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. 2016 Jul 1;594(13):3561-74.
doi: 10.1113/JP271925. Epub 2016 May 7.

Genetic alteration of the metal/redox modulation of Cav3.2 T-type calcium channel reveals its role in neuronal excitability

Tiphaine Voisin  1   2   3 , Emmanuel Bourinet  1   2   3 , Philippe Lory  1   2   3
Affiliations

Genetic alteration of the metal/redox modulation of Cav3.2 T-type calcium channel reveals its role in neuronal excitability

Tiphaine Voisin et al. J Physiol. .

Abstract

Key points: In this study, we describe a new knock-in (KI) mouse model that allows the study of the H191-dependent regulation of T-type Cav3.2 channels. Sensitivity to zinc, nickel and ascorbate of native Cav3.2 channels is significantly impeded in the dorsal root ganglion (DRG) neurons of this KI mouse. Importantly, we describe that this H191-dependent regulation has discrete but significant effects on the excitability properties of D-hair (down-hair) cells, a sub-population of DRG neurons in which Cav3.2 currents prominently regulate excitability. Overall, this study reveals that the native H191-dependent regulation of Cav3.2 channels plays a role in the excitability of Cav3.2-expressing neurons. This animal model will be valuable in addressing the potential in vivo roles of the trace metal and redox modulation of Cav3.2 T-type channels in a wide range of physiological and pathological conditions.

Abstract: Cav3.2 channels are T-type voltage-gated calcium channels that play important roles in controlling neuronal excitability, particularly in dorsal root ganglion (DRG) neurons where they are involved in touch and pain signalling. Cav3.2 channels are modulated by low concentrations of metal ions (nickel, zinc) and redox agents, which involves the histidine 191 (H191) in the channel's extracellular IS3-IS4 loop. It is hypothesized that this metal/redox modulation would contribute to the tuning of the excitability properties of DRG neurons. However, the precise role of this H191-dependent modulation of Cav3.2 channel remains unresolved. Towards this goal, we have generated a knock-in (KI) mouse carrying the mutation H191Q in the Cav3.2 protein. Electrophysiological studies were performed on a subpopulation of DRG neurons, the D-hair cells, which express large Cav3.2 currents. We describe an impaired sensitivity to zinc, nickel and ascorbate of the T-type current in D-hair neurons from KI mice. Analysis of the action potential and low-threshold calcium spike (LTCS) properties revealed that, contrary to that observed in WT D-hair neurons, a low concentration of zinc and nickel is unable to modulate (1) the rheobase threshold current, (2) the afterdepolarization amplitude, (3) the threshold potential necessary to trigger an LTCS or (4) the LTCS amplitude in D-hair neurons from KI mice. Together, our data demonstrate that this H191-dependent metal/redox regulation of Cav3.2 channels can tune neuronal excitability. This study validates the use of this Cav3.2-H191Q mouse model for further investigations of the physiological roles thought to rely on this Cav3.2 modulation.

