Resurgent sodium current promotes action potential firing in the avian auditory brainstem.

KEY POINTS: Auditory brainstem neurons of all vertebrates fire phase-locked action potentials (APs) at high rates with remarkable fidelity, a process controlled by specialized anatomical and biophysical properties. This is especially true in the avian nucleus magnocellularis (NM) - the analogue of the mammalian anteroventral cochlear nucleus. In addition to high voltage-activated potassium (KHVA ) channels, we report, using whole cell physiology and modelling, that resurgent sodium current (INaR ) of sodium channels (NaV ) is equally important and operates synergistically with KHVA channels to enable rapid AP firing in NM. Anatomically, we detected strong NaV 1.6 expression near hearing maturation, which was less distinct during hearing development despite functional evidence of INaR , suggesting that multiple NaV channel subtypes may contribute to INaR . We conclude that INaR plays an important role in regulating rapid AP firing for NM neurons, a property that may be evolutionarily conserved for functions related to similar avian and mammalian hearing. ABSTRACT: Auditory brainstem neurons are functionally primed to fire action potentials (APs) at markedly high rates in order to rapidly encode the acoustic information of sound. This specialization is critical for survival and the comprehension of behaviourally relevant communication functions, including sound localization and distinguishing speech from noise. Here, we investigated underlying ion channel mechanisms essential for high-rate AP firing in neurons of the chicken nucleus magnocellularis (NM) - the avian analogue of bushy cells of the mammalian anteroventral cochlear nucleus. In addition to the established function of high voltage-activated potassium channels, we found that resurgent sodium current (INaR ) plays a role in regulating rapid firing activity of late-developing (embryonic (E) days 19-21) NM neurons. INaR of late-developing NM neurons showed similar properties to mammalian neurons in that its unique mechanism of an 'open channel block state' facilitated the recovery and increased the availability of sodium (NaV ) channels after depolarization. Using a computational model of NM neurons, we demonstrated that removal of INaR reduced high-rate AP firing. We found weak INaR during a prehearing period (E11-12), which transformed to resemble late-developing INaR properties around hearing onset (E14-16). Anatomically, we detected strong NaV 1.6 expression near maturation, which became increasingly less distinct at hearing onset and prehearing periods, suggesting that multiple NaV channel subtypes may contribute to INaR during development. We conclude that INaR plays an important role in regulating rapid AP firing for NM neurons, a property that may be evolutionarily conserved for functions related to similar avian and mammalian hearing.

Introduction

Voltage‐dependent sodium (NaV) channels play a critical role in generation of action potentials (APs), the firing pattern of which is fundamental for information processing in the nervous system (Eijkelkamp et al. 2012). A unique property of some NaV channels is a resurgent sodium current (I NaR) (Raman & Bean, 1997). I NaR is the result of a voltage‐dependent open channel block of the NaV α‐subunit by an intracellular particle that competes with the classic inactivation gate (i.e. the cytoplasmic linker between the III and IV domains of the α‐subunit) during depolarization. Channels that are in the ‘blocked’ state, but not the ‘classic‐inactivated’ state, generate I NaR during AP repolarization because the open channel blocker loses affinity for the α‐subunit at repolarized membrane potentials. As a result, I NaR provides a small depolarizing drive to the membrane near the AP threshold – a property that promotes repetitive neuronal firing. In addition, activation of the open channel blocker facilitates the recovery of NaV channels after depolarization and increases NaV channel availability by competing against classic inactivation (Raman & Bean, 2001). This specific blocker has been identified as the β4‐subunit in cerebellar Purkinje cells (Grieco et al. 2005; Aman et al. 2009), whereas multiple α‐subunits (e.g. NaV1.2, 1.5, 1.6 and 1.7) have been shown to be capable of carrying I NaR (Rush et al. 2005; Jarecki et al. 2010). As a result of unique features of the open channel blocker, studies show that I NaR plays an important role in promoting high rates of AP firing in numerous mammalian neurons, underlying their highly specialized information processing patterns (Lewis & Raman, 2014).

In the auditory brainstem of all vertebrates, neurons are known for their remarkable ability to fire APs at high rates (Oertel, 1997). This specialization is important for encoding sound in an ultrafast and temporally precise manner, properties essential for survival and comprehension of behaviourally relevant communication functions, including sound localization and signal extraction in complex listening environments (Shannon et al. 1995; Anderson et al. 2010; Grothe et al. 2010). An exemplar of this ability is well studied in the avian auditory brainstem. Neurons in nucleus magnocellularis (NM) – the avian analogue of bushy cells of the mammalian anteroventral cochlear nucleus – are able to phase‐lock to inputs up to 1000 Hz with high fidelity, as demonstrated by single‐unit recordings from barn owls (Koppl & Carr, 1997). Similarly, in the mammalian auditory brainstem, the calyx of Held is a pivotal synapse involved in the microsecond precision of sound localization computations. Here, fast NaV channel kinetics (Leao et al. 2005) and abundant expression of high voltage‐activated potassium channels (KHVA) (Wang et al. 1998) along with I NaR present at both pre‐ and postsynaptic sites (Leao et al. 2006; Kim et al. 2010) promote high rates of AP firing. Based on the aforementioned properties of I NaR in mammals, as well as the established and shared functional phenotypes between species (Carr & Soares, 2002), we hypothesized that this unique current also plays an important role in shaping the fast firing pattern observed for avian NM neurons.

In this study we tested whether NM neurons present with I NaR. By investigating the function of I NaR in AP firing rates of chicken NM neurons both experimentally and computationally, we found that NM neurons have robust I NaR that significantly increases NaV availability immediately after depolarization and facilitates NaV channel recovery. Removal of I NaR in a model NM neuron undermines its ability to fire at high rates. We also examined the maturation of I NaR relative to hearing onset and the potential NaV channel subtype that carries this current in developing NM using immunocytochemistry. We conclude that I NaR plays an important role in regulating rapid AP firing for NM neurons, a property that may be evolutionarily conserved for functions related to similar avian and mammalian hearing.

