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Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Methods
  5. Results
  6. Discussion
  7. References
  8. Appendix
  9. Supporting Information

Acid-sensing ion channels (ASICs) are proton-gated cation channels that play important roles in the CNS including synaptic plasticity and acidosis-mediated neuronal injury. ASIC1a and ASIC2a subunits are predominant in CNS neurons, where homomultimeric and heteromultimeric channel configurations co-exist. Since ASIC1a and ASIC2a have dramatic differences in pH sensitivity, Ca2+ permeability and channel kinetics, any change in the level of individual subunits may have significant effects on the properties and functions of ASICs. Using patch-clamp recording, fluorescent Ca2+ imaging and molecular biological techniques, we show dramatic developmental changes in the properties of ASICs in mouse cortical neurons. For example, the amplitude of ASIC currents increases whereas desensitization decreases with neuronal maturation. Decreased H+ affinity and acid-evoked [Ca2+]i but increased Zn2+ potentiation were also recorded in mature neurons. RT-PCR revealed significant increases in the ratio of ASIC2/ASIC1 mRNA with neuronal maturation. Thus, contributions of ASIC1a and ASIC2a to overall ASIC-mediated responses undergo distinct developmental changes. These findings may help in understanding the precise role of ASICs in physiological and pathological conditions at different developmental stages.

Abbreviations 
ASIC

acid-sensing ion channel

PcTX 1

psalmotoxin 1

Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Methods
  5. Results
  6. Discussion
  7. References
  8. Appendix
  9. Supporting Information

Acid-sensing ion channels (ASICs) are H+-gated cation channels, which belong to the epithelial sodium channel/degenerin superfamily (Waldmann & Lazdunski, 1998; Krishtal, 2003). Molecular cloning has identified seven distinct ASIC subunits: ASIC1a, ASIC1b, ASICβ, ASIC2a, ASIC2b, ASIC3 and ASIC4 (Wemmie et al. 2006). When expressed in heterologous systems, all subunits except ASIC2b and ASIC4 can form functional homomultimeric channels with distinct electrophysiological and pharmacological properties. When co-expressed, the heteromultimeric channels may demonstrate characteristics dramatically different from their homomeric counterparts (Escoubas et al. 2000; Babinski et al. 2000; Baron et al. 2001; Benson et al. 2002; Chu et al. 2004; Hesselager et al. 2004). Therefore, alterations in the property and/or expression of individual ASIC subunits in neurons could result in dramatic change in the properties of H+-gated currents and overall acid-mediated neuronal signalling. Unlike peripheral sensory neurons where most of the cloned ASIC subunits have been detected, ASIC1a and ASIC2a are the major functional ASIC subunits in CNS neurons (Baron et al. 2002; Askwith et al. 2004; Xiong et al. 2004).

ASIC1a- and ASIC2a-mediated responses can be distinguished by their distinct pH sensitivity, pharmacology and kinetic properties. For example, ASIC1a subunits have high sensitivity to H+ with a half-maximum activation pH value (pH50) of ∼6.2 (Waldmann et al. 1997), whereas ASIC2a subunits have rather low sensitivity to H+ with a pH50 of ∼4.4 (Waldmann et al. 1999; Hesselager et al. 2004). Although both ASIC1a and ASIC2a channels are sensitive to blockade by amiloride, only homomeric ASIC1a channels are inhibited by the peptide psalmotoxin 1 (PcTX1) (Escoubas et al. 2000). While low nanomolar concentrations of Zn2+ inhibit ASIC1a-containing channels (Chu et al. 2004), high micromolar Zn2+ selectively potentiates the activities of ASIC2a-containing channels (Baron et al. 2001).

Activation of Ca2+-permeable ASIC1a channels in the brain has been shown to be involved in synaptic plasticity, learning/memory (Wemmie et al. 2002, 2003) and in acidosis-mediated neuronal injury (Yermolaieva et al. 2004; Xiong et al. 2004; Gao et al. 2005). The role of ASIC2a in the brain is less clear. A previous study has suggested that an increased ASIC2a expression may be associated with neuronal survival following global ischaemia (Johnson et al. 2001). As synaptic plasticity, learning/memory and ischaemic brain injury are functionally and neuropathologically different in developing brain, evolution of ASICs during development is of interest and potentially of therapeutic importance.

Accordingly, to understand the developmental role of ASICs in physiological and pathological processes, we determined the age-dependent changes of ASIC properties. We show dramatic differences in pH sensitivity, Ca2+ permeability, pharmacology and kinetics of ASICs in mouse cortical neurons at different culture days and in acutely dissociated neurons from different ages of mice. Although the exact mechanism underlying these changes is not clear, our data suggest that alterations in the level of ASIC1a and ASIC2a expression might be partially responsible.

Methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Methods
  5. Results
  6. Discussion
  7. References
  8. Appendix
  9. Supporting Information

Ethical approval

The protocol for the use of mice for neuronal culture and acute dissociation of neurons was reviewed and approved by the Institutional Animal Care and Use Committee of Legacy Clinical Research and Technology Center.

Primary neuronal cultures

Mouse cortical neurons were cultured as described previously (Chu et al. 2004). Briefly, time-pregnant Swiss mice (embryonic day 16) were anaesthetized with halothane followed by cervical dislocation. Brains of fetuses were removed rapidly and placed in Ca2+/Mg2+-free ice-cold PBS. Cerebral cortices were dissected under a dissection microscope and incubated with 0.05% trypsin–EDTA for 10 min at 37°C, followed by trituration with fire-polished glass pipettes. Cells were counted and plated in poly-l-ornithine-coated culture dishes or 24-well plates at a density of 1 × 106 cells per 35 mm diameter dish or 2 × 105 cells per well. Neurons were cultured with Neurobasal medium supplemented with B27 and maintained at 37°C in a humidified 5% CO2 atmosphere incubator. Cultures were fed twice a week. Neurons at the following culture stages were used for the experiments: day 7, 14, 21 and 28 or above. The neurons with large pyramidal-shaped cell bodies and thick apical dendritic processes were chosen for the study (Supplemental Fig. S1).

