Atypical changes in DRG neuron excitability and complex pain phenotype associated with a Nav1.7 mutation that massively hyperpolarizes activation

Sodium channel Nav1.7 plays a central role in pain-signaling: gain-of-function Nav1.7 mutations usually cause severe pain and loss-of-function mutations produce insensitivity to pain. The Nav1.7 I234T gain-of-function mutation, however, is linked to a dual clinical presentation of episodic pain, together with absence of pain following fractures, and corneal anesthesia. How a Nav1.7 mutation that produces gain-of-function at the channel level causes clinical loss-of-function has remained enigmatic. We show by current-clamp that expression of I234T in dorsal root ganglion (DRG) neurons produces a range of membrane depolarizations including a massive shift to >−40 mV that reduces excitability in a small number of neurons. Dynamic-clamp permitted us to mimic the heterozygous condition via replacement of 50% endogenous wild-type Nav1.7 channels by I234T, and confirmed that the I234T conductance could drastically depolarize DRG neurons, resulting in loss of excitability. We conclude that attenuation of pain sensation by I234T is caused by massively depolarized membrane potential of some DRG neurons which is partly due to enhanced overlap between activation and fast-inactivation, impairing their ability to fire. Our results demonstrate how a Nav1.7 mutation that produces channel gain-of-function can contribute to a dual clinical presentation that includes loss of pain sensation at the clinical level.

37˚C. Transfected HEK293 cells were treated with G418 for several weeks to establish the stable cell lines expressing sodium currents.

Voltage-clamp recordings
Voltage-clamp recordings at room temperature (22 ± 1°C) were performed using an EPC-10 amplifier and the Patchmaster program (v 53; HEKA Elektronik). Recordings at 33˚C were obtained using an Axon MultiClamp 700B amplifier (Molecular Devices, US). Data were digitized via an analogue to digital converter Digidata 1440a (Molecular Devices, US). Acquired data were analyzed using pClamp v10.6, Origin v9.1, Excel and SPSS 24. Temperature was controlled using an adapted Warner Instruments CL-100 temperature controller and SC-20 Dual Inline Heater/Cooler system (Warner Instruments).
Fire-polished electrodes were fabricated from 1.6 mm outer diameter borosilicate glass micropipettes (World Precision Instruments) using a Sutter Instruments P-97 puller and had a resistance of 0.7-1.5 MΩ. Pipette potential was adjusted to zero before seal formation. Liquid junction potential was not corrected. To reduce voltage errors, 80%-90% series resistance compensation was applied. Cells were excluded from analysis if the predicted voltage error exceeded 3 mV. Linear leak currents were subtracted out using the P/N method. Sodium current recordings were initiated after a 5 min equilibration period once whole-cell configuration was achieved. Current traces were sampled at 50 kHz and filtered with a low-pass Bessel setting of 10 kHz. The pipette solution contained the following (in mM): 140 CsF, 10 NaCl, 1 EGTA, and 10 HEPES, 10 dextrose, pH 7.30 with CsOH (adjusted to 310 mOsmol/L with sucrose). The extracellular bath solution contained the following (in mM): 140 NaCl, 3 KCl, 1 MgCl2, 1 CaCl2, 10 HEPES, 10 dextrose, pH 7.30 with NaOH (adjusted to 320 mOsmol/L with sucrose). TTX 3 (300 nM) was included in the bath to block the endogenous sodium currents in HEK293 cells.
Cells were held at -120 mV for all parameters examined. Recovery of hNav1.7 channels from fast inactivation at room temperature was examined using a two-pulse protocol with interpulse intervals varying from 1 to 513 ms. Recovery rates were measured by normalizing peak current elicited by the test pulse (10 ms depolarization to -10 mV) to that of the prepulse (20 ms at -10 mV) at voltages ranging from -120 mV to -70 mV in 10 mV increments. Normalized peak current at each individual voltage was plotted against the interpulse interval and fit with a singleexponential equation of the form I = A × exp (−t/τ) + Ic, where A is the amplitude of the fit, t is time, τ is the time constant of decay, and Ic is the asymptotic minimum to which the tail currents decay. To assess the recovery rate from slow-inactivation at room temperature, cells were prepulsed with a 30 s stimulus at -10 mV to allow the channel to enter slow-inactivation, followed by a range of conditioning voltages from -130 mV to -90 mV in 10 mV increments varying from 4-16384 ms. A 100 ms step to -120 mV was applied immediately afterwards to remove fast inactivation before a 20 ms depolarizing step to -10 mV as a final step to elicit a test response, which reflects the remaining currents that have recovered from slow-inactivation.
Sweep interval was increased to 60 s to allow the channel to recover from slow-inactivation.
To assess the biophysical properties of I234T mutant channels at skin/more physiological temperature, we performed voltage-clamp experiments at 33˚C in HEK293 cells stably expressing WT or I234T Nav1.7 channels. Current-voltage relationships were obtained by applying a series of 100 ms depolarizing steps from -80 to +50 mV in 5 mV increments from a holding potential of -120 mV. Conductance was calculated as G = I / (Vm -ENa), normalized to the maximum conductance and fit with Boltzmann equation. Steady-state fast inactivation curves were generated by applying 500 ms inactivating potentials from -120 to -10 mV, followed by a 4 40 ms step to -10 mV. Slow inactivation properties were investigated by stepping the membrane potential from -140 to +10 mV in 10 mV increments for 30 s from Vhold of -120 mV, followed by a 100 ms pulse to -120 mV in order to allow recovery from fast inactivation, after which a 40 ms depolarizing step to -10 mV was applied in order to recruit the remaining channels. Normalized conductance values were fit to Boltzmann equation in order to quantify the voltage dependence of steady-state fast and slow inactivation. Deactivation time constants for WT and I234T channels were obtained by fitting single exponential function to tail currents elicited by applying repolarization pulses from -120 to -50 mV after activating the channels at -10 mV for 0.5 ms.
Persistent currents were measured as mean amplitudes of currents between 93 and 98 ms after the onset of depolarization, and are presented as a percentage of the maximal peak current.
Independent t test was used in statistical analysis unless otherwise mentioned. P < 0.05 is considered as statistically significant.      9 Figure S2