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Figures

Figure 1
Figure 1. Generation of a knock‐in (KI) mouse line carrying the mutation H191Q on the T‐type Cav3.2 channel
A, the Cav3.2 allele was created by homologous recombination (see Materials and Methods). The CAC codon encoding histidine 191 (H191) in exon 4 of the WT allele (upper panel) was replaced by the codon CAG encoding glutamine (Q), and a loxP‐flanked neo cassette was introduced 3′ from exon 4 to select for embryonic stem cells harbouring the knock‐in (KI) allele (middle panel). The final mutant allele was obtained after excision of the neo cassette by a Cre recombinase treatment of embryonic stem cells (lower panel). B, henotyping of the Cav3.2‐H191Q KI mice. PCR analysis using specific primers positioned in A revealed the WT allele (488 bp) in both WT and heterozygous animals as well as the mutant allele (533 bp) in both KI and heterozygous animals. The 45 bp difference between mutant and WT alleles results from the remaining loxP site in the mutant allele. C, representative Western‐blot analysis of the expression level of the Cav3.2 protein in the whole brain of WT and KI mice; a whole brain extract from knock‐out mice for Cav3.2 (KO) was used as a negative control in these experiments. The expression level of α‐GAPDH was used as loading control in these experiments (lower panel).
Figure 2
Figure 2. Electrophysiological properties of T‐type currents in DRG neurons from WT and KI mice
A, T‐type calcium current density in D‐hair DRG neurons from wild‐type (WT) and knock‐in (KI) mice, in control conditions (left) and in the presence of 10 μm of the T‐type channel inhibitor TTA‐A2 (right). For each cell, the current amplitude (in pA) was measured for a depolarization step from −80 to −30 mV and normalized for the cell capacitance (pF) in current density (pA pF–1). Values are expressed as mean ± SEM. Inset: bright field image of a typical D‐hair neuron with its ‘rosette’ morphology. Scale bar = 30 μm. B, normalized I–V relationship for the WT (grey square) and the KI (black circle) current evoked by 90 ms depolarizing step pulses from a holding potential at −80 mV. C, steady‐state inactivation curves obtained by stepping the membrane potential at −30 mV from conditioning depolarizing pulses ranging from −100 to −30 mV. D, deactivation kinetics as a function of the voltage.
Figure 3
Figure 3. Metal/redox sensitivity of T‐type currents in DRG neurons from WT and KI mice
A, representative T‐type current traces recorded from D‐hair cells isolated from a WT mouse (upper trace) and a KI mouse (lower trace) evoked by 90 ms depolarization steps from −80 to −30 mV, before and after exposure to 1 and 10 μm zinc. B, dose–response curves obtained for zinc inhibition of T‐type currents in neurons from WT mice and KI mice. Current amplitude was normalized to the peak current in the absence of zinc, and the remaining current was plotted against zinc concentration. The IC50 values were obtained from fitted data using a sigmoidal dose–response with variable Hill slope equation. Values are mean ± SEM (***P < 0.001; two‐way ANOVA). C, the ability of ascorbate (300 μm) and nickel ions (1 and 30 μm) to inhibit T‐type currents in D‐hair neurons from WT (black bars) and KI (grey bars) mice is represented as the percentage of remaining current following drug application. D, effect of zinc chelator TPEN (10 μm) and l‐cysteine (100 μm) on T‐type current. Note that TPEN and l‐cysteine in control conditions fail to modulate either the WT or the KI current. WT TPEN vs. KI TPEN: P = 0.52; WT l‐cysteine vs. KI l‐cysteine: P = 0.0734; Mann–Whitney). E, effect of TPEN (10 μm) and l‐cysteine (100 μm) on the T‐type current from WT and KI mice in the presence of 1 μm zinc. Values are expressed as mean ± SEM (**P < 0.01; ***P < 0.001; Mann–Whitney).
Figure 4
Figure 4. Action potential (AP) properties in DRG neurons from WT and KI mice
A, typical traces of evoked APs. Depolarization currents of increasing amplitude were injected to trigger an AP in the absence (left) or presence of 1 μm zinc (right) in D‐hair neurons from WT (upper panels) and KI (lower panels) mice. The resting potential of these neurons was maintained at −70 mV. B, threshold current for an AP generation (Rheobase) in control condition and with 30 μm nickel for neurons from WT mice (left) and KI mice (right). Statistical analysis was performed with repeated‐measures two‐way ANOVA (WT Ctrl vs. WT Ni 30 μm, *P < 0.05; WT Ctrl vs. KI Ctrl, P = 0.52; WT Ni 30 μm vs. KI Ni 30 μm, P = 0.08; KI Ctrl vs. KI Ni 30 μm, P = 0.95).
Figure 5
Figure 5. Afterdepolarization (ADP) properties in DRG neurons from WT and KI mice
A, representative APs with ADP before and after application of zinc (1 and 30 μm) in D‐hair neurons of WT (left) and KI mice (right). The resting membrane potential of the neurons was maintained at −70 mV. B, ADP amplitude (in mV) in control condition is determined as the difference in voltage from the resting potential value (*P < 0.05; Mann‐–Whitney). C and D, effect of the application of zinc at 1 and 30 μm (C) and nickel at 30 μm (D) on ADP amplitude. Statistical analysis was performed with two‐way ANOVA followed by Sidak's test (*P < 0.05; ***P < 0.001; KI Ctrl vs. KI Zn 1 μm, P = 0.22; WT Ctrl vs. WT Ni 1 μm, P = 0.11; KI Ctrl vs. KI Ni 1 μm, P = 0.61; KI Ctrl vs. KI Ni 30 μm, P = 0.43).
Figure 6
Figure 6. Rebound activity, low threshold calcium spike (LTCS) and its modulation by zinc in DRG neurons from WT and KI mice
A, typical traces of the rebound potential LTCS evoked following 2 s hyperpolarizing steps of increasing amplitude in D‐hair neurons in control (left panel) and in the presence of zinc (1 μm, right panel) from WT (upper panel) and KI (lower panel) mice. B, LTCS threshold in control condition (circle) and in the presence of zinc (1 μm, square) in D‐hair neurons from WT mice (black symbols) and KI (grey symbols) mice. Statistical analysis was performed with repeated‐measures two‐way ANOVA (**P < 0.01; KI Ctrl vs. KI Zn 1 μm, P = 0.99; WT Ctrl vs. KI Ctrl, P = 0.09; WT Zn 1 μm vs. KI Zn 1 μm, P = 0.70).
Figure 7
Figure 7. Zinc modulation of LTCS amplitude in DRG neurons from WT and KI mice
A, typical traces of LTCS evoked by a 2 s hyperpolarizing pulse in D‐hair neurons from WT (left) and KI mice (right) before and after exposure to 1 μm zinc. B, averaged amplitude of the LTCS in DRG neurons from WT and KI mice. The amplitude of the LTCS is measured as the difference from the resting membrane potential value (P = 0.27; Mann–Whitney). C, normalized amplitude of the LTCS before and after exposure to 1 and 30 μm zinc. Statistical analysis was performed with two‐way ANOVA followed by Sidak's test (***P < 0.001; KI Ctrl vs. KI Zn 1 μm, P = 0.19). D, normalized amplitude of the LTCS before and after exposure to 1 and 30 μm nickel. Statistical analysis was performed with two‐way ANOVA followed by Sidak's test (*P < 0.05; **P < 0.01; ***P < 0.001; KI Ctrl vs. KI Ni 1 μm, P = 0.96).
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