Methods

In vitro electrophysiology in brainstem slices

Slice preparation

All animal procedures were approved by the Northwestern University and Florida State University Institutional Animal Care and Use Committees and conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Acute brainstem slices were prepared from 96 White Leghorn chicken (Gallus gallus domesticus) embryos of either sex as previously described (Sanchez et al. 2011; Hong et al. 2016). Briefly, embryos were rapidly decapitated and the brain was dissected from the skull to isolate the brainstem region of interest. This procedure is consistent with the recommendation from the Panel on Euthanasia of the American Veterinary Medical Association and is appropriate for the species, stages of development and size of the embryos. For electrophysiological experiments, eggs were obtained from Sunnyside Farms, Inc. (Beaver Dam, WI, USA) and incubated in the central auditory physiology laboratory at Northwestern University. For immunocytochemical experiments, eggs were obtained from Charles River Laboratories (Wilmington, MA, USA) and incubated in a Florida State University vivarium. The authors understand and conform to the principles and regulations described by The Journal of Physiology (Grundy, 2015).

To study the stimulus frequency–firing pattern of NM neurons and the properties and function of I NaR, brainstem slices were taken from chickens at embryonic days (E) 19–21. Frequency‐firing pattern is defined as the calculated firing probability (see below) of APs as a function of stimulus frequency. At this age range, near‐mature hearing ability is established (Saunders et al. 1973; Rebillard & Rubel, 1981; Jones et al. 2006) and NM neurons have obtained mature‐like morphology and physiology (Jhaveri & Morest, 1982b). To study the development of I NaR, along with the development of frequency–firing pattern, chickens at the age of E11–12 and E14–16 were included in the current study, corresponding to before and during hearing onset, respectively, while chickens at E19–21 were considered as after hearing onset (Jones et al. 2006). The brainstem was dissected and isolated in ice‐cold (∼0°C) oxygenated low‐Ca2+, high‐Mg2+ modified artificial cerebral spinal fluid (ACSF) containing the following (in mm): 130 NaCl, 2.5 KCl, 1.25 NaH2PO4, 26 NaHCO3, 3 MgCl2, 1 CaCl2 and 10 glucose. ACSF was continuously bubbled with a mixture of 95% O2–5% CO2 (pH 7.4, osmolarity 295–310 mosmol l−1). The brainstem was blocked coronally, affixed to the stage of a vibrating blade microtome slicing chamber (Ted Pella, Inc., Redding, CA) and submerged in ice‐cold ACSF. Bilaterally symmetrical coronal slices were made (200–300 μm thick) and approximately three to seven slices (depending on age) containing NM were taken from caudal to rostral, roughly representing the low‐to‐high frequency regions, respectively. All neurons reported here were obtained from the rostral one‐half of the entire nucleus, roughly representing the mid‐ to high‐frequency regions of NM.

Slices were collected in a custom holding chamber and allowed to equilibrate for 1 h at room temperature in normal ACSF containing the following (in mm): 130 NaCl, 2.5 KCl, 1.25 NaH2PO4, 26 NaHCO3, 1 MgCl2, 3 CaCl2 and 10 glucose. Normal ACSF was continuously bubbled with a mixture of 95% O2–5% CO2 (pH 7.4, osmolarity 295–310 mosmol l−1). Slices were transferred to a recording chamber mounted on an Olympus BX51W1 (Center Valley, PA, USA) microscope for electrophysiological experiments. The microscope was equipped with a CCD camera, ×60 water‐immersion objective and infrared differential interference contrast optics. The recording chamber was superfused continuously with a motorized pump (Welco, Tokyo, Japan) at room temperature (monitored continuously with a bath emerged thermometer at ∼24°C, Warner Instruments, Hamden, CT, USA) in oxygenated normal ACSF at a rate of 1.5–2 ml min−1.

Whole cell electrophysiology

Voltage‐clamp and current‐clamp experiments were performed using an Axon Multiclamp 700B amplifier (Molecular Devices, Sunnyvale, CA, USA). Patch pipettes were pulled to a tip diameter of 1–2 μm using a P‐97 Flaming–Brown micropipette puller (Sutter Instrument, Novato, CA, USA) and had resistances ranging from 3 to 6 MΩ. For voltage‐clamp experiments of isolated NaV currents, the internal solution was caesium‐based and contained the following (in mm): 150 CsCl, 10 NaCl, 0.2 EGTA and 10 HEPES, pH adjusted to 7.3–7.4 with CsOH. The Cs+‐based internal solution was used to block KV currents and reduce space‐clamp issues. The junction potential was ∼−3 mV. Series resistance was compensated for by ∼80% in all voltage‐clamp recordings. For current‐clamp experiments, the internal solution was potassium‐based and contained the following (in mm): 105 potassium gluconate, 35 KCl, 1 MgCl2, 10 HEPES‐K, 5 EGTA, 4 4‐Mg2ATP, and 0.3 4‐Tris2GTP, pH adjusted to 7.3–7.4 with KOH. The junction potential was ∼−10 mV. Data in both voltage clamp and current clamp experiments were not corrected for junction potentials.

Pipettes were visually guided to NM and neurons were identified and distinguished from surrounding tissue based on cell morphology, known structure and location of the nucleus within the slice. After a gigaohm seal was attained, membrane patches were ruptured and neurons were first held in the voltage clamp mode of the whole‐cell configuration. A small hyperpolarizing (−1 mV, 30 ms) voltage command was presented to monitor whole‐cell parameters (i.e. cell membrane capacitance, series resistance and input resistance). NM neurons were included in the data analysis only if they had series resistances <15 MΩ. Raw data were low‐pass filtered at 5 kHz and digitized at 50 kHz using a Digidata 1440A (Molecular Devices).