Acute isolation of mouse cortical neurons

Acute dissociation of mouse cortical neurons was performed as described previously (Xiong et al. 1999; Li et al. 2004), with minor modifications. Briefly, Swiss mice from postnatal day 1 to day 28 were anaesthetized with halothane. Cortical tissues were dissected and incubated in oxygenated ice-cold dissociation solution containing (in mm): Na2SO4 82, K2SO4 30, MgCl2 5, Hepes 10, glucose 10, pH 7.3. Transverse cortical slices (400–500 μm) were cut with a microtome (Leica VT 1000) followed by incubation in dissociation solution containing 3.5 mg ml−1 papain at room temperature for 20–30 min. Slices were then washed three times and incubated in enzyme-free solution for at least 30 min before mechanical dissociation. For dissociation, slices were triturated using a series of fire-polished Pasteur pipettes with decreasing tip diameters. Recording began 15 min after the mechanical dissociation.

Whole-cell patch-clamp recording

ASIC currents were recorded with the conventional whole-cell patch-clamp technique as described previously (Xiong et al. 2004). Unless otherwise specified, cells were voltage-clamped at −60 mV. Patch pipettes were pulled from borosilicate glass (1.5 mm diameter; WPI, Sarasota, FL, USA) on a two-stage puller (PP83, Narishige, Tokyo, Japan). Pipettes had a resistance of 2–4 MΩ when filled with the intracellular solution (see below). Membrane capacitance was recorded for each neuron as a measure of cell size. For rapid changes of extracellular solutions, a multi-barrel perfusion system (SF-77, Warner Instruments, Hamden, CT, USA) was used. During each experiment, a voltage step of −10 mV from the holding potential was applied periodically to monitor cell capacitance and access resistance. Recordings in which access resistance or capacitance changed by more than 10% during the experiment were excluded for data analysis (Xiong et al. 1998; Chu et al. 2004). Whenever possible, neurons with a pyramidal shape were selected for recordings.

Solutions and chemicals

Standard extracellular solution (ECF) contained (in mm): 140 NaCl, 5.4 KCl, 2 CaCl2, 1 MgCl2, 20 Hepes, 10 glucose. The Na+-, K+- and Mg+-free high-Ca2+ solutions contained (in mm): 140 choline chloride, 10 CaCl2, 25 Hepes, 10 glucose. pH was adjusted to 7.4 or the values indicated, with NaOH/HCl, and the osmolarity was adjusted to 320–335 mosmol l−1. For solutions with pH ≤ 6.0, Hepes was replaced by Mes for more reliable pH buffering (Bassler et al. 2001; Chu et al. 2002). Intracellular solution contained (in mm): 140 CsF, 2 TEACl, 5 EGTA, 10 Hepes, 1 CaCl2, 4 MgCl2; pH 7.3 adjusted with CsOH/HCl, and the osmolarity was adjusted to 290–300 mosmol l−1.

In general, acidic solutions were applied to neurons at 2 min intervals, to allow for complete recovery of the ASICs from desensitization. To ensure high quality voltage-clamp, only the recordings with an access resistance of less than 10 MΩ and a leak current less than 100 pA at −60 mV were included for data analysis. ZnCl2 and amiloride were purchased from Sigma. PcTX1 was purchased from Spider Pharm Inc. (Yarnell, AZ, USA).

Ca2+ imaging

Fura-2 fluorescent Ca2+ imaging was performed as described previously (Xiong et al. 2004). Cortical neurons grown on 25 × 25 mm glass coverslips were washed three times with ECF and incubated with 5 mm fura-2 AM for ∼40 min at room temperature. Neurons were then washed three times and incubated in normal ECF for 30 min. Coverslips with fura-2-loaded neurons were transferred to a perfusion chamber on the stage of an inverted microscope (Nikon TE300). Cells were illuminated using a xenon lamp (75 W) and observed with a 40× UV fluor oil-immersion objective lens. Video images were obtained using a cooled CCD camera (Sensys KAF 1401, Photometrics, Tucson, AZ, USA). Digitized images were acquired, stored and analysed in a PC controlled by Axon Imaging Workbench software (AIW2.1, Axon Instruments). The shutter and filter wheel (Lambda 10-2, Sutter Instrument, Novato, CA, USA) were also controlled by AIW to allow timed illumination of cells at 340 and 380 nm excitation wavelengths. Fura-2 fluorescence was detected at an emission wavelength of 510 nm. Ratio images of 340/380 nm were analysed by averaging pixel ratio values in circumscribed regions of cells in the field of view. The values were exported from AIW to SigmaPlot for further analysis and plotting.

Multiplex RT-PCR

Total RNAs were extracted from cortical tissues of mice at different ages with an RNeasy kit (Qiagen) according to manufacturer's instruction. cDNAs were then synthesized from 0.2 μg total RNA in 20 μl volume by reverse transcription using oligo(dT)15 and SuperscriptII (Invitrogen) according to the manufacturer's protocol. A forward oligonucleotide primer (5′-CACATGCCAGGGGATGCCCC-3′) was synthesized for both ASIC1 and ASIC2, while the reverse primer was specific for individual ASICs (ASIC1; 5′-AGCCGGTGCTTAATGACCTC-3′, ASIC2; 5′-GTGTCAGCAGGCAATCTCCTCC-3′). PCR reactions were performed using 0.25 μl of cDNAs as templates in 1× PCR reaction buffer, 0.2 mm each dNTP, 0.2 ml DNA polymerase mix (Advantage cDNA Polymerase Mix, BD Biosciences), and 333 nm each primer in a 10 μl reaction volume. The PCR amplification consisted of denaturation at 94°C for 3 min, 27 cycles of denaturation at 94°C for 30 s, annealing at 61°C for 15 s, and extension at 72°C for 30 s. PCR products were separated by electrophoresis on a 1.5% agarose gel, detected using ethidium bromide, and sequenced. The detected signals were quantified by laser densitometry, and the relative ratios were calculated by dividing the density of the ASIC2 band by that of the ASIC1 band.

Data analysis

Electrophysiology data were acquired using an Axopatch 200B amplifier with pCLAMP 8.1 software (Axon Instruments). Data were analysed using Clampfit (Axon Instruments). The pH50 values for H+ dose–response and steady-state inactivation curves were fitted using the following equation (Wang et al. 2006): I=a/(1+(C50/pH)n), where a is the normalized amplitude of the ASIC current, C50 is the pH at which a half-maximal response occurs, and n is the Hill coefficient. The time constant of desensitization (τ) and recovery from desensitization were determined using a mono-exponential fit.

