelopmental disorders such as schizophrenia and autism, can be a cause of deficits in learning and memory in ASIC1 knockout mice.
5.2.2. Site of ASIC1a function: presynaptic or postsynaptic?
ASIC1a is localized in postsynaptic membrane of dendritic spines (Wemmie et al.
2002; Zha et al. 2006). Postsynaptic localization of ASIC1a suggests that ASIC1a may regulate synaptic transmission through postsynaptic mechanism. Indeed, ASIC1a has been reported to regulate the density of dendritic spines (Zha et al. 2006). However, we observed presynaptic alteration of synaptic transmission in ASIC1 knockout neurons whereas there was no difference in the quantal size of mEPSC, suggesting that postsynaptic mechanism was not changed in ASIC1 knockout mice. There are several examples that proteins expressed on postsynaptic side can regulate presynaptic functions
(Futai et al. 2007). Therefore, even though ASIC1a is expressed on postsynaptic membrane, it is still possible that postsynaptic ASIC1a affect presynaptic neurotransmitter release. Because it is not clear whether ASIC1a is also expressed at presynaptic terminals, it is also possible that ASIC1a may regulate neurotransmitter release from presynaptic side.
It is an important question from which side (presynaptic or postsynaptic) ASIC1a regulates neurotransmitter release. Coculture of wild-type and ASIC1 knockout neurons can be used to address this question. Hippocampal neurons isolated from wild-type and
145 ASIC1 knockout mice are plated in the same culture dish allowing synapse formation
between wild-type and ASIC1 knockout neurons. Neurotransmitter release probability can be compared between synaptic transmissions of presynaptic wild-type to postsynaptic
ASIC1 knockout and presynaptic ASIC1 knockout to postsynaptic wild-type neurons.
5.2.3. Ionotropic mechanism of postsynaptic ASIC1a: retrograde signaling
One of the popular hypotheses on ASIC function in synaptic transmission and
plasticity is that postsynaptic ASICs may be activated by protons released from synaptic
vesicles during synaptic transmission affecting the activity of postsynaptic AMPA and
NMDA receptors. However, ASIC-mediated postsynaptic currents evoked by synaptic
activity were not been observed in our system. Acute activation of ASICs by applying
pH 6.0 acidic solution or acute inhibition of ASICs by amiloride did not affect AMPAR
EPSC, either. Thus, the acute activation of ASIC by protons released from synaptic vesicles during synaptic activity is not likely to be the underlying mechanism of how
ASIC1a regulates presynaptic neurotransmitter release. However, it is still possible that
long-term sustained activation of postsynaptic ASICs by either protons or as-yet-
unidentified ligands during synaptic activity may induce signal transduction cascades in
postsynaptic neurons. Because ASIC1a homomultimers are Ca2+-permeable, Ca2+ influx through postsynaptic ASIC1a may induce the generation of nitric oxide and endocannabinoids (Fig. 2A1). These retrograde messengers may affect presynaptic neurotransmitter release. Activators or inhibitors of these signal transduction pathways could be used to test this hypothesis.
146 5.2.4. Ionotropic mechanism of presynaptic ASIC1a: signaling within presynaptic
terminals
Although it is not known whether ASIC1a is expressed at presynaptic terminals,
ASIC1a may modulate neurotransmitter release from presynaptic side. Presynaptic
ASIC1a may be activated by protons released from synaptic vesicles. In retinal ribbon
synapses, protons released during previous synaptic transmission attenuate the following
synaptic transmission by inhibiting presynaptic voltage-gated Ca2+ channels directly
(DeVries 2001; Hosoi et al. 2005; Palmer et al. 2003; Vessey et al. 2005). Ca2+- permeable ASIC1a at presynaptic terminals may be activated in similar manner, and affect neurotransmitter release through signaling mechanisms involving CaMKII, PKC, or PKA within presynaptic terminals (Fig. 2A2). In our system, it is difficult to test whether presynaptic ASIC1a is activated by released protons during synaptic activity because presynaptic currents cannot be recorded directly in conventional neuronal preparations. However, other experimental systems such as Calyx of Held synapses can be used to test this hypothesis.
5.2.5. Non-ionotropic mechanism of ASIC1a
Although most research on ASICs have been focused on H+-dependent ion- conducting (ionotropic) function, ASICs may affect synaptic transmission through non- ionotropic mechanism. This hypothesis is supported by a recent report on non-H+- dependent function of ASICs in neuronal development (Coric et al. 2008). By examining whether mutant ASIC1a with reduced pH sensitivity can rescue ASIC1 knockout
147 neurons, it can be determined whether H+-dependent ionotropic mechanism is required
for the regulation of neurotransmitter release by ASIC1a.
There are several possibilities how ASIC1a affect presynaptic function through non-ionotropic mechanisms. Presynaptic ASIC1a may interact directly with the SNARE
complex, which is required for the exocytosis of synaptic vesicles (Fig. 2B1). This
hypothesis is supported by a previous report that ASIC1a/ASIC2a/γENaC heteromeric
channels interact with syntaxin 1A, a component of SNARE complex in glioma cell line,
although it is not known whether this interaction also occur in neurons. Alternatively,
ASIC1a may also function as a modulatory subunit of presynaptic voltage-gated ion
channels, especially voltage-gated Ca2+ channels, of which activation allows Ca2+ influx at presynaptic terminals and triggers the exocytosis of synaptic vesicles (Fig. 2B2).
The modulation of presynaptic neurotransmitter release by ASICs is specific to
ASIC1a because there was no difference in presynaptic function between wild-type and
ASIC2 knockout neurons. Therefore, chimeras of ASIC1 and ASIC2 subunits can be
used to identify the domain involved in potential non-ionotropic function of ASIC1a.
Once the domain is identified, it should be examined whether recombinant peptides
corresponding to the domain rescue ASIC1 knockout neurons when applied
intracellularly or extracellularly.