All experiments were conducted in the presence of the GABAA receptor antagonist picrotoxin (100 μm). Synaptic glutamate transmission was continuously blocked using dl‐2‐amino‐5‐phosphonopentanoic acid (dl‐APV, 100 μm, an NMDA receptor antagonist) and 6‐cyano‐7‐nitroquinoxaline‐2,3‐dione (CNQX, 20 μm, an AMPA receptor antagonist). In current‐clamp experiments, tetraethylammonium (TEA, 1 mm) was used to block KV3‐containing channels. Current commands of square pulse trains (duration 1 s) were injected into the soma of E19–21 NM neurons before and during TEA application. The current strength was 1 nA and individual square pulse width was 2 ms. Square pulse trains were applied at varying frequencies: 50, 100, 150, 200, 250 and 300 Hz. In order to profile the frequency–firing pattern for NM neurons, firing probability per square pulse (for simplicity, ‘firing probability’) was calculated as the number of APs divided by the total number of square pulses and plotted as a function of stimulus frequency. To study the development of frequency–firing pattern, the same current commands of square pulse trains were used for NM neurons at E14–16. For NM neurons at E11–12, however, individual square pulse width was extended to 5 ms, because neurons at this age are presented with significantly wider AP half‐width (average = 4.6 ms) than the other two age groups (Hong et al. 2016). In addition, square pulse trains were applied at different frequencies: 10, 30, 50, 70, 100, 120 and 150 Hz.

In voltage‐clamp experiments, isolated NaV currents were recorded with bath application of TEA (3 mm), 4‐AP (30 μm) and CdCl2 (0.2 mm) to block potassium and calcium channels. Classic voltage‐clamp protocols to elicit I NaR were used for NM neurons (Raman & Bean, 1997; see Results). Briefly, repolarizations at membrane voltages from −70 to 0 mV (in a step of 5 mV) were applied after a depolarizing conditioning step. Three levels of conditioning step were used, +30, 0 and −30 mV, along with two durations, 10 and 100 ms. I NaR properties were characterized separately under these six experimental conditions. Afterwards, tetrodotoxin (TTX, 1 μm) was bath applied and the same voltage‐clamp protocols were repeated during TTX application. Data reported in this study were obtained by subtracting the TTX‐insensitive current traces from the control traces. Capacitive currents generated during voltage‐clamp recordings were blanked or reduced offline.

Data analysis

Recording protocols were written and run using Clampex acquisition and Clampfit analysis software (v. 10.3; Molecular Devices). Statistical analyses and graphing protocols were performed using Prism (v. 7.0b; GraphPad Software, La Jolla, CA, USA) and MATLAB (v. R2014b; The MathWorks, Natick, MA, USA) software. Student's t test or analysis of variance (ANOVA) with post hoc Bonferroni adjusted t test was used to determine significance. The standard for significant differences was defined as P < 0.05. Numerical values in the text are reported as means ± standard deviation (SD). Numerical values in Table 3 are reported as means ± standard error of the mean (SEM). Error bars in all figures represent SEM.

Table 3

Sodium channel current properties

Amplitude	E11–12 (n)	E14–16 (n)	E19–21 (n)	ANOVA
Transient I NaV a (nA)	−1.57 ± 0.13 (7)	−2.65 ± 0.30 (12)	−3.85 ± 0.40 (12)	P = 0.0005
Resurgent I NaV b (nA)	−0.10 ± 0.01 (10)	−0.44 ± 0.05 (12)	−0.60 ± 0.04 (17)	P < 0.0001
Persistent I NaV b , c (nA)	−0.05 ± 0.01 (10)	−0.12 ± 0.02 (12)	−0.14 ± 0.01 (16)	P = 0.001

Transient NaV current = amplitude calculated at step depolarization to −30 mV (holding voltage = −90 mV). Resurgent and persistent NaV current = amplitude calculated at the repolarizing membrane voltage of −30 mV. The conditioning step = +30 mV for 10 ms. Persistent NaV current = measured at the end of the 100 ms repolarizing membrane voltage.

Reagents

All bath‐applied drugs except for TTX were allowed to perfuse through the recording chamber for ∼10 min before subsequent recordings. TTX application was allowed for 3–5 min before subsequent recordings. dl‐APV, CNQX and all other salts and chemicals were obtained from Sigma‐Aldrich (St Louis, MO, USA). Picrotoxin was obtained from Tocris Bioscience (Ellisville, MO, USA). TTX was obtained from Alomone Labs (Jerusalem, Israel). TEA was obtained from VWR (Radnor, PA, USA).

Computational modelling

Based on NM models previously described in detail elsewhere (Howard & Rubel, 2010; Lu et al. 2017), a single‐compartment computational model was constructed using NEURON 7.1 (Table 1) (Hines & Carnevale, 1997). This model contains currents mediated by low and high voltage‐activated potassium channels (KLVA and KHVA, respectively), NaV and passive leak channels. Modelling schemes were identical to those described in Lu et al. (2017) for all membrane currents except for NaV currents (Table 2). The Hodgkin–Huxley style formalism, previously employed to model NaV current, was replaced with a Markovian 13‐state NaV‐channel model, which generates the transient (I NaT), persistent (I NaP) and resurgent (I NaR) current components simultaneously (Fig. 1) (Khaliq et al. 2003). We adopted a modified version described previously (Akemann & Knopfel, 2006), which includes the Q 10 parameter to adjust the NaV current to match the experimental recording temperature of 24°C. Q 10 is a measure of the degree to which a biological process depends on temperature. It is defined as the ratio between the rate of a biological process at two temperatures separated by 10°C. In the context of ion channels, it can be applied to the temperature dependence of the rate of channel opening and closing and to the dependence of maximum channel conductance on temperature.