Relative permeability ratio of Ca2+/Cs+ was determined using the constant field equation (Jia et al. 1996):

  • image

where E is the reversal potential, and F, R and T have their conventional meaning. PCa and PCs represent the permeability coefficients of Ca2+ and Cs+.

All data are reported as mean ±s.e.m. One-way ANOVA and Student's t test were used where appropriate to examine the statistical significance of the difference between groups of data. The criterion for significance was set at P < 0.05.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Methods
  5. Results
  6. Discussion
  7. References
  8. Appendix
  9. Supporting Information

Increase in the amplitude and density of ASIC currents with neuronal maturation

As reported previously (Varming, 1999; Xiong et al. 2004), transient ASIC currents were activated in all cortical neurons when extracellular pH (pHo) is reduced from 7.4 to below 7.0. To determine whether there is a change in the ASIC response with neuronal maturation, we first compared the peak amplitude of the acid-activated current by a pH drop from 7.4 to 6.0 in neurons at different days in culture. To be consistent, the amplitude of ASIC current recorded at 2 min after the establishment of whole-cell configuration was compared among different groups of neurons. As shown in Fig. 1 and Table 1A, the amplitude of the ASIC currents increased gradually from day 7 to day 28 (−378.10 ± 40.24 pA at day 7, −1093.98 ± 144.11 pA at day 14, −1556.42 ± 136.03 pA at day 21, and −1798.40 ± 160.98 pA at day 28, n= 43–48, P < 0.01 between day 7 vs. day 14, day 21 and day 28, one-way ANOVA with Bonferroni's post hoc tests). To determine whether the increase in the current amplitude was simply due to an increase in the size of neurons, or due to a change in the current density, membrane capacitance was recorded as a measure of the cell size. The current density was then calculated by dividing the peak current amplitude by the value of cell capacitance. The cell capacitance gradually increased from day 7 to day 28 (38.40 ± 1.10 pF at day 7, 62.39 ± 3.07 pF at day 14, 74.77 ± 4.21 pF at day 21, and 87.00 ± 4.41 pF at day 28, n= 43–48, data not shown), indicating an increase of cell size with neuronal maturation. Similar to the current amplitude, the density of the ASIC currents also increased with neuronal maturation (−9.69 ± 0.99 pA pF−1 at day 7; −17.30 ± 1.89 pA pF−1 at day 14; −20.78 ± 1.43 pA pF−1 at day 21; and −22.30 ± 1.85 pA pF−1 at day 28). The differences between day 7 vs. day 14, day 21 and day 28 were statistically significant (P < 0.01, n= 43–48, one-way ANOVA with Bonferroni's post hoc tests, Fig. 1C).

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Figure 1. Developmental change in the amplitude and density of the ASIC currents in cultured mouse cortical neurons A, representative traces showing the ASIC current activated by pH drop from 7.4 to 6.0 recorded in neurons at different culture days. The amplitude of the ASIC current increased gradually from day 7 to day 28. B and C, the summary data showing the change of the amplitude (B) and the density (C) of the ASIC current in cultured cortical neurons (n= 43–48, **P < 0.01 between day 7 vs. day 14, 21 and 28, one-way ANOVA with Bonferroni's post hoc tests). D and E, the summary data showing the change of the amplitude and the density of the ASIC currents in acute dissociated cortical neurons (n= 25–38, **P < 0.01 for day 1 vs. day 7, 14 and 28, one-way ANOVA with Bonferroni's post hoc tests).

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Table 1.  Developmental changes in the properties of ASIC currents in cultured and acutely dissociated mouse cortical neurons
A. Cultured mouse cortical neurons
Days in cultureAmplitude (pA, at pH 6.0)Half-maximum activation (pH50)Decay time constant (τd, s, at pH 6.0)Recovery time constant (τrec, s)Half-maximum inactivation (pH50)
Day 7 −378.10 ± 40.24 (n= 48)6.24 ± 0.04 (n= 8)1.18 ± 0.09 (n= 26)5.40 ± 1.04 (n= 7)7.27 ± 0.02 (n= 11)
Day 14−1093.98 ± 144.11 (n= 44)**6.00 ± 0.07 (n= 7)*1.79 ± 0.12 (n= 26)*3.30 ± 0.81 (n= 13)*7.23 ± 0.04 (n= 10)*
Day 21−1556.42 ± 136.03 (n= 43)**†5.75 ± 0.10 (n= 6)**2.26 ± 0.24 (n= 31)**†2.02 ± 0.63 (n= 12)**†7.05 ± 0.06 (n= 11)**†
Day 28−1798.40 ± 160.98 (n= 45)**‡5.74 ± 0.08 (n= 7)**†2.52 ± 0.19 (n= 33)**‡1.23 ± 0.40 (n= 9)**†7.03 ± 0.04 (n= 10)**‡
Data are presented as mean ±s.e.m.*P < 0.05 compared with day 7; **P < 0.01 compared with day 7; †P < 0.05 compared with day 14; ‡P < 0.01 compared with day 14; one-way ANOVA with Bonferroni's post hoc tests
B. Acute dissociated mouse cortical neurons
Postnatal dayAmplitude (pA, at pH 6.0)Half-maximum activation (pH50)Recovery time constant (τrec, s)Half-maximum inactivation (pH50)
  1. *P < 0.05 compared with day 1; **P < 0.01 compared with day 1; †P < 0.05 compared with day 7; ‡P < 0.01 compared with day 7; one-way ANOVA with Bonferroni's post hoc tests

Day 1 −546.89 ± 71.73 (n= 38)6.51 ± 0.14 (n= 7)4.15 ± 0.99 (n= 7)7.27 ± 0.04 (n= 7)
Day 7−1223.44 ± 156.09 (n= 31)**6.33 ± 0.05 (n= 7)*2.03 ± 0.64 (n= 7)**7.12 ± 0.03 (n= 7)*
Day 14−1317.92 ± 149.05 (n= 30)**5.89 ± 0.09 (n= 5)**‡1.05 ± 0.17 (n= 7)**7.11 ± 0.02 (n= 13)**
Day 28−1387.64 ± 201.11 (n= 25)**5.80 ± 0.09 (n= 6)**‡0.78 ± 0.15 (n= 7)**†7.08 ± 0.04 (n= 7)**

The increases in current amplitude and density were also observed in acutely dissociated mouse cortical neurons (Fig. 1D and E, Table 1B). For example, the amplitude of currents increased from −546.89 ± 71.73 pA at day 1 to −1387.64 ± 201.11 pA at day 28, and the density of currents increased from −17.62 ± 2.50 pA at day 1 to −57.83 ± 5.34 pA at day 28 (n= 25 and 38, P < 0.01).