148 5.3. Regulation of tonic firing of cerebellar Purkinje cells by ASICs
5.3.1. How do ASICs affect tonic firing of Purkinje cells: potential mechanisms
We found altered tonic firing of Purkinje cells in ASIC1/ASIC2 double knockout
(ASIC1/2 DKO) mice. In ASIC1/2 DKO mice, tonic firing rate was higher in either
presence or absence of endogenous synaptic activity, suggesting the alteration of intrinsic
properties of Purkinje cells. Many voltage-gated ion channels involved in the generation
and regulation of intrinsic pacemaker activity of cerebellar Purkinje cells (Raman and
Bean 1999a). Therefore, altered spontaneous firing rate may be due to changes in the
activity of voltage-gated ion channels in ASIC1/2 DKO mice (Fig. 3). A recent report raises a possibility that ASICs may regulate voltage-gated ion channels by direct interaction (Petroff et al. 2008). To test this hypothesis, it is necessary to examine whether voltage-gated ion channel activity is altered in Purkinje cells from ASIC1/2
DKO mice.
We also examined the regularity of Purkinje cell firing in the presence or absence
of endogenous synaptic transmission. When endogenous synaptic transmission was
inhibited pharmacologically, there was no difference in regularity in Purkinje cell firing
between wild-type and ASIC knockout mice, suggesting that the regularity of intrinsic
firing was not altered in ASIC knockout mice. The regularity of Purkinje cell firing was
decreased when endogenous synaptic transmission was allowed by perfusing cerebellar
slices with aCSF without inhibitors of fast synaptic transmission. This is consistent with
the previous finding that the regularity of Purkinje cell firing is modulated by endogenous
synaptic transmission (Hausser and Clark 1997). Interestingly, we did observe an
149 increased regularity in Purkinje cell firing in ASIC1/2 DKO mice compared to wild-type mice when endogenous synaptic transmission was allowed. Because synaptic activity decreases the regularity (Hausser and Clark 1997), this observation suggests that endogenous synaptic activity onto Purkinje cells may be reduced in ASIC1/2 DKO mice
(Fig. 3). This hypothesis is supported by previous reports that both ASIC1 and ASIC2 are known to regulate the spine density in hippocampal pyramidal neurons (Zha et al.
2006), although it is not clear whether spine density of Purkinje cells is also regulated by
ASICs. It is necessary to examine whether endogenous synaptic activity to Purkinje cells is altered in ASIC1/2 DKO mice by comparing the spine density and postsynaptic currents of Purkinje cells.
5.3.2. Are ASICs sufficient to modulate Purkinje cell firing?
Compared to wild-type mice, there was no significant difference in tonic firing of
Purkinje cells from ASIC1 knockout or ASIC2 knockout mice. However, tonic firing of
Purkinje cells was altered in ASIC1/2 DKO mice. These suggest that both ASIC1 and
ASIC2 subunits may affect Purkinje cell firing independently and compensate for each other in ASIC1 knockout or ASIC2 knockout mice. However, the alteration of tonic firing of ASIC1/2 DKO Purkinje cells may be due to indirect effects resulted from developmental compensation for unknown function of ASICs. Thus, it should be examined whether ASIC1 and ASIC2 are sufficient to modulate tonic firing of cerebellar
Purkinje cells. ASIC-encoding viruses could be used to deliver exogenous ASIC1 or
ASIC2 constructs to ASIC1/2 DKO Purkinje cells in vivo to examine whether transient expression of ASICs rescue tonic firing in ASIC1/2 DKO mice.
150
5.3.3. Functional significance
Purkinje cells have intrinsic pacemaker activity, which allows rapid and regular
spontaneous firing of action potentials (Raman and Bean 1999b). By firing action
potentials spontaneously, they send tonic inhibitory signal to deep cerebellar nuclei.
Proper regulation of tonic spontaneous firing of Purkinje cells is required for normal
cerebellar function (Raman et al. 1997). Thus, altered tonic firing of Purkinje cells may affect the cerebellar function in ASIC1/2 DKO mice (Fig. 3). To determine the
physiologic significance of altered tonic firing, it is necessary to perform behavioral studies comparing cerebellar function between wild-type and ASIC1/2 DKO mice.
Although ASIC1 knockout mice were reported to display impaired eyeblink conditioning
(Wemmie et al. 2002), we did not observe alteration in tonic firing of Purkinje cells in
ASIC1 knockout mice. Thus, the functional deficit in ASIC1 knockout mice may be due
to a mechanism other than altered tonic firing of Purkinje cells. Because synaptic
plasticity in Purkinje cells and neurons in deep cerebellar nuclei is implicated as
mechanisms underlying eyeblink conditioning, further investigation on synaptic transmission and plasticity in these neurons is required to explain the mechanism of
behavioral deficits in ASIC1 knockout mice.
151 A. Acute potentiation B. Long-lasting potentiation
Zn2+ Zn2+ Zn2+ SH SH S S DTT or GSH S S S S
2+ SH S S Zn S S S S SH
DTT or GSH
Zn2+
Zn2+-bound hASIC1a Zn2+-free hASIC1a Oxidized hASIC1a Reduced hASIC1a
DTT or GSH DTT or GSH / Wash pH 6.5 pH 6.5
Figure 5.1: Potentiation of ASICs by sulfhydryl compounds through two different mechanisms
152 A. Ionotropic mechanism
A1. Postsynaptic ASIC1a A2. Presynaptic ASIC1a
Na+/Ca2+
ASIC1 Retrograde messengers
ASIC1
Na+/Ca2+
B. Non-ionotropic mechanism
B1. Interaction with SNARE B2. Interaction with VGCC
Ca2+
SNARE ASIC1 ASIC1 VGCC
Figure 5.2: Regulation of presynaptic neurotransmitter release by ASIC1a : working models 153 Loss of ASIC1 and ASIC2 in ASIC1/2 DKO
Reduced spine density Altered voltage-gated in Purkinje cells? ion channel activity?
Reduced synaptic activity Increased excitability onto Purkinje cells?
Increased rate of intrinsic More regular firing of firing of Purkinje cells Purkinje cells
Deficits in cerebellar function in ASIC1/2 DKO mice?
Figure 5.3: Potential mechanisms of altered tonic firing in ASIC1/2 DKO Purkinje cells
154
BIBLIOGRAPHY
Ahmed IT. Formation constants of ternary complexes involving some metal ions, tricine, dicarboxylic amino acids, as well as N-(2-Acetamido)iminodiacetic acid and 3-amino-5- mercapto-1,2,4-triazole. J Chem Eng Data 48: 272-276, 2003.