Table 1

Single compartment model

Parameter	Value
Axial resistance	50 Ω cm
Temperature	24°C
E Na	44 mV
E K	−80 mV
Length	20 μm
Diameter	20 μm
g Leak	0.002 S cm−2
g NaV	0.025 S cm−2
g KLVA	0.0069 S cm−2
g KHVA	0.004 S cm−2

Table 2

Model parameters

Parameter	Value
I NaV, Markovian model
α	150exp(V/20) ms−1
β	3exp(−V/20) ms−1
γ	150 ms−1
δ	40 ms−1
ε	1.75 ms−1
ζ	0.03exp(−V/25) ms−1
C on	0.005 ms−1
C off	0.5 ms−1
O on	0.75 ms−1
O off	0.005 ms−1
a	(O on/C on)1/4
b	(O off/C off) 1/4
Q 10	3
T 0	22°C
I KLVA, Hodgkin–Huxley model
w∞	1/(1 + exp(−(V + 67)/8)
z∞	1/(1 + exp(−(V + 71)/10)
τw	(100/(6 × exp((V + 60)/6) + 16 × exp(−(V + 60)/45)) + 1.5
τz	(100/(exp((V + 60)/20) + exp(−(V + 60)/8)) + 50
Q 10	3
T 0	22°C
I KHVA, Hodgkin–Huxley model
n∞	1/(1 + exp(−(V + 42.5)/4.5)
p∞	1/(1 + exp(−(V + 11.2)/6)
τn	(100/(11 × exp((V + 60)/24) + 21 × exp(−(V + 60)/23)) + 0.7
τp	(100/(4 × exp((V + 60)/32) + 5 × exp(−(V + 60)/22)) + 5
Q 10	3
T 0	22°C

V, C and O denote voltage, closed and opened states, respectively.

Figure 1

Markovian state model of the sodium (NaV) channel

Reproduced from the model described by Khaliq et al. (2003). C, I, O, and OB denote closed, inactivated, open and open‐blocked states, respectively. The values of the kinetic parameters are shown in Table 2.

To remove the I NaR component, the rate constant for the O→OB transition, ε, was set to zero. This resulted in considerable slowing of I NaT decay (see Fig. 6 B, blue trace). To modify I NaR without affecting I NaT and I NaP, we applied the method described previously (Magistretti et al. 2006). Kinetic constants were modified as follows: ε was set to 0; O on and O off were increased to 2.15 ms−1 and 0.01433 ms−1 to restore I NaT and I NaP, respectively (see Fig. 6 B and C).

Figure 6

Simulations of NaV currents from the model NM neuron

Aa, simulated current traces (upper panel) in response to the voltage‐clamp protocol (lower panel) consisting of a 10 ms conditioning step to +30 mV, followed by step repolarizations to −10, −30, −40, −50 and −70 mV. Arrowhead and small arrow indicate I NaT and I NaR, respectively. Ab, current–voltage relationship of simulated I NaR. B, simulated I NaT (upper panel) was evoked by depolarizing steps to 0 mV (lower panel). The current obtained under control condition is shown in black. Switching off I NaR by setting the rate constant for the O→OB transition, ε, to 0 resulted in considerable slowing of I NaT decay (0‐I NaR− condition, blue trace). I NaT was restored under 0‐I NaR+ condition (ε = 0, O on = 2.15 ms−1, and O off = 0.01433 ms−1; red trace). C, simulated I NaR under control (black trace) and 0‐I NaR+ condition (red trace). Removal of I NaR under the 0‐I NaR+ condition has no effect on persistent NaV current (I NaP).

Immunocytochemistry

Chicken embryo brains (E11, E15 and E21; n = 3 for each age) were dissected out of the skull and immediately immersed in modified periodate–lysin–paraformaldehyde (PLP) fixative: 0.2% (w/v) paraformaldehyde, 2.7% (w/v) lysin HCl, 0.21% (w/v) NaIO4, and 0.1% (w/v) Na2HPO4 (Kuba et al. 2005). Brains of all ages were then transferred to 30% sucrose in 0.1 m phosphate buffer for 3 days and sectioned in the coronal plane at 30 μm on a freezing sliding microtome. Each section was collected in 0.01 m phosphate‐buffered saline (PBS) with 0.02% sodium azide. Alternate serial sections were immunocytochemically stained for anti‐NaV1.6, generously provided by Dr Hiroshi Kuba at Kyoto University (Kuba et al. 2006). Briefly, free‐floating sections were incubated with primary antibody solutions (1:1000) diluted in PBS with 0.3% Triton X‐100 overnight at 4°C, followed by Alexa‐Fluor secondary antibodies (Thermo Fisher Scientific, Waltham, MA, USA) at 1:400 overnight at 4°C. Sections from E21 chickens were double stained with Neurofilament 200 (Sigma‐Aldrich, N0142). Sections of all ages were then mounted on gelatin‐coated slides and coverslipped with Fluoromount‐G mounting medium® (Southern Biotech, Birmingham, AL, USA). Images were captured with an Olympus FV1200 confocal microscope. Image brightness, gamma and contrast adjustments were performed in Adobe Photoshop (Adobe Systems, San Jose, CA, USA).

Results

Frequency–firing pattern of NM neurons

Auditory brainstem neurons are able to fire APs at high rates (Oertel, 1997; Trussell, 1997, 1999). Therefore, we first tested whether late‐developing NM neurons (E19–21) follow varying frequency rates with high fidelity. Square pulse current trains at frequencies from 50 to 300 Hz were injected into the soma of NM neurons. Firing probability at each frequency was defined as the average number of APs per square pulse (see Methods). The majority of NM neurons (11 out of 12 neurons) were able to follow square pulse trains of 150 Hz in a one‐to‐one manner (i.e. high‐fidelity, Fig. 2 Aa and D). Firing probability dropped to ∼0.6 at 200 Hz when obvious failures of spike generation are observed during the stimulus (Fig. 2 Ca and D, asterisks). When increasing the stimulus frequency beyond 200 Hz, firing probability was further reduced below 0.5 and in response to 300 Hz stimulation, NM neurons only fired a single onset AP (data not shown). Therefore, NM neurons are capable of firing APs at rates up to 200 Hz with good fidelity, and this ability of frequency firing is similar to other auditory brainstem neurons (Gao & Lu, 2008).