Decrease in the sensitivity of ASICs to H+ with neuronal maturation

We next determined whether the maturation process of neurons increases the sensitivity of ASICs to H+, which might also explain the increase of the current amplitude. For this reason, H+ dose–response relationships were constructed in neurons at different days in culture. To reduce the possibility that initial current run-down or tachyphylaxis with acid applications (Chen & Grunder, 2007) might interfere with accurate construction of the H+ dose–response curve, ASIC currents activated by different test pH values were studied at ∼10 min following the establishment of whole-cell configuration when stable amplitude of the ASIC current activated at pH 6.0 had been recorded. To our surprise, there was an apparent shift in the H+ dose–response curve towards more acidic pH values from day 7 to day 28, indicating a decrease, rather than an increase, in the sensitivity of ASICs to H+ (Fig. 2A and B). For example, the pH50 value was shifted from 6.24 ± 0.04 to 5.74 ± 0.08 from day 7 to day 28 (n= 7 and 8, P < 0.01, Table 1A). There was no significant change in Hill coefficient (1.19 ± 0.04 vs. 1.03 ± 0.03 at day 7 and 28, respectively). A decrease in the sensitivity of ASICs to H+ with neuronal maturation was also observed in acutely dissociated mouse cortical neurons (Fig. 2C and D, Table 1B). Since homomeric ASIC1a channels have high sensitivity to H+ (pH50 around 6.2), whereas homomeric ASIC2a channels have the least sensitivity to H+ (pH50 around 4.4) (Waldmann et al. 1999; Hesselager et al. 2004), the shift of pH50 to more acidic values might suggest an increase in the expression of the ASIC2a subunit, or in the relative ratio of ASIC2a/ASIC1a, with neuronal maturation.

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Figure 2. pH-dependent activation of the ASIC currents in cultured and dissociated mouse cortical neurons A, representative current traces showing pH-dependent activation of the ASIC currents in cultured cortical neurons at day 7 and day 28. B, summary data showing H+ dose–response curves for ASICs recorded at day 7 and day 28 in culture (n= 6–8). C, representative current traces showing pH-dependent activation of the ASIC currents in acutely dissociated mouse cortical neurons at postnatal day 1 to day 28. D, summary data showing H+ dose–response curves for ASICs in acutely dissociated mouse cortical neurons recorded at postnatal day 1 to day 28 (n= 5–7).

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Decrease in desensitization of the ASIC currents with neuronal maturation

The desensitization of ASICs is another important parameter for these channels since it determines how long the channels can stay open in the continuous presence of acidosis. Therefore, we next determined whether the desensitization properties of ASICs also change with neuronal maturation. The desensitization time constant of ASICs recorded at different culture days was measured by fitting the decay phase of the current with a single exponential function (Chu et al. 2002; Xiong et al. 2004). As shown in Fig. 3A and B, there is a gradual decrease in the desensitization of ASICs with neuronal maturation. For example, the decay time constant (τd) of the ASIC currents activated at pH 6.0 was 1.18 ± 0.09 s at day 7, but it increased to 2.52 ± 0.19 s at day 28 (P < 0.01 n= 26 and 33, Table 1A). Since the currents mediated by different ASIC subunits have dramatic difference in their desensitization kinetics (Askwith et al. 2004; Hesselager et al. 2004), the increase in the decay time constant of the ASIC currents in mature neurons might also suggest changes in the expression of ASIC subunits. Of course, other factors contributing to the changes of ASIC desensitization cannot be ruled out.

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Figure 3. Developmental change in the rate of desensitization of the ASIC currents in cultured mouse cortical neurons A, representative traces showing a gradual decrease in the rate of desensitization of ASIC currents recorded in neurons at day 7 to day 28 in culture. B, summary data showing the change of the rate of desensitization time constant of the ASIC current recorded in neurons at day 7 to day 28 in culture (n= 26–33). *P < 0.05, **P < 0.01.

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Increased rate of recovery of ASICs from desensitization in mature neurons

A potential change in the rate of recovery of ASICs from desensitization was then studied as previously described (Wang et al. 2006). This parameter is important as it defines how fast the channels can recover from desensitization and be activated by repeated acid pulses, a paradigm expected during intense excitatory neurotransmission. ASIC currents were activated by pairs of acid exposures (from 7.4 to 6.0) with increasing time intervals between the end of the first acid exposure (15 s in duration) and the beginning of the second acid exposure (5 s in duration). The relative amplitude of the second current to the first was then plotted against the time intervals, and the recovery time constant derived from an exponential fit (Wang et al. 2006). As shown in Fig. 4A and B, the recovery of the ASIC currents from desensitization became faster with neuronal maturation. For example, the recovery time constant (τrec) of the ASIC currents decreased from 5.40 ± 1.04 s at day 7 to 1.23 ± 0.40 s at day 28 (P < 0.01, n= 7 and 9, Fig. 4C and D, Table 1A).

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Figure 4. Developmental change in the recovery of ASIC currents from desensitization A and B, representative traces showing time-dependent recovery of the ASIC current from desensitization. The currents were recorded in cultured mouse cortical neurons at either day 7 (A) or day 28 (B) in culture. Neurons were first exposed to acidic solution (pH 6.0) for 15 s in order to achieve a complete desensitization of the ASIC current. At different time intervals following the end of the acid pulse, ASIC current was activated again by a second acid pulse (5 s in duration). C, summary data showing the relative amplitude of the ASIC current in cultured neurons (the second amplitude divided by the first one) plotted against the time intervals. D, summary bar graph showing developmental change in the time constant for recovery of ASICs from desensitization recorded in cultured mouse cortical neurons. The time constants were derived by single exponential fit to the curves in C. n= 7–13, *P < 0.05, **P < 0.01. E, summary data showing the relative amplitude of the ASIC current in acutely dissociated neurons plotted against the time intervals. F, summary bar graph showing developmental change in the time constant for recovery of ASICs from desensitization recorded in acutely dissociated mouse cortical neurons. The time constants were derived by single exponential fit to the curves in E. n= 7, **P < 0.01.