Allen NJ and Attwell D. Modulation of ASIC channels in rat cerebellar Purkinje neurons by ischaemia-related signals. J Physiol 543: 521-529, 2002.
Almaguer-Melian W, Cruz-Aguado R, and Bergado JA. Synaptic plasticity is impaired in rats with a low glutathione content. Synapse 38: 369-374, 2000.
Alvarez de la Rosa D, Krueger SR, Kolar A, Shao D, Fitzsimonds RM, and Canessa CM. Distribution, subcellular localization and ontogeny of ASIC1 in the mammalian central nervous system. J Physiol 546: 77-87, 2003.
Alvarez de la Rosa D, Zhang P, Shao D, White F, and Canessa CM. Functional implications of the localization and activity of acid-sensitive channels in rat peripheral nervous system. Proc Natl Acad Sci U S A 99: 2326-2331, 2002.
Alvarez VA, Ridenour DA, and Sabatini BL. Distinct Structural and Ionotropic Roles of NMDA Receptors in Controlling Spine and Synapse Stability 10.1523/JNEUROSCI.0956-07.2007. J Neurosci 27: 7365-7376, 2007.
Amar M, Perin-Dureau F, and Neyton J. High-affinity Zn block in recombinant N- methyl-D-aspartate receptors with cysteine substitutions at the Q/R/N site. Biophys J 81: 107-116, 2001.
Andine P, Orwar O, Jacobson I, Sandberg M, and Hagberg H. Extracellular acidic sulfur-containing amino acids and gamma-glutamyl peptides in global ischemia: postischemic recovery of neuronal activity is paralleled by a tetrodotoxin-sensitive increase in cysteine sulfinate in the CA1 of the rat hippocampus. J Neurochem 57: 230- 236, 1991.
Andrey F, Tsintsadze T, Volkova T, Lozovaya N, and Krishtal O. Acid sensing ionic channels: modulation by redox reagents. Biochim Biophys Acta 1745: 1-6, 2005.
155
Anzai N, Deval E, Schaefer L, Friend V, Lazdunski M, and Lingueglia E. The multivalent PDZ domain-containing protein CIPP is a partner of acid-sensing ion channel 3 in sensory neurons. J Biol Chem 277: 16655-16661, 2002.
Askwith CC, Cheng C, Ikuma M, Benson C, Price MP, and Welsh MJ. Neuropeptide FF and FMRFamide potentiate acid-evoked currents from sensory neurons and proton- gated DEG/ENaC channels. Neuron 26: 133-141, 2000.
Askwith CC, Wemmie JA, Price MP, Rokhlina T, and Welsh MJ. Acid-sensing ion channel 2 (ASIC2) modulates ASIC1 H+-activated currents in hippocampal neurons. J Biol Chem 279: 18296-18305, 2004.
Babini E, Paukert M, Geisler HS, and Grunder S. Alternative splicing and interaction with di- and polyvalent cations control the dynamic range of acid-sensing ion channel 1 (ASIC1). J Biol Chem 277: 41597-41603, 2002.
Babinski K, Catarsi S, Biagini G, and Seguela P. Mammalian ASIC2a and ASIC3 subunits co-assemble into heteromeric proton-gated channels sensitive to Gd3+. J Biol Chem 275: 28519-28525, 2000.
Baron A, Deval E, Salinas M, Lingueglia E, Voilley N, and Lazdunski M. Protein kinase C stimulates the acid-sensing ion channel ASIC2a via the PDZ domain-containing protein PICK1. J Biol Chem 277: 50463-50468, 2002.
Baron A, Schaefer L, Lingueglia E, Champigny G, and Lazdunski M. Zn2+ and H+ are coactivators of acid-sensing ion channels. J Biol Chem 276: 35361-35367, 2001.
Bassilana F, Champigny G, Waldmann R, de Weille JR, Heurteaux C, and Lazdunski M. The acid-sensitive ionic channel subunit ASIC and the mammalian degenerin MDEG form a heteromultimeric H+-gated Na+ channel with novel properties. J Biol Chem 272: 28819-28822, 1997.
Bassler EL, Ngo-Anh TJ, Geisler HS, Ruppersberg JP, and Grunder S. Molecular and functional characterization of acid-sensing ion channel (ASIC) 1b. J Biol Chem 276: 33782-33787, 2001.
Beaumont V, Zhong N, Froemke RC, Ball RW, and Zucker RS. Temporal synaptic tagging by I(h) activation and actin: involvement in long-term facilitation and cAMP- induced synaptic enhancement. Neuron 33: 601-613, 2002.
Beaumont V and Zucker RS. Enhancement of synaptic transmission by cyclic AMP modulation of presynaptic Ih channels. Nat Neurosci 3: 133-141, 2000.
156 Bekkers JM and Stevens CF. Excitatory and inhibitory autaptic currents in isolated hippocampal neurons maintained in cell culture. Proc Natl Acad Sci U S A 88: 7834- 7838, 1991.
Benson CJ, Eckert SP, and McCleskey EW. Acid-evoked currents in cardiac sensory neurons: A possible mediator of myocardial ischemic sensation. Circ Res 84: 921-928, 1999.
Benson CJ and Sutherland SP. Toward an understanding of the molecules that sense myocardial ischemia. Ann N Y Acad Sci 940: 96-109, 2001.
Benson CJ, Xie J, Wemmie JA, Price MP, Henss JM, Welsh MJ, and Snyder PM. Heteromultimers of DEG/ENaC subunits form H+-gated channels in mouse sensory neurons. Proc Natl Acad Sci U S A 99: 2338-2343, 2002.
Berdiev BK, Xia J, McLean LA, Markert JM, Gillespie GY, Mapstone TB, Naren AP, Jovov B, Bubien JK, Ji HL, Fuller CM, Kirk KL, and Benos DJ. Acid-sensing ion channels in malignant gliomas. J Biol Chem 278: 15023-15034, 2003.
Bianchi L and Driscoll M. Protons at the gate: DEG/ENaC ion channels help us feel and remember. Neuron 34: 337-340, 2002.
Blitz DM, Foster KA, and Regehr WG. Short-term synaptic plasticity: a comparison of two synapses. Nat Rev Neurosci 5: 630-640, 2004.