Figure 2

Frequency–firing pattern of NM neurons at E19–21

Aa and b, representative voltage responses recorded from an E20 NM neuron to current injections of square pulse trains at 150 Hz, before and during TEA (1 mm) application, respectively. B, overlaid enlargement of representative voltage responses shown in Aa and b. Ca and b, representative voltage responses to current injections of square pulse trains at 200 Hz, before and during TEA application, respectively. Asterisk indicates action potential failure. D, population data showing firing probability of NM neurons as a function of square pulse frequency. The strength of all square pulse trains is 1 nA with a duration of 1 s. Individual square pulse width is 2 ms. Error bar, SEM.

Our current‐clamp recordings were conducted at room temperature (∼24°C). Chickens have a body temperature of ∼42°C. Increasing recording temperature changes the physiology of neurons dramatically (Kushmerick et al. 2006). Our previous study recorded APs at near‐physiological temperature (∼35°C) and we showed a significant improvement of AP kinetics for NM neurons (Hong et al. 2016). For example, AP half‐width was reduced by ∼30% and fall rate increased by ∼40% when increasing the temperature to ∼35°C. Based on these observations, one would expect that NM neurons recorded at higher temperatures could follow stimulations higher than 200 Hz. Indeed, a recent study using current‐clamp recordings at 38–40°C reveals that NM neurons are able to follow square pulse trains up to 500 Hz with good fidelity, whereas the firing probability drops dramatically at 667 Hz (Kuba et al. 2015).

KV3‐containing channels, which belong to the family of KHVA channels, promote the AP repolarizing phase and have been shown to shape the frequency–firing pattern of many neurons (Wang & Kaczmarek, 1998; Johnston et al. 2010; Hong et al. 2016). Based on previous findings, we hypothesized that Kv3‐containing channels play a major role in regulating the rapid firing capability NM neurons. To test this hypothesis, we bath applied TEA (1 mm) to block KV3‐containing channels. Representative responses to 150 Hz square pulse trains before and during TEA application are shown in Fig. 2 Aa and b. The enlargement of two overlaid APs from the same neuron is shown in Fig. 2 B. During TEA application, the majority of NM neurons (10 out of 12 neurons) were still able to follow pulse trains of 150 Hz with 100% fidelity, despite a significant widening of APs (P = 0.0034, Fig. 2 B and D). Correspondingly, no changes in firing probability at 50 or 100 Hz were observed with blockade of KV3‐containing channels (Fig. 2 D). In contrast, we observed reduction in firing probability when increasing the stimulus frequency to 200 or 250 Hz (Fig. 2 Ca and b and D). In response to 200 Hz stimulation, firing probability was reduced by ∼19% on average. Although this reduction is statistically significant (P = 0.0011), nearly half of the recorded neurons (5 out of 12 neurons) showed a reduction of only ∼10% or less in firing probability. These observations with TEA application indicate that KV3‐containing channels contribute partially to higher frequency firing of NM neurons and thus gave rise to an important question: what other factor(s) regulate the ability of NM neurons to follow inputs up to 200 Hz with relatively high fidelity?

Resurgent NaV current of NM neurons

A number of previous studies suggest that I NaR shapes the rapid firing and burst generation in many mammalian neurons (Lewis & Raman, 2014). Therefore, we speculated that NM neurons have I NaR, which is likely one of the factors that regulate the frequency–firing pattern of NM neurons. We first examined whether I NaR is present in NM by using classic voltage‐clamp protocols (Raman & Bean, 1997; Fig. 3). NM neurons were held at −90 mV before giving a depolarizing conditioning step to +30 mV (duration = 10 ms). A transient NaV current (I NaT) was evoked by the conditioning step (Fig. 3 Aa, arrowhead). Afterwards, neurons were repolarized to varying membrane voltages from −70 mV to 0 mV (Δ step = 5 mV). Inward‐going I NaR, which could be eliminated by TTX application, was elicited by repolarization and its amplitude was plotted as a function of repolarizing membrane voltage (Fig. 3 Aa, small arrow, and Ac). The I–V curve of I NaR for NM neurons peaked at −40 mV and presented with the typical ‘V’ shape that largely resembled that of I NaR reported in mammalian neurons (Lewis & Raman, 2014). The maximal amplitude of I NaR is usually less than 1 nA for NM neurons recorded from slice preparation, which is much smaller than the amplitude of I NaT (Fig. 3 Ac). This property is also similar to that observed in other auditory brainstem neurons (Leao et al. 2006; Kim et al. 2010). Therefore, by using the conditioning step of +30 mV at 10 ms, NM neurons show robust generation of I NaR that is present with similar amplitude and voltage dependence properties to mammalian neurons.

Figure 3

Resurgent sodium current (I NaR) of NM neurons at E19–21

Aa and b, Ba and b and Ca and b, representative current traces in response to voltage‐clamp protocols that elicit I NaR shown below traces. The amplitude of the conditioning step is +30 mV in Aa and b, 0 mV in Ba and b and −30 mV in Ca and b. The duration of the conditioning step is 10 ms in Aa–Ca and 100 ms in Ab–Cb. Arrowhead and small arrow in Aa indicate the transient sodium current (I NaT) and I NaR, respectively. Ac, Bc and Cc, population data showing the I NaR amplitude as a function of repolarizing membrane voltage (V MEMBRANE) in response to the conditioning steps shown in Aa and b, Ba and b and Ca and b, respectively. Error bar, SEM.