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An increase in the rate of recovery from desensitization with neuronal development was also observed in acutely dissociated mouse cortical neurons. The τrec was 4.15 ± 0.99 s in cortical neurons dissociated from postnatal day 1 mice (n= 7), but it was 0.78 ± 0.15 s in neurons from day 28 mice (n= 7, P < 0.01, Fig. 4E and F, Table 1B).

Decreased steady-state inactivation of ASICs in mature neurons

A minor and sustained pH drop (e.g. 7.4 to 7.2), insufficient to activate ASICs itself, can induce substantial inactivation of these channels. This steady-state inactivation is an important parameter for ASICs since it determines how many channels are available for activation following a minor acidosis. Therefore, we next compared the steady-state inactivation of ASICs in neurons at different culture stages. Neurons were exposed to extracellular solutions at different conditioning pH values ranging from 8.0 to 6.8 for 2 min before lowering pH to 6.0. The amplitude of the ASIC currents recorded with different conditioning pHs was normalized to that recorded with the conditioning pH of 8.0 (where there is no inactivation) and then plotted against the conditioning pH values to generate the steady-state inactivation curve (Wang et al. 2006). As shownin Fig. 5A and B, the steady-state inactivation curvewas shifted toward more acidic pH values from day 7 to day 28, indicating a reduced inactivation of ASICs in mature neurons. The pH50 for steady-state inactivation was 7.27 ± 0.02 at day 7, whereas it was 7.03 ± 0.04 at day 28 (n= 10 and 11, P < 0.01, Table 1A).

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Figure 5. Age-dependent shift of steady-state inactivation of the ASIC currents A, representative traces showing the ASIC currents in cultured mouse cortical neurons activated at pH 6.0 from various conditioning pH. B, summary data showing the steady-state inactivation curve for ASICs in cultured mouse cortical neurons at different days in culture. n= 10–11. C, representative traces showing the ASIC currents in acutely dissociated mouse cortical neurons activated at pH 6.0 from various conditioning pH. D, summary data showing the steady-state inactivation curve for ASICs in acutely dissociated mouse cortical neurons at different postnatal days. n= 7–13.

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A similar shift in the steady-state inactivation curve was recorded in acutely dissociated neurons (Fig. 5C and D). The pH50 value for steady-state inactivation was 7.27 ± 0.04 at day 1, whereas it was 7.08 ± 0.04 at day 28 (n= 7, P < 0.01, Table 1B).

Decrease in Ca2+ permeability of ASICs with neuronal maturation

In addition to Na+, homomeric ASIC1a channels also conduct Ca2+ (Waldmann et al. 1997; Yermolaieva et al. 2004; Xiong et al. 2004). The entry of Ca2+ through these channels plays an important role in acidosis-mediated neuronal injury (Yermolaieva et al. 2004; Xiong et al. 2004). For this reason, we determined whether the Ca2+ permeability of ASICs in mouse cortical neurons also changes with neuronal maturation. With 10 mm Ca2+ as the only charge carrier in the extracellular solution, detectable inward currents can be recorded in cultured neurons upon lowering pHo to 6.0, indicating Ca2+ permeability of ASICs (Fig. 6A). This acid-activated, Ca2+-mediated inward currents can be inhibited by amiloride and PcTX1, confirming that the acid-evoked Ca2+ current was carried by homomeric ASIC1a channels (Fig. 6A). To quantify the change of Ca2+ permeability of ASICs with neuronal maturation, the reversal potential of acid-activated, Ca2+-mediated current was measured at different culture days. As shown in Fig. 6B (right), the reversal potential of the acid-activated Ca2+ current shifted to more negative values with neuronal maturation, indicating a decrease in the relative Ca2+ permeability of ASICs in mature neurons. The reversal potential of the Ca2+-mediated currents was −12.31 ± 1.03, −18.82 ± 1.22, −23.17 ± 1.48 and −23.76 ± 0.24 mV at culture day 7, 14, 21 and 28, respectively (n= 5). The relative permeability of Ca2+/Cs+ was 4.29 ± 0.35, 3.03 ± 0.16, 2.24 ± 0.14 and 1.95 ± 0.18 for day 7, 14, 21 and 28, respectively. A similar shift in the reversal potential to more negative values was observed in acutely dissociated mouse cortical neurons (Fig. 6C, right panel). The reversal potential was −0.72 ± 0.40, −5.64 ± 0.75, −16.51 ± 1.33 and −18.28 ± 1.33 mV at postnatal day 1, 7, 14 and 28, respectively (n= 5). The relative permeability of Ca2+/Cs+ was 8.18 ± 0.20, 5.95 ± 0.26, 3.45 ± 0.24 and 3.2 ± 0.23 for day 1, 7, 14 and 28, respectively.

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Figure 6. Age-dependent change in the reversal potential of acid-activated, Ca2+-mediated currents in cultured and acutely dissociated mouse cortical neurons A, representative traces showing blockade of Ca2+-mediated current by amiloride and PcTX1. The currents were recorded in the presence of 10 mm Ca2+ but absence of Na+ and K+ ions in the extracellular solution B, representative traces showing the acid-activated Ca2+-mediated currents in cultured mouse cortical neurons recorded at different membrane potentials indicated (left). Right panel showing the current–voltage relationship (I–V curve) of the acid-activated, Ca2+-mediated current in cultured mouse cortical neurons at different cultured days. The reversal potentials of the acid-activated Ca2+ current were −12.09 ± 1.10 mV at day 7, −18.69 ± 5.06 mV at day 14, −23.04 ± 3.61 mV at day 21, and −23.74 ± 4.56 mV at day 28, n= 5, P < 0.05 for day 7 vs. day 21, and P < 0.01 for day 7 vs. day 28 n= 5. C, representative traces showing the acid-activated Ca2+-mediated currents in acutely dissociated mouse cortical neurons recorded at different membrane potentials (left). Right panel showing the current–voltage relationship (I–V curve) of the acid-activated, Ca2+-mediated current in dissociated mouse cortical neurons at different postnatal days. The reversal potential was −0.72 ± 3.43 mV at day 1, −5.67 ± 2.56 mV at day 7, −16.66 ± 2.80 mV at day 14, and −17.29 ± 1.82 at day 28, respectively. P < 0.01 for day 1 vs. day 14 or day 28, n= 5–8. A junction potential of −7 mV has been corrected.