Bolshakov KV, Essin KV, Buldakova SL, Dorofeeva NA, Skatchkov SN, Eaton MJ, Tikhonov DB, and Magazanik LG. Characterization of acid-sensitive ion channels in freshly isolated rat brain neurons. Neuroscience 110: 723-730, 2002.
Bubien JK, Ji HL, Gillespie GY, Fuller CM, Markert JM, Mapstone TB, and Benos DJ. Cation selectivity and inhibition of malignant glioma Na+ channels by Psalmotoxin 1. Am J Physiol Cell Physiol 287: C1282-1291, 2004.
Cadiou H, Studer M, Jones NG, Smith ES, Ballard A, McMahon SB, and McNaughton PA. Modulation of acid-sensing ion channel activity by nitric oxide. J Neurosci 27: 13251-13260, 2007.
Catarsi S, Babinski K, and Seguela P. Selective modulation of heteromeric ASIC proton-gated channels by neuropeptide FF. Neuropharmacology 41: 592-600, 2001.
Chai S, Li M, Lan J, Xiong ZG, Saugstad JA, and Simon RP. A kinase-anchoring protein 150 and calcineurin are involved in regulation of acid sensing ion channels ASIC1a and ASIC2a. J Biol Chem, 2007.
157 Chavis P and Westbrook G. Integrins mediate functional pre- and postsynaptic maturation at a hippocampal synapse. Nature 411: 317-321, 2001.
Chen CC, England S, Akopian AN, and Wood JN. A sensory neuron-specific, proton- gated ion channel. Proc Natl Acad Sci U S A 95: 10240-10245, 1998.
Chen CC, Zimmer A, Sun WH, Hall J, and Brownstein MJ. A role for ASIC3 in the modulation of high-intensity pain stimuli. Proc Natl Acad Sci U S A 99: 8992-8997, 2002.
Chen X, Kalbacher H, and Grunder S. Interaction of acid-sensing ion channel (ASIC) 1 with the tarantula toxin psalmotoxin 1 is state dependent. J Gen Physiol 127: 267-276, 2006a.
Chen X, Kalbacher H, and Grunder S. The tarantula toxin psalmotoxin 1 inhibits acid- sensing ion channel (ASIC) 1a by increasing its apparent H+ affinity. J Gen Physiol 126: 71-79, 2005.
Chen X, Paukert M, Kadurin I, Pusch M, and Grunder S. Strong modulation by RFamide neuropeptides of the ASIC1b/3 heteromer in competition with extracellular calcium. Neuropharmacology, 2006b.
Chi P, Greengard P, and Ryan TA. Synapsin dispersion and reclustering during synaptic activity. Nat Neurosci 4: 1187-1193, 2001.
Cho JH and Askwith CC. Potentiation of acid-sensing ion channels by sulfhydryl compounds. Am J Physiol Cell Physiol 292: C2161-2174, 2007.
Chu XP, Close N, Saugstad JA, and Xiong ZG. ASIC1a-specific modulation of acid- sensing ion channels in mouse cortical neurons by redox reagents. J Neurosci 26: 5329- 5339, 2006.
Chu XP, Miesch J, Johnson M, Root L, Zhu XM, Chen D, Simon RP, and Xiong ZG. Proton-gated channels in PC12 cells. J Neurophysiol 87: 2555-2561, 2002.
Chu XP, Wemmie JA, Wang WZ, Zhu XM, Saugstad JA, Price MP, Simon RP, and Xiong ZG. Subunit-dependent high-affinity zinc inhibition of acid-sensing ion channels. J Neurosci 24: 8678-8689, 2004.
Coric T, Passamaneck YJ, Zhang P, Di Gregorio A, and Canessa CM. Simple chordates exhibit a proton-independent function of acid-sensing ion channels. Faseb J, 2008.
158 Coric T, Zheng D, Gerstein M, and Canessa CM. Proton sensitivity of ASIC1 appeared with the rise of fishes by changes of residues in the region that follows TM1 in the ectodomain of the channel. J Physiol 568: 725-735, 2005.
Cruz-Aguado R, Almaguer-Melian W, Diaz CM, Lorigados L, and Bergado J. Behavioral and biochemical effects of glutathione depletion in the rat brain. Brain Res Bull 55: 327-333, 2001. de Weille J and Bassilana F. Dependence of the acid-sensitive ion channel, ASIC1a, on extracellular Ca(2+) ions. Brain Res 900: 277-281, 2001. de Weille JR, Bassilana F, Lazdunski M, and Waldmann R. Identification, functional expression and chromosomal localisation of a sustained human proton-gated cation channel. FEBS Lett 433: 257-260, 1998.
Deval E, Baron A, Lingueglia E, Mazarguil H, Zajac JM, and Lazdunski M. Effects of neuropeptide SF and related peptides on acid sensing ion channel 3 and sensory neuron excitability. Neuropharmacology 44: 662-671, 2003.
Deval E, Salinas M, Baron A, Lingueglia E, and Lazdunski M. ASIC2b-dependent regulation of ASIC3, an essential acid-sensing ion channel subunit in sensory neurons via the partner protein PICK-1. J Biol Chem 279: 19531-19539, 2004.
DeVries SH. Exocytosed protons feedback to suppress the Ca2+ current in mammalian cone photoreceptors. Neuron 32: 1107-1117, 2001.
Duggan A, Garcia-Anoveros J, and Corey DP. The PDZ domain protein PICK1 and the sodium channel BNaC1 interact and localize at mechanosensory terminals of dorsal root ganglion neurons and dendrites of central neurons. J Biol Chem 277: 5203-5208, 2002.
Escoubas P, De Weille JR, Lecoq A, Diochot S, Waldmann R, Champigny G, Moinier D, Menez A, and Lazdunski M. Isolation of a tarantula toxin specific for a class of proton-gated Na+ channels. J Biol Chem 275: 25116-25121, 2000.
Ettaiche M, Deval E, Cougnon M, Lazdunski M, and Voilley N. Silencing acid- sensing ion channel 1a alters cone-mediated retinal function. J Neurosci 26: 5800-5809, 2006.
Ettaiche M, Guy N, Hofman P, Lazdunski M, and Waldmann R. Acid-sensing ion channel 2 is important for retinal function and protects against light-induced retinal degeneration. J Neurosci 24: 1005-1012, 2004.