In mammalian neurons, the I NaR amplitude is dependent on the duration and level of the conditioning step (Raman & Bean, 2001). This is due to two competing inactivation mechanisms of NaV channels: open channel block induced by an intracellular particle and the classic inactivation induced by the ‘gate’ (i.e. cytoplasmic III–IV linker) within the α‐subunit. As demonstrated in cerebellar Purkinje cells, the majority of NaV channels will be open channel blocked if neurons are exposed to a short and more positive depolarization (e.g. an AP). The unbinding of open channel block at moderately negative membrane voltage (e.g. −40 mV) under this condition generates large I NaR. With longer and less positive depolarization, NaV channels will mainly adopt the classic inactivation gate that requires hyperpolarization of membrane voltage to be released, and the generation of I NaR will be minimal. In order to investigate whether I NaR in NM has similar properties, we applied conditioning steps with different durations and levels and compared the I NaR amplitude among the conditions. For example, the duration of +30 mV depolarization was extended from 10 to 100 ms. Fig. 3 Ab shows representative current traces in response to the prolonged conditioning step. The generation of I NaR can still be observed but with a smaller amplitude. Indeed, comparison between the two I–V curves revealed a significant reduction in current amplitude when the conditioning step was elongated to 100 ms (Fig. 3 Ac). Interestingly, we also observed a shift in peak membrane voltage towards the positive direction by ∼5 mV.

Next, we changed the level of the conditioning step to 0 mV and −30 mV while maintaining the same duration (i.e. 10 ms). Fig. 3 Ba and Ca shows representative current traces in response to conditioning steps of 0 mV and −30 mV, respectively. By comparing the I–V curves shown in Fig. 3 Ac–Cc, we observed a decreasing trend (albeit not significant) in I NaR amplitude when the conditioning step was switched from positive to negative voltages. The voltage dependence did not change with different levels of conditioning steps (i.e. all peaked at −40 mV). Finally, we applied a long duration (i.e. 100 ms) conditioning step at non‐positive levels (i.e. 0 and −30 mV), with the prediction that NaV channels will be unlikely to adopt the open channel block and thus elicit minimal I NaR. Indeed, a dramatic reduction in current amplitude was observed in both cases (Fig. 3 Bb and c and Cb and c) along with a shift in voltage dependence by ∼10 mV. Under the condition of −30 mV, the reduction was so great that the typical ‘V’ shape of the I NaR I–V curve was lost (Fig. 3 Cc), which can be explained by the following two factors. First, I NaR, if any, was hardly detectable in response to the conditioning step of −30 mV at 100 ms (Fig. 3 Cb). Instead, steady‐state current was evident following the repolarization. Second, low‐level contaminant noise and/or other channel conductances can obscure small I NaR, which prevents the amplitude of I NaR from being reliably measured (Afshari et al. 2004; Aman et al. 2009).

We further characterized the kinetics of I NaR, which is considered to be another important property. Two variables of kinetics were used based on previous studies: time to peak and decay time constant (tau; Raman & Bean, 1997; Lewis & Raman, 2011). Fig. 4 A shows the calculation of two variables in the current study. Time to peak is defined as the time interval between the onset of repolarization and the I NaR peak. Decay time constant was calculated by fitting a single exponential to the decay phase of I NaR. All calculations were conducted under the conditioning step of +30 mV at 10 ms. Across the population of NM neurons, the average time to peak was 3.97 ± 1.19 ms when measured at the membrane voltage that elicited the maximum current. The membrane voltage for the representative trace shown in Fig. 4 A was −35 mV. The decay time constant was plotted as a function of repolarizing membrane voltage (Fig. 4 B). Again, similar to mammalian neurons, the decay time constant of I NaR increased with more depolarized membrane voltage and was generally larger than the decay variable of I NaT (Ming & Wang, 2003; Rush et al. 2005), which confirmed the slower kinetics of I NaR. In summary, these observations suggest that the use of the open channel blocker, the inactivation mechanisms and the I NaR kinetics of NM neurons closely resemble those of mammalian neurons, properties likely conserved across various species and neural structures.

Figure 4

I NaR kinetics of NM neurons at E19–21

A, representative current trace showing the calculation of time to peak and decay time constant (tau) for I NaR. Decay time constant (tau) is obtained by fitting a single exponential (red trace) to the decay phase of I NaR. The conditioning step is +30 mV at 10 ms. B, population data showing the decay time constant (tau) of I NaR as a function of repolarizing membrane voltage (V MEMBRANE). Error bar, SEM.

Function of resurgent NaV current for NM neurons: experimental results

We hypothesized that I NaR, along with the underlying open channel blocker, plays an important role in promoting the AP firing rates of NM neurons. We predicted that a neuron's activation of open channel blocker would promote NaV channel availability and recovery from depolarization, which is ultimately important for subsequent and rapid AP firing. Therefore, we applied two voltage‐clamp protocols based on previous studies to test our prediction (Raman & Bean, 2001; Patel et al. 2015). In the first protocol, NM neurons were held at −90 mV before being depolarized to +30 mV for 5 ms (Fig. 5). According to our previous observations (see Fig. 3), this conditioned the majority of NaV channels to be occupied by the open channel blocker (referred here as the ‘open channel block state’). The membrane voltage was then set at −65 mV for NM neurons to recover. The recovery time varied from 2 to 50 ms (Δ step = 2 ms). Finally, a depolarization to 0 mV was applied to evoke an I NaT. In the second protocol, the conditioning step was changed to −30 mV for 40 ms, in order to maximize the occupancy of the classic inactivation gate (referred to here as the ‘inactivation state’). Fig. 5 A and B shows representative current traces in response to the two protocols from the same neuron. When neurons were given a short and positive conditioning step, abrupt repolarization to the resting state resulted in an obvious generation of I NaR (arrow in Fig. 5 A), while this was not observed when conditioned to the inactivation state (Fig. 5 B). The generation of I NaR further supported the expectation that the occupancy of the open channel blocker was primary for the first protocol.