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Ca2+ imaging was performed, in the presence of the blockers of glutamate receptors and voltage-gated Ca2+ channels, to further demonstrate changes of Ca2+ permeability of ASICs with neuronal maturation. As reported previously (Xiong et al. 2004; Wang et al. 2006), reduction of pHo from 7.4 to 6.0 induced a dramatic increase of [Ca2+]i in the majority of cultured mouse cortical neurons independent of activation of glutamate receptors and voltage-gated Ca2+ channels. The amplitude of this acid-induced increase of [Ca2+]i decreased gradually in cultured cortical neurons from day 7 to day 28 (Fig. 7).

image

Figure 7. Developmental changes in acid-induced [Ca2+]i increase A and B, representative images and 340/380 ratios showing the amplitude of acid-induced increase of [Ca2+]i decreased gradually in cultured mouse cortical neurons from day 7 to day 28. Neurons were bathed in normal ECF containing 2 mm CaCl2 with blockers for voltage-gated Ca2+ channels (5 μm nimodipine) and glutamate receptors (10 μm MK801 and 20 μm CNQX). C, summary data showing gradual decrease of acid-induced increase of [Ca2+]i with neuronal maturation. The 340/380 ratio was 7.2 ± 3.4, 2.6 ± 0.5, 1.3 ± 0.3 and 0.8 ± 0.4 at day 7, day 14, day 21 and day 28, respectively (n= 8–25, *P < 0.05 for day 7 vs. day 14, **P < 0.01 for day 7 vs. day 21 and day 28).

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Increased sensitivity of ASICs to Zn2+ potentiation with neuronal maturation

Zinc, an endogenous divalent cation, modulates the activity of ASICs in a subunit-dependent manner (Baron et al. 2001; Chu et al. 2004). At high micromolar concentrations (e.g. 100–300 μm), Zn2+ potentiates the ASIC currents mediated by ASIC2a-containing channels (Baron et al. 2001), whereas at nanomolar concentrations, Zn2+ inhibits the ASIC currents mediated by ASIC1a-containing channels (Chu et al. 2004). Next, we examined the potential change in the potentiation of ASIC currents by 200 μm Zn2+ with neuronal maturation. As shown in Fig. 8, the potentiation of ASIC currents by Zn2+ increased from 25.15 ± 12.16% at day 7 to 56.40 ± 11.40% at day 28 (n= 12 and 14, P < 0.05, Fig. 8A and B), suggesting an increased ASIC2a expression with neuronal maturation. Similar, in acutely dissociated neurons, the potentiation of ASIC currents by 200 μm Zn2+ increased from 24.87 ± 12.25% (n= 7) at postnatal day 1 to 59.85 ± 6.33 (n= 13) at day 28 (P < 0.05, Fig. 8C and D).

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Figure 8. Age-dependent potentiation of the ASIC current by ZnCl2 A and C, representative current traces showing potentiation of the ASIC current by 200 μm ZnCl2 in cultured (A) and acutely dissociated (C) mouse cortical neurons. B and D, summary data showing age-dependent increase in the sensitivity of ASIC currents to zinc potentiation in culture (B) and acutely dissociated (D) mouse cortical neurons. The percentage potentiation of the ASIC current by zinc increased from 25.15 ± 12.16 at day 7 to 56.40 ± 11.40 at day 28 in cultured neurons (n= 12 and 14, *P < 0.05), and from 24.87 ± 12.25 at postnatal day 1 to 59.85 ± 6.33 at day 28 in acutely dissociated neurons (n= 7 and 13, *P < 0.05).

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Decreased sensitivity of the ASIC currents to PcTX1 inhibition in mature neurons

PcTX1 specifically inhibits homomeric ASIC1a channels with little effect on other configurations of ASICs (Escoubas et al. 2000; Xiong et al. 2004; Chen et al. 2005). To further evaluate the changes in the relative abundance of the ASIC2a/ASIC1a channels with neuronal maturation, we examined the sensitivity of the ASIC currents to PcTX1 inhibition at culture day 7 and day 28. After recording the stable ASIC currents activated by pH drops from 7.4 to 6.5, PcTX1 venom at 200,000 times dilution (or 100 ng ml−1 total venom protein) was added to the bath solution. Previous studies have shown that, at this concentration, it specifically inhibits the homomeric ASIC1a current without any effect on the current mediated by other configurations of ASICs or ligand-gated channels (Xiong et al. 2004). As shown in Fig. 9A and B, the percentage inhibition of the ASIC currents by PcTX1 decreased from 88.71 ± 3.68 at day 7 to 63.52 ± 6.81 at day 28 (n= 12, P < 0.01). Similar to cultured neurons, reduction in the sensitivity of the ASIC currents to PcTX1 inhibition was seen in acutely dissociated neurons. The percentage inhibition of the ASIC currents by PcTX1 decreased from 90.27 ± 5.64 at day 1 to 32.68 ± 8.76 at day 28 (n= 5–6, P < 0.01, Fig. 9C and D). The decrease in the sensitivity of the ASIC currents to PcTX1 suggests a relative reduction of homomeric ASIC1a channels in mature neurons.

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Figure 9. Age-dependent decrease in the sensitivity of ASIC currents to PcTX1 inhibition A and C, representative current traces showing the inhibition of ASIC currents by PcTX1 in cultured (A) and acutely dissociated (C) mouse cortical neurons. B and D, summary data showing age-dependent decrease in the sensitivity of ASIC currents to PcTX1 in cultured (B) and acutely dissociated (D) mouse cortical neurons. The percentage inhibition of ASIC currents by PcTX1 decreased from 88.71 ± 3.68 at day 7 to 63.52 ± 6.81 at day 28 in cultured neurons (n= 12, **P < 0.01), and from 90.27 ± 5.64 at postnatal day 1 to 32.68 ± 8.76 at day 28 in acutely dissociated neurons (n= 5–6, **P < 0.01).