159 Firsov D, Robert-Nicoud M, Gruender S, Schild L, and Rossier BC. Mutational analysis of cysteine-rich domains of the epithelium sodium channel (ENaC). Identification of cysteines essential for channel expression at the cell surface. J Biol Chem 274: 2743-2749, 1999.
Futai K, Kim MJ, Hashikawa T, Scheiffele P, Sheng M, and Hayashi Y. Retrograde modulation of presynaptic release probability through signaling mediated by PSD-95- neuroligin. Nat Neurosci 10: 186-195, 2007.
Gao J, Duan B, Wang DG, Deng XH, Zhang GY, Xu L, and Xu TL. Coupling between NMDA receptor and acid-sensing ion channel contributes to ischemic neuronal death. Neuron 48: 635-646, 2005.
Gao J, Wu LJ, Xu L, and Xu TL. Properties of the proton-evoked currents and their modulation by Ca2+ and Zn2+ in the acutely dissociated hippocampus CA1 neurons. Brain Res 1017: 197-207, 2004.
Gao Y, Liu SS, Qiu S, Cheng W, Zheng J, and Luo JH. Fluorescence resonance energy transfer analysis of subunit assembly of the ASIC channel. Biochem Biophys Res Commun 359: 143-150, 2007.
Garcia-Anoveros J, Derfler B, Neville-Golden J, Hyman BT, and Corey DP. BNaC1 and BNaC2 constitute a new family of human neuronal sodium channels related to degenerins and epithelial sodium channels. Proc Natl Acad Sci U S A 94: 1459-1464, 1997.
Gozlan H and Ben-Ari Y. NMDA receptor redox sites: are they targets for selective neuronal protection? Trends Pharmacol Sci 16: 368-374, 1995.
Green AR and Ashwood T. Free radical trapping as a therapeutic approach to neuroprotection in stroke: experimental and clinical studies with NXY-059 and free radical scavengers. Curr Drug Targets CNS Neurol Disord 4: 109-118, 2005.
Grunder S, Geissler HS, Bassler EL, and Ruppersberg JP. A new member of acid- sensing ion channels from pituitary gland. Neuroreport 11: 1607-1611, 2000.
Hausser M and Clark BA. Tonic synaptic inhibition modulates neuronal output pattern and spatiotemporal synaptic integration. Neuron 19: 665-678, 1997.
Herin GA, Du S, and Aizenman E. The neuroprotective agent ebselen modifies NMDA receptor function via the redox modulatory site. J Neurochem 78: 1307-1314, 2001.
Hesselager M, Timmermann DB, and Ahring PK. pH Dependency and desensitization kinetics of heterologously expressed combinations of acid-sensing ion channel subunits. J Biol Chem 279: 11006-11015, 2004.
160
Hildebrand MS, de Silva MG, Klockars T, Rose E, Price M, Smith RJ, McGuirt WT, Christopoulos H, Petit C, and Dahl HH. Characterisation of DRASIC in the mouse inner ear. Hear Res 190: 149-160, 2004.
Hosoi N, Arai I, and Tachibana M. Group III metabotropic glutamate receptors and exocytosed protons inhibit L-type calcium currents in cones but not in rods. J Neurosci 25: 4062-4072, 2005.
Hovatta I, Tennant RS, Helton R, Marr RA, Singer O, Redwine JM, Ellison JA, Schadt EE, Verma IM, Lockhart DJ, and Barlow C. Glyoxalase 1 and glutathione reductase 1 regulate anxiety in mice. Nature 438: 662-666, 2005.
Hruska-Hageman AM, Wemmie JA, Price MP, and Welsh MJ. Interaction of the synaptic protein PICK1 (protein interacting with C kinase 1) with the non-voltage gated sodium channels BNC1 (brain Na+ channel 1) and ASIC (acid-sensing ion channel). Biochem J 361: 443-450, 2002.
Immke DC and McCleskey EW. Lactate enhances the acid-sensing Na+ channel on ischemia-sensing neurons. Nat Neurosci 4: 869-870, 2001.
Immke DC and McCleskey EW. Protons open acid-sensing ion channels by catalyzing relief of Ca2+ blockade. Neuron 37: 75-84, 2003.
Jasti J, Furukawa H, Gonzales EB, and Gouaux E. Structure of acid-sensing ion channel 1 at 1.9 A resolution and low pH. Nature 449: 316-323, 2007.
Jones NG, Slater R, Cadiou H, McNaughton P, and McMahon SB. Acid-induced pain and its modulation in humans. J Neurosci 24: 10974-10979, 2004.
Jones RC, 3rd, Xu L, and Gebhart GF. The mechanosensitivity of mouse colon afferent fibers and their sensitization by inflammatory mediators require transient receptor potential vanilloid 1 and acid-sensing ion channel 3. J Neurosci 25: 10981- 10989, 2005.
Jovov B, Tousson A, McMahon LL, and Benos DJ. Immunolocalization of the acid- sensing ion channel 2a in the rat cerebellum. Histochem Cell Biol 119: 437-446, 2003.
Kellenberger S and Schild L. Epithelial sodium channel/degenerin family of ion channels: a variety of functions for a shared structure. Physiol Rev 82: 735-767, 2002.
Krezel A, Lesniak W, Jezowska-Bojczuk M, Mlynarz P, Brasun J, Kozlowski H, and Bal W. Coordination of heavy metals by dithiothreitol, a commonly used thiol group protectant. J Inorg Biochem 84: 77-88, 2001.
161 Krishtal O. The ASICs: signaling molecules? Modulators? Trends Neurosci 26: 477- 483, 2003.
Krishtal OA, Osipchuk YV, Shelest TN, and Smirnoff SV. Rapid extracellular pH transients related to synaptic transmission in rat hippocampal slices. Brain Res 436: 352- 356, 1987.
Krishtal OA and Pidoplichko VI. Receptor for protons in the membrane of sensory neurons. Brain Res 214: 150-154, 1981a.
Krishtal OA and Pidoplichko VI. A receptor for protons in the membrane of sensory neurons may participate in nociception. Neuroscience 6: 2599-2601, 1981b.