Figure 5

I NaR helps increase NaV channel availability and facilitates NaV recovery

A and B, representative current traces in response to voltage‐clamp protocols shown below traces. The conditioning step is +30 mV at 5 ms in A (open channel block state) and −30 mV at 40 ms in B (inactivation state). Δt represents the varying recovery time, increasing from 2 to 50 ms in steps of 2 ms. Arrow in A indicates the generation of I NaR. C, population data showing the NaV channel availability (%) as a function of recovery time. In order to calculate NaV channel availability, a reference pulse to 0 mV was applied to NM neurons (not shown in the figure), and the amplitude of I NaT after the recovery was normalized to this ‘reference amplitude’. Before the normalization, the amplitude of I NaT was first adjusted by subtracting the steady‐state current that remained at the end of the conditioning step. The recovery trajectory is fit by a single exponential, in order to obtain recovery time constant (tau) shown in D. D, population data showing the recovery time constant (tau) under two different condition states. * P < 0.05. Numbers on bars represent sample size. E, representative current traces shown in A and B are normalized and overlaid for recovery time periods of 2 ms (left), 20 ms (middle) and 40 ms (right). Error bar, SEM.

To determine NaV channel availability under the two protocols, we calculated the normalized ratio, which indicates the amount of available NaV channels after the recovery period. To do this, a reference pulse to 0 mV was applied to NM neurons prior to the implementation of the two protocols described above, and the amplitude of I NaT after the recovery was normalized to this ‘reference amplitude’. The normalized ratio was plotted as a function of the recovery time in Fig. 5 C. We observed a clear separation of recovery trajectories between the two conditions, which indicates that the presence of open channel block significantly increased NaV channel availability during the recovery. The recovery trajectory was fit with a single exponential in order to obtain a recovery time constant (tau). The open channel block significantly shortened the recovery time constant, facilitating the recovery of NaV channels (Fig. 5 D). These observations provide supporting evidence for our prediction that the use of the open channel blocker can promote the availability and recovery of NaV channels.

In addition, we found that the distinction between the two states was larger when the amount of recovery time was short. For example, when NM neurons were only given 2 ms to recover, the availability of NaV channels was reduced by ∼67% on average from the open channel block state to the inactivation state (Fig. 5 E, left). But this difference reduced to ∼20% when the recovery time was longer than 20 ms (Fig. 5 E, middle and right). This suggests that the role of open channel block in recovering NaV channels is more critical when the recovery time is limited, which is reminiscent of a highly restricted interspike interval when NM neurons are performing rapid auditory tasks (Warchol & Dallos, 1990; Jones & Jones, 2000).

Function of resurgent NaV current for NM neurons: computational results

We further used a computational model to examine the function of I NaR in NM. This model is designed based on our previous study (Lu et al. 2017), in combination with the I NaR model from Khaliq et al. (2003). The model NM neuron was held at −90 mV before giving a depolarizing conditioning step to +30 mV (duration = 10 ms). An I NaT was evoked by the conditioning step (Fig. 6 Aa, arrowhead). The model NM neuron was then repolarized to varying membrane voltages from −70 mV to 0 mV (Δ step = 5 mV). The model NM neuron displayed slightly smaller I NaR amplitudes (Fig. 6 Aa, small arrow) but comparable voltage dependence properties to our experimental data (Figs 6 Ab and 3 Ac). In order to remove I NaR, we first set the rate constant ε to zero (see Fig. 1 for reference). This modification, however, significantly slowed down the falling phase of I NaT (Fig. 6 B, the ‘0‐I NaR−’ condition, blue trace). The slower falling phase of I NaT is because the Markovian 13‐state NaV‐channel model sets the O→OB transition (with the rate constant ε, see Fig. 1) as a major exit path from the open state (Magistretti et al. 2006). Removing this path resulted in the slower speed of channels exiting the open state and thus led to a slower I NaT falling phase. Therefore, we next increased the rate constant O on to 2.15 ms−1 and the O off to 0.01433 ms−1 to restore the normal decay kinetics of I NaT and amplitude of I NaP, respectively (Fig. 6 B and C, the ‘0‐I NaR+’ condition, red trace). After these two modifications, I NaR was successfully eliminated (Fig. 6 C). Both 0‐I NaR− (blue) and 0‐I NaR+ (red) conditions were used to characterize the spiking activity of model NM neuron without I NaR. The only difference between 0‐I NaR+ and control conditions was the absence of I NaR, while the 0‐I NaR− condition showed both the absence of I NaR and slower I NaT decay kinetics.

Our model NM neuron generated similar voltage responses to square pulse current trains with varying frequencies, as compared to our experimental data. For example, the model NM neuron responded to a 200 Hz current injection with ∼0.6 AP firing probability (Fig. 7 Aa and c). It should be noted that an AP was identified only if the evoked voltage peak exceeded −30 mV. Voltages below this value were considered as failures (Fig. 7 Ac, asterisks). Fig. 7 Ab shows the total underlying NaV current required for AP generation in Fig. 7 Aa. With the current and time scale expanded, we observed a clear but small inward NaV current immediately after a large I NaT (Fig. 7 Ad, arrow). This inward NaV current is reminiscent of I NaR that occurs during the AP repolarizing phase and closely resembles the data reported by previous ‘AP‐clamp’ studies in mammalian neurons (Raman & Bean, 1997, 1999). When we removed I NaR in the model NM neuron, the reduction in firing probability was identical in both 0‐I NaR− and 0‐I NaR+ conditions. Firing probability at 200 Hz was reduced by ∼21% (Fig. 7 Ba–c, asterisks; Fig. 7 Ca–c, asterisks) and the small inward NaV current after I NaT was no longer visible (Fig. 7 Bd and Cd, arrow). This result further confirmed that the small inward current is I NaR and is induced by the mechanism of open channel block. In addition, when we removed both I NaR and KHVA currents, firing probability in response to 200 Hz stimulation was reduced by ∼23% from the control – just slightly larger than the reduction we observed with I NaR removal only (data not shown).