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Increase in relative ratio of ASIC2/ASIC1 mRNA with neuronal maturation

Although various factors might have contributed to the changes of properties of ASICs with neuronal maturation, our electrophysiological and pharmacological data all suggested a potential change in the level of ASIC1a and/or ASIC2a expression. Therefore, semi-quantitative PCR was performed to provide molecular biological evidence whether such a change does take place with neuronal maturation. When PCR was conducted for each single target, fragments of the expected sizes were yielded, suggesting that individual primer pairs were adequate for the amplification of both ASIC1 and ASIC2 gene products (data not shown). To compare the relative amounts of ASIC1 and ASIC2 mRNA in cortical tissues from different ages of mice, multiplex RT-PCR was performed with the mixture of ASIC1 and ASIC2 primers. As shown in Fig. 10, the intensity of the ASIC1 fragment was stronger than that of ASIC2 at day 1. However, at day 7 or later, the ASIC2 band became clearly stronger than the ASIC1 band. Thus, the relative ratio of ASIC2/ASIC1 increased from postnatal day 1 to day 28. This observation suggests that a developmental increase in the expression of ASIC2 subunit in mouse cortical neurons might be responsible, at least partially, for the changes of the electrophysiological and pharmacological properties of ASICs.

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Figure 10. Relative change of ASIC1 and ASIC2 mRNA with neuronal development A, multiplex RT-PCR assay for simultaneous comparison of ASIC1 and ASIC2 gene expression during development. Total RNAs were isolated from mouse cortical brains at indicated ages. Equal amounts of total RNA were reverse-transcribed and PCR-amplified using specific primers for ASIC1 and ASIC2 as described in Methods. The RT-PCR products were electrophoresed on 1.5% gel and visualized. B, summary data showing age-dependent increase of relative ratio of ASIC2/ASIC1 mRNA. Bar chart indicates relative ratio between ASIC1 and ASIC2 at different ages. The ratio of ASIC2/ASIC1 mRNA (Mean ± SD) was 0.83 ± 0.20, 1.22 ± 0.12, 1.47 ± 0.18, and 1.42 ± 0.13 at day 1, day 7, day 14 and day 28, respectively (n= 5–6, **P < 0.01 for day 1 vs. day 7, day 14 and day 28, P < 0.05 for day 7 vs. day 21 and day 28).

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Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Methods
  5. Results
  6. Discussion
  7. References
  8. Appendix
  9. Supporting Information

ASICs are H+-gated cation channels abundant in neurons of the peripheral sensory and the central nervous system (CNS). Recent studies have demonstrated an important role for ASICs in a variety of physiological and pathological processes including nociception, mechanosensation, synaptic plasticity, learning/memory and acidosis-mediated neuronal injury (Wemmie et al. 2002, 2006; Krishtal, 2003; Voilley, 2004; Gao et al. 2005; Xiong et al. 2006; Friese et al. 2007; Mazzuca et al. 2007). Since ASICs in native neurons are formed by a combination of homomeric and heteromeric subunits with distinct H+ sensitivity, ion permeability and kinetics (Baron et al. 2002, 2008; Benson et al. 2002; Krishtal, 2003; Askwith et al. 2004; Wemmie et al. 2006), the overall physiological/pathological roles they can play are, therefore, largely influenced by the property of individual ASIC subunits and their relative abundance. Although ASIC1a, 1b, 2a, 2b and 3 subunits are present in peripheral sensory neurons, homomeric ASIC1a and heteromeric ASIC1a/ASIC2a channels are the most common configurations of functional ASICs in CNS neurons (Baron et al. 2002; Askwith et al. 2004; Chu et al. 2004). Thus, any change in the property and/or level of ASIC1a or ASIC2a expression, or the relative ratio of these two subunits, probably has significant impact on acid-mediated signalling in CNS neurons.

In this study, we examined the changes of electrophysiological/pharmacological properties of ASIC currents and the expression of ASIC1a and ASIC2a subunits in mouse cortical neurons at different developmental stages. We demonstrated dramatic differences in the properties of ASIC currents in mature as compared to immature neurons: (1) An increase in the amplitude and the density of the ASIC currents; (2) A decrease in the sensitivity of ASICs to H+; (3) A reduced desensitization of ASICs, and an increased rate of recovery from desensitization; (4) A decrease in the pH sensitivity of steady-state inactivation of ASICs; (5) A decrease in Ca2+ permeability; (6) An increased potentiation of the current amplitude by high concentration of extracellular Zn2+; (7) A reduced sensitivity of the currents to PcTX1. The detailed mechanism underlying these changes is not clear. However, some of the changes may be explained by an increase in the relative ratio of ASIC2a/ASIC1a expression. Although both ASIC1a and ASIC2a subunits are abundant in CNS neurons, the influence of each subunit to the properties of acid-activated currents in neurons differs dramatically. Using ASIC1 and ASIC2 knockout mice, for example, Askwith and colleagues showed a differential contribution of ASIC1a and ASIC2a subunits to the electrophysiological properties of ASICs in mouse hippocampal neurons (Askwith et al. 2004). They demonstrated that the ASIC1a subunit is critical for the amplitude of the ASIC currents whereas the ASIC2a subunit is important in determining the channel kinetics. Based on their studies, a possible interpretation to our findings is that there is a developmental increase of both ASIC1a and ASIC2a subunits in mouse cortical neurons, but the increase of ASIC2a subunit predominates. The gradual increase in the amplitude and the density of ASIC currents with neuronal maturation might suggest an increase in the expression of the ASIC1a subunit. On the other hand, an increased rate of recovery from desensitization, a decreased PcTX1 sensitivity and an increased Zn2+ potentiation are consistent with an increase in the expression of the ASIC2a subunit (Bassilana et al. 1997; Baron et al. 2001, 2008; Askwith et al. 2004). Consistent with the electrophysiological and pharmacological findings, our RT-PCR data support a relative increase in ASIC2a/ASIC1a expression. Previous studies by Baron and colleagues have shown that the expression of ASIC1a and ASIC2a subunits in mouse spinal cord neurons also increases with neuronal development (Baron et al. 2008). On the other hand, studies by de la Rosa et al. suggested that the ASIC1a expression remained at approximately the same level throughout development (Alvarez et al. 2003). In their study, whole brain tissue from rat was used, while in our current studies, we focused on the cortical tissue/neurons from mouse. It is therefore possible that the developmental changes in ASIC1a and ASIC2a might be regional and species specific.