Krishtal OA and Pidoplichko VI. A receptor for protons in the nerve cell membrane. Neuroscience 5: 2325-2327, 1980.
Krishtal OA and Pidoplichko VI. A "receptor" for protons in small neurons of trigeminal ganglia: possible role in nociception. Neurosci Lett 24: 243-246, 1981c.
Lalo U, Pankratov Y, North RA, and Verkhratsky A. Spontaneous autocrine release of protons activates ASIC-mediated currents in HEK293 cells. J Cell Physiol 212: 473- 480, 2007.
Landolt H, Lutz TW, Langemann H, Stauble D, Mendelowitsch A, Gratzl O, and Honegger CG. Extracellular antioxidants and amino acids in the cortex of the rat: monitoring by microdialysis of early ischemic changes. J Cereb Blood Flow Metab 12: 96-102, 1992.
Leonard AS, Yermolaieva O, Hruska-Hageman A, Askwith CC, Price MP, Wemmie JA, and Welsh MJ. cAMP-dependent protein kinase phosphorylation of the acid-sensing ion channel-1 regulates its binding to the protein interacting with C-kinase-1. Proc Natl Acad Sci U S A 100: 2029-2034, 2003.
Lingueglia E, de Weille JR, Bassilana F, Heurteaux C, Sakai H, Waldmann R, and Lazdunski M. A modulatory subunit of acid sensing ion channels in brain and dorsal root ganglion cells. J Biol Chem 272: 29778-29783, 1997.
Lipton SA, Choi YB, Takahashi H, Zhang D, Li W, Godzik A, and Bankston LA. Cysteine regulation of protein function--as exemplified by NMDA-receptor modulation. Trends Neurosci 25: 474-480, 2002.
Lisman J, Schulman H, and Cline H. The molecular basis of CaMKII function in synaptic and behavioural memory. Nat Rev Neurosci 3: 175-190, 2002.
162 Llinas R, Gruner JA, Sugimori M, McGuinness TL, and Greengard P. Regulation by synapsin I and Ca(2+)-calmodulin-dependent protein kinase II of the transmitter release in squid giant synapse. J Physiol 436: 257-282, 1991.
MacDermott AB, Role LW, and Siegelbaum SA. Presynaptic ionotropic receptors and the control of transmitter release. Annu Rev Neurosci 22: 443-485, 1999.
Mamet J, Baron A, Lazdunski M, and Voilley N. Proinflammatory mediators, stimulators of sensory neuron excitability via the expression of acid-sensing ion channels. J Neurosci 22: 10662-10670, 2002.
McCormick DA and Thompson RF. Cerebellum: essential involvement in the classically conditioned eyelid response. Science 223: 296-299, 1984.
Mennerick S, Que J, Benz A, and Zorumski CF. Passive and synaptic properties of hippocampal neurons grown in microcultures and in mass cultures. J Neurophysiol 73: 320-332, 1995.
Mennerick S and Zorumski CF. Paired-pulse modulation of fast excitatory synaptic currents in microcultures of rat hippocampal neurons. J Physiol 488 ( Pt 1): 85-101, 1995.
Mercado F, Lopez IA, Acuna D, Vega R, and Soto E. Acid-sensing ionic channels in the rat vestibular endorgans and ganglia. J Neurophysiol, 2006.
Miesenbock G, De Angelis DA, and Rothman JE. Visualizing secretion and synaptic transmission with pH-sensitive green fluorescent proteins. Nature 394: 192-195, 1998.
Mogil JS, Breese NM, Witty MF, Ritchie J, Rainville ML, Ase A, Abbadi N, Stucky CL, and Seguela P. Transgenic expression of a dominant-negative ASIC3 subunit leads to increased sensitivity to mechanical and inflammatory stimuli. J Neurosci 25: 9893- 9901, 2005.
Monyer H, Sprengel R, Schoepfer R, Herb A, Higuchi M, Lomeli H, Burnashev N, Sakmann B, and Seeburg PH. Heteromeric NMDA receptors: molecular and functional distinction of subtypes. Science 256: 1217-1221, 1992.
Moulder KL, Meeks JP, Shute AA, Hamilton CK, de Erausquin G, and Mennerick S. Plastic elimination of functional glutamate release sites by depolarization. Neuron 42: 423-435, 2004.
Ogawa A, Yoshimoto T, Kikuchi H, Sano K, Saito I, Yamaguchi T, and Yasuhara H. Ebselen in acute middle cerebral artery occlusion: a placebo-controlled, double-blind clinical trial. Cerebrovasc Dis 9: 112-118, 1999.
163 Ostrovskaya O, Moroz L, and Krishtal O. Modulatory action of RFamide-related peptides on acid-sensing ionic channels is pH dependent: the role of arginine. J Neurochem 91: 252-255, 2004.
Palmer MJ, Hull C, Vigh J, and von Gersdorff H. Synaptic cleft acidification and modulation of short-term depression by exocytosed protons in retinal bipolar cells. J Neurosci 23: 11332-11341, 2003.
Paoletti P, Ascher P, and Neyton J. High-affinity zinc inhibition of NMDA NR1-NR2A receptors. J Neurosci 17: 5711-5725, 1997.
Paukert M, Babini E, Pusch M, and Grunder S. Identification of the Ca2+ blocking site of acid-sensing ion channel (ASIC) 1: implications for channel gating. J Gen Physiol 124: 383-394, 2004.
Paukert M, Chen X, Polleichtner G, Schindelin H, and Grunder S. Candidate amino acids involved in H+ gating of acid-sensing ion channel 1a. J Biol Chem 283: 572-581, 2008.
Peng BG, Ahmad S, Chen S, Chen P, Price MP, and Lin X. Acid-sensing ion channel 2 contributes a major component to acid-evoked excitatory responses in spiral ganglion neurons and plays a role in noise susceptibility of mice. J Neurosci 24: 10167-10175, 2004.
Pignataro G, Simon RP, and Xiong ZG. Prolonged activation of ASIC1a and the time window for neuroprotection in cerebral ischaemia. Brain 130: 151-158, 2007.
Poirot O, Berta T, Decosterd I, and Kellenberger S. Distinct ASIC currents are expressed in rat putative nociceptors and are modulated by nerve injury. J Physiol, 2006.
Poirot O, Vukicevic M, Boesch A, and Kellenberger S. Selective regulation of acid- sensing ion channel 1 by serine proteases. J Biol Chem 279: 38448-38457, 2004.
Price MP, Lewin GR, McIlwrath SL, Cheng C, Xie J, Heppenstall PA, Stucky CL, Mannsfeldt AG, Brennan TJ, Drummond HA, Qiao J, Benson CJ, Tarr DE, Hrstka RF, Yang B, Williamson RA, and Welsh MJ. The mammalian sodium channel BNC1 is required for normal touch sensation. Nature 407: 1007-1011, 2000.
Price MP, McIlwrath SL, Xie J, Cheng C, Qiao J, Tarr DE, Sluka KA, Brennan TJ, Lewin GR, and Welsh MJ. The DRASIC cation channel contributes to the detection of cutaneous touch and acid stimuli in mice. Neuron 32: 1071-1083, 2001.
Price MP, Snyder PM, and Welsh MJ. Cloning and expression of a novel human brain Na+ channel. J Biol Chem 271: 7879-7882, 1996.
164 Priller C, Bauer T, Mitteregger G, Krebs B, Kretzschmar HA, and Herms J. Synapse formation and function is modulated by the amyloid precursor protein. J Neurosci 26: 7212-7221, 2006.
Raman IM and Bean BP. Ionic currents underlying spontaneous action potentials in isolated cerebellar Purkinje neurons. J Neurosci 19: 1663-1674, 1999a.
Raman IM and Bean BP. Properties of sodium currents and action potential firing in isolated cerebellar Purkinje neurons. Ann N Y Acad Sci 868: 93-96, 1999b.
Raman IM, Sprunger LK, Meisler MH, and Bean BP. Altered subthreshold sodium currents and disrupted firing patterns in Purkinje neurons of Scn8a mutant mice. Neuron 19: 881-891, 1997.
Rhee JS, Betz A, Pyott S, Reim K, Varoqueaux F, Augustin I, Hesse D, Sudhof TC, Takahashi M, Rosenmund C, and Brose N. Beta phorbol ester- and diacylglycerol- induced augmentation of transmitter release is mediated by Munc13s and not by PKCs. Cell 108: 121-133, 2002.
Rosenmund C, Clements JD, and Westbrook GL. Nonuniform probability of glutamate release at a hippocampal synapse. Science 262: 754-757, 1993.
Rosenmund C and Stevens CF. Definition of the readily releasable pool of vesicles at hippocampal synapses. Neuron 16: 1197-1207, 1996.
Sanhueza M, McIntyre CC, and Lisman JE. Reversal of synaptic memory by Ca2+/calmodulin-dependent protein kinase II inhibitor. J Neurosci 27: 5190-5199, 2007.
Saugstad JA, Roberts JA, Dong J, Zeitouni S, and Evans RJ. Analysis of the membrane topology of the acid-sensing ion channel 2a. J Biol Chem 279: 55514-55519, 2004.
Sausbier M, Hu H, Arntz C, Feil S, Kamm S, Adelsberger H, Sausbier U, Sailer CA, Feil R, Hofmann F, Korth M, Shipston MJ, Knaus HG, Wolfer DP, Pedroarena CM, Storm JF, and Ruth P. Cerebellar ataxia and Purkinje cell dysfunction caused by Ca2+-activated K+ channel deficiency. Proc Natl Acad Sci U S A 101: 9474-9478, 2004.
Sherwood TW and Askwith CC. Endogenous Arginine-Phenylalanine-Amide-related Peptides Alter Steady-state Desensitization of ASIC1a. J Biol Chem 283: 1818-1830, 2008.
Siesjo BK. Acidosis and ischemic brain damage. Neurochem Pathol 9: 31-88, 1988. Slivka A and Cohen G. Brain ischemia markedly elevates levels of the neurotoxic amino acid, cysteine. Brain Res 608: 33-37, 1993.
165 Smith ES, Cadiou H, and McNaughton PA. Arachidonic acid potentiates acid-sensing ion channels in rat sensory neurons by a direct action. Neuroscience 145: 686-698, 2007a.
Smith ES, Zhang X, Cadiou H, and McNaughton PA. Proton binding sites involved in the activation of acid-sensing ion channel ASIC2a. Neurosci Lett 426: 12-17, 2007b.
Smith RM and Martell AE. Critical Stability Constants. New York: Plenum Press, 1974.
Staruschenko A, Dorofeeva NA, Bolshakov KV, and Stockand JD. Subunit-dependent cadmium and nickel inhibition of acid-sensing ion channels. J Neurobiol 67: 97-107, 2006.
Steullet P, Neijt HC, Cuenod M, and Do KQ. Synaptic plasticity impairment and hypofunction of NMDA receptors induced by glutathione deficit: relevance to schizophrenia. Neuroscience 137: 807-819, 2006.
Sullivan JM, Traynelis SF, Chen HS, Escobar W, Heinemann SF, and Lipton SA. Identification of two cysteine residues that are required for redox modulation of the NMDA subtype of glutamate receptor. Neuron 13: 929-936, 1994.
Sutherland SP, Benson CJ, Adelman JP, and McCleskey EW. Acid-sensing ion channel 3 matches the acid-gated current in cardiac ischemia-sensing neurons. Proc Natl Acad Sci U S A 98: 711-716, 2001.
Thompson RJ, Zhou N, and MacVicar BA. Ischemia opens neuronal gap junction hemichannels. Science 312: 924-927, 2006.
Vessey JP, Stratis AK, Daniels BA, Da Silva N, Jonz MG, Lalonde MR, Baldridge WH, and Barnes S. Proton-mediated feedback inhibition of presynaptic calcium channels at the cone photoreceptor synapse. J Neurosci 25: 4108-4117, 2005.
Vila-Carriles WH, Kovacs GG, Jovov B, Zhou ZH, Pahwa AK, Colby G, Esimai O, Gillespie GY, Mapstone TB, Markert JM, Fuller CM, Bubien JK, and Benos DJ. Surface expression of ASIC2 inhibits the amiloride-sensitive current and migration of glioma cells. J Biol Chem 281: 19220-19232, 2006.
Vukicevic M and Kellenberger S. Modulatory effects of acid-sensing ion channels on action potential generation in hippocampal neurons. Am J Physiol Cell Physiol 287: C682-690, 2004.