In addition to a change of overall ASIC expression, alterations in the relative distribution of the ASIC subunit, e.g. membrane surface vs. intracellular sites, could also contribute to the changes in electrophysiological/pharmacological properties. For example, our recent studies have shown that, in the normal condition, ASIC1a is predominantly localized to the endoplasmic reticulum. It, however, can be rapidly translocated to the cell surface membrane upon certain stimulations mimicking ischemia, e.g. serum deprivation or insulin depletion, resulting in increased surface expression and the potentiation of ASIC current (Chai et al. 2010). Thus, further experiments would be needed to determine whether the expression of ASICs in cell surface membrane changes with neuronal development.

In addition to the change in the level of ASIC expression, one cannot rule out the possibility that any change in the level and properties of endogenous signalling molecules which are known to modulate the behaviour of ASICs (Askwith et al. 2000; Gao et al. 2005; Xu & Xiong, 2007; Sherwood & Askwith, 2009), could also contribute to the changes in the properties of ASICs.

The influence of the ASIC2a subunit on the rate of desensitization of the heteromeric ASIC1a/ASIC2a channels has been controversial. While some studies show reduced desensitization of the heteromeric ASIC1a/ASIC2a currents compared with the homomeric ASIC1a channels (Bassilana et al. 1997), other studies have suggested the opposite (Askwith et al. 2004). The reason for this controversy was not clear; however, a recent study by Baron et al. (2008) has provided some tentative explanation. In this study, Baron and colleagues demonstrated that the influence of ASIC2a on the desensitization kinetics of the heteromeric ASIC1a/ASIC2a channels depends on the relative abundance of ASIC2a. If the ratio of ASIC2a/ASIC1a is 1:2, the rate of desensitization is increased. However, if the ratio of ASIC2a/ASIC1a is 2:1, a decrease, rather than an increase, in the rate of desensitization of the heteromeric currents is detected (Baron et al. 2008). Thus, our data showing a decrease in the rate of desensitization with neuronal maturation are consistent with a relative increase of ASIC2a subunit or the ratio of ASIC2a/ASIC1a.

The pharmacological properties of ASICs also depend largely on the subunit composition (Baron et al. 2002; Askwith et al. 2004; Chu et al. 2004). For example, potentiation of the ASIC currents by high concentrations of zinc (100–300 mm) depends on the presence of ASIC2a-containing channels, e.g. homomeric ASIC2a and heteromeric ASIC1a/ASIC2a channels (Baron et al. 2001; Chu et al. 2004). A relative increase in the expression of ASIC2a should favour the formation of heteromeric ASIC1a/ASIC2a channels thus increasing the sensitivity of the ASIC currents to zinc potentiation. On the other hand, PcTX1 only inhibits the current mediated by homomeric ASIC1a channels (Escoubas et al. 2000; Xiong et al. 2004; Chen et al. 2005). A reduction in the sensitivity of overall ASIC currents to PcTX1 inhibition suggests a decrease in the relative contribution by homomeric ASIC1a channels to total acid-activated currents.

In addition to Na+ permeability, homomeric ASIC channels composed of ASIC1a subunits exhibit Ca2+ permeability (Waldmann et al. 1997; Yermolaieva et al. 2004; Xiong et al. 2004; Wu et al. 2004). A relative increase of ASIC2a expression in mature neurons and the formation of heteromeric ASIC1a/ASIC2a channels are expected to reduce the permeability of ASICs to Ca2+. Consistent with this notion, we show a shift in the reversal potential of acid-activated Ca2+ currents towards more negative values and a reduced acid-induced [Ca2+]i accumulation in mature neurons than younger neurons. Since activation of homomeric ASIC1a channels and the entry of Ca2+ play important roles in acidosis-mediated signalling, e.g. synaptic plasticity (Wemmie et al. 2002) and acidosis-induced neuronal injury (Yermolaieva et al. 2004; Xiong et al. 2004; Gao et al. 2005), our findings may suggest a reduced contribution of ASIC1a to the learning/memory process and ischaemic injury in mature or aged neurons than the younger neurons. Preliminary data from our on-going studies suggest that this might be the case. The changes of Ca2+ permeability of ASICs during development may suggest some roles of ASIC in the neural development.

Our studies showed some difference in the properties of ASICs between cultured neurons and acutely dissociated neurons. One noticeable difference is the reversal potential of Ca2+-mediated currents. For example, in the 7-day-cultured neurons, the reversal potential is −12.31 mV. However, it is −5.64 mV in acutely dissociated neurons from day 7 mice. One likely possibility for this is the difference in the maturation status of the cultured neurons and the acutely dissociated neurons. For example, the neurons after 7 days in culture are more mature than the neurons isolated from postnatal day 7. Thus, the Ca2+ permeability of ASICs in 7-day-cultured neurons is lower than that of acutely dissociated neurons from day 7 mice, as reflected by more negative reversal potential in cultured neurons.

In this study, we only focused on the changes of electrophysiological and pharmacological properties of ASICs in excitatory neurons based on their pyramidal shape. It is not clear whether ASICs in inhibitory interneurons undergo similar developmental changes. Since recent studies by Ziemann and colleagues have suggested that increased activation of ASIC1a channels in interneurons is involved in acidosis-mediated seizure termination (Ziemann et al. 2008), it will be interesting to explore the potential developmental changes of ASICs in these neurons in the future.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Methods
  5. Results
  6. Discussion
  7. References
  8. Appendix
  9. Supporting Information

Appendix

  1. Top of page
  2. Abstract
  3. Introduction
  4. Methods
  5. Results
  6. Discussion
  7. References
  8. Appendix
  9. Supporting Information

Author contributions

M.L. conducted experiments, analysed data and wrote the manuscript. E.K. conducted cell culture. K.I. conducted the multiple RT-PCR. R.P.S. provided partial funding and revised the paper. Z.-G.X. designed the project, interpreted the data, provided the funding, and revised the paper. All authors approved the final version of the manuscript.

Acknowledgements

These studies were supported by the grants from NIH (R01NS047506 to ZGX, R01NS050610 to R.P.S.), American Heart Association (0840132N to Z.-G.X.), and Legacy Research Advisory Committee.

Supporting Information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Methods
  5. Results
  6. Discussion
  7. References
  8. Appendix
  9. Supporting Information

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