Waldmann R. Proton-gated cation channels--neuronal acid sensors in the central and peripheral nervous system. Adv Exp Med Biol 502: 293-304, 2001.
166 Waldmann R, Bassilana F, de Weille J, Champigny G, Heurteaux C, and Lazdunski M. Molecular cloning of a non-inactivating proton-gated Na+ channel specific for sensory neurons. J Biol Chem 272: 20975-20978, 1997a.
Waldmann R, Champigny G, Bassilana F, Heurteaux C, and Lazdunski M. A proton-gated cation channel involved in acid-sensing. Nature 386: 173-177, 1997b.
Waldmann R, Champigny G, Voilley N, Lauritzen I, and Lazdunski M. The mammalian degenerin MDEG, an amiloride-sensitive cation channel activated by mutations causing neurodegeneration in Caenorhabditis elegans. J Biol Chem 271: 10433-10436, 1996.
Walter JT, Alvina K, Womack MD, Chevez C, and Khodakhah K. Decreases in the precision of Purkinje cell pacemaking cause cerebellar dysfunction and ataxia. Nat Neurosci 9: 389-397, 2006.
Wang W, Duan B, Xu H, Xu L, and Xu TL. Calcium-permeable acid-sensing ion channel is a molecular target of the neurotoxic metal ion lead. J Biol Chem 281: 2497- 2505, 2006.
Wang W, Yu Y, and Xu TL. Modulation of acid-sensing ion channels by Cu(2+) in cultured hypothalamic neurons of the rat. Neuroscience, 2007.
Waxham MN, Malenka RC, Kelly PT, and Mauk MD. Calcium/calmodulin- dependent protein kinase II regulates hippocampal synaptic transmission. Brain Res 609: 1-8, 1993.
Wemmie JA, Askwith CC, Lamani E, Cassell MD, Freeman JH, Jr., and Welsh MJ. Acid-sensing ion channel 1 is localized in brain regions with high synaptic density and contributes to fear conditioning. J Neurosci 23: 5496-5502, 2003.
Wemmie JA, Chen J, Askwith CC, Hruska-Hageman AM, Price MP, Nolan BC, Yoder PG, Lamani E, Hoshi T, Freeman JH, Jr., and Welsh MJ. The acid-activated ion channel ASIC contributes to synaptic plasticity, learning, and memory. Neuron 34: 463-477, 2002.
Wemmie JA, Coryell MW, Askwith CC, Lamani E, Leonard AS, Sigmund CD, and Welsh MJ. Overexpression of acid-sensing ion channel 1a in transgenic mice increases acquired fear-related behavior. Proc Natl Acad Sci U S A 101: 3621-3626, 2004.
Wemmie JA, Price MP, and Welsh MJ. Acid-sensing ion channels: advances, questions and therapeutic opportunities. Trends Neurosci 29: 578-586, 2006.
167 Wierda KD, Toonen RF, de Wit H, Brussaard AB, and Verhage M. Interdependence of PKC-dependent and PKC-independent pathways for presynaptic plasticity. Neuron 54: 275-290, 2007.
Wilkins ME and Smart TG. Redox modulation of GABAA receptors obscured by Zn2+ complexation. Neuropharmacology 43: 938-944, 2002.
Wu LJ, Duan B, Mei YD, Gao J, Chen JG, Zhuo M, Xu L, Wu M, and Xu TL. Characterization of acid-sensing ion channels in dorsal horn neurons of rat spinal cord. J Biol Chem 279: 43716-43724, 2004.
Xie J, Price MP, Wemmie JA, Askwith CC, and Welsh MJ. ASIC3 and ASIC1 mediate FMRFamide-related peptide enhancement of H+-gated currents in cultured dorsal root ganglion neurons. J Neurophysiol 89: 2459-2465, 2003.
Xiong ZG, Chu XP, and Simon RP. Acid sensing ion channels--novel therapeutic targets for ischemic brain injury. Front Biosci 12: 1376-1386, 2007.
Xiong ZG, Chu XP, and Simon RP. Ca2+ -permeable acid-sensing ion channels and ischemic brain injury. J Membr Biol 209: 59-68, 2006.
Xiong ZG, Zhu XM, Chu XP, Minami M, Hey J, Wei WL, MacDonald JF, Wemmie JA, Price MP, Welsh MJ, and Simon RP. Neuroprotection in ischemia: blocking calcium-permeable acid-sensing ion channels. Cell 118: 687-698, 2004.
Yagi J, Wenk HN, Naves LA, and McCleskey EW. Sustained Currents Through ASIC3 Ion Channels at the Modest pH Changes That Occur During Myocardial Ischemia. Circ Res, 2006.
Yamaguchi T, Sano K, Takakura K, Saito I, Shinohara Y, Asano T, and Yasuhara H. Ebselen in acute ischemic stroke: a placebo-controlled, double-blind clinical trial. Ebselen Study Group. Stroke 29: 12-17, 1998.
Yermolaieva O, Leonard AS, Schnizler MK, Abboud FM, and Welsh MJ. Extracellular acidosis increases neuronal cell calcium by activating acid-sensing ion channel 1a. Proc Natl Acad Sci U S A 101: 6752-6757, 2004.
Zangerle L, Cuenod M, Winterhalter KH, and Do KQ. Screening of thiol compounds: depolarization-induced release of glutathione and cysteine from rat brain slices. J Neurochem 59: 181-189, 1992.
Zha XM, Wemmie JA, Green SH, and Welsh MJ. Acid-sensing ion channel 1a is a postsynaptic proton receptor that affects the density of dendritic spines. Proc Natl Acad Sci U S A 103: 16556-16561, 2006.
168 Zhang P, Sigworth FJ, and Canessa CM. Gating of acid-sensitive ion channel-1: release of Ca2+ block vs. allosteric mechanism. J Gen Physiol 127: 109-117, 2006.
Zhong N, Beaumont V, and Zucker RS. Calcium influx through HCN channels does not contribute to cAMP-enhanced transmission. J Neurophysiol 92: 644-647, 2004.
Zucker RS and Regehr WG. Short-term synaptic plasticity. Annu Rev Physiol 64: 355- 405, 2002.
169