Figure 7

I NaR promotes frequency firing of model NM neuron

Aa–d, Ba–d and Ca–d, simulated voltage and current responses to square pulse current injections of 200 Hz, under the control, 0‐I NaR− (i.e. I NaR removal and slower I NaT) and 0‐I NaR+ (i.e. I NaR removal) conditions, respectively. Aa, Ba and Ca are model output of membrane voltage (V MEMBRANE); enlarged traces shown in Ac, Bc and Cc. Ab, Bb and Cb are model output of NaV currents; enlarged traces are shown in Ad, Bd and Cd. Asterisk indicates action potential failure. Arrows indicate the generation (Ad) or elimination (Bd and Cd) of I NaR.

It should be noted that I NaT generated in the 0‐I NaR− condition was obviously wider than in the 0‐I NaR+ condition, though the same amount of reduction in firing probability was observed for both conditions (Fig. 7 Bd and Cd). This difference is due to the slower decay kinetics of I NaT in the 0‐I NaR− condition that means NaV channels inactivate more slowly (see Fig. 6 B). As a result, APs generated in the 0‐I NaR− condition showed longer half‐width and larger amplitude than in the 0‐I NaR+ condition (Fig. 7 Bc and Cc). In addition, we also observed a difference between two conditions in a small I NaT that failed to generate an AP after a large I NaT (Fig. 7 Bd and Cd, asterisks). In the 0‐I NaR− condition, NaV currents after a large I NaT, the amplitude of which indicates the number of NaV channels recovered after an AP, were present with smaller amplitude than those in the 0‐I NaR+ condition (Fig. 7 Bd and Cd, asterisks). This is because of the slower NaV channel kinetics in the 0‐I NaR− condition and the highly restricted recovery time in response to 200 Hz square pulse trains. Nevertheless, the identical reduction in firing probability in both conditions confirmed that the reduced ability of the model NM neuron to follow inputs at high rates was not due to a slower I NaT but to the absence of I NaR. Together, our modelling results indicate that I NaR helps shape AP firing rate of NM neurons.

Development of resurgent NaV current in NM

We confirmed the generation of I NaR in late‐developing NM neurons (i.e. E19–21), when near‐mature hearing ability is established (Jones et al. 2006). Next, we investigated whether NM neurons in the earlier periods of development have I NaR. Voltage‐clamp experiments were first performed on NM neurons at E14–16, which corresponds to a developmental period when crude hearing ability is just established for chickens (i.e. during hearing onset). To our surprise, we observed robust generation of I NaR at E14–16 when using the conditioning step of +30 mV, 10 ms, with the peak amplitude just slightly smaller than that of late‐developing NM neurons (Fig. 8 A and D). Also similar to late‐developing neurons, the I NaR amplitude was reduced when we changed the level of the conditioning step to either less positive voltages (i.e. 0 and −30 mV) or extended the step duration (i.e. 10 and 100 ms) (Fig. 8 A and B and D–F). Additionally, we observed a shift in the peak of I–V curve in the positive direction by ∼15 mV when we prolonged the duration of the step from 10 to 100 ms. These observations suggest that during hearing onset, the mechanisms of open channel block are established and show near‐mature properties.

Figure 8

I NaR properties of NM neurons at E14–16

A and B, representative current traces in response to voltage‐clamp protocols shown below traces. The strength of the conditioning step is +30 mV (black trace), 0 mV (grey trace) and −30 mV (blue trace) and the duration is 10 ms in A and 100 ms in B. C, population data showing the decay time constant (tau) of I NaR as a function of repolarizing membrane voltage (V MEMBRANE). D–F, population data showing the I NaR amplitude as a function of repolarizing membrane voltage (V MEMBRANE) in response to different conditioning steps. The strength of the conditioning step is +30 mV in D, 0 mV in E and −30 mV in F, and the duration is 10 and 100 ms. Error bar, SEM.

Despite these similarities, we also noticed that some properties of I NaR at E14–16 were different from late‐developing neurons. It should be noted that the following differences are discussed using the conditioning step of +30 mV, 10 ms. Although the I NaR amplitude peaked at −40 mV for both age groups, it reduced more dramatically at older ages when membrane voltage became more positive or more negative (see Figs 3 Ac and 8 D). According to a previous study (Lewis & Raman, 2011), the relative amplitude of I NaR depends on two mechanisms. First, the rate of the open channel block unbinding from the NaV channel α‐subunit, which indicates the affinity between these two particles, and second, after removal, the rate of the α‐subunit entering the classic inactivation or closed states (depending on the voltage). Time to peak is a good index for the first mechanism (i.e. the rate of unbinding), while decay time constant is dependent on both mechanisms. We did not observe a significant difference of time to peak between two age groups (see Fig. 9 D, P = 0.88). Furthermore, we plotted the decay time constant as a function of membrane voltage for E14–16 NM neurons and compared it with E19–21 NM neurons (Figs 8 C and 4 B, respectively). Indeed, E14–16 NM neurons showed a generally larger decay time constant than the older age group. Therefore, we suggest that NaV channels at E14–16 inactivate or close (at the membrane voltage more positive or more negative than −40 mV, respectively) significantly slower after the unbinding of the open channel blocker, resulting in a less dramatic fall‐off of I NaR amplitude and thus a shallower slope of their I–V curve than NaV channels at E19–21. Although additional experiments are required to test this suggestion, we speculate that longer I NaR decay kinetics are partially related to the type of NaV α‐subunit(s) expressed at E14–16 compared to older ages (see below).

Figure 9

I NaR properties of NM neurons at E11–12

A, representative current traces in response to voltage‐clamp protocols shown below traces. The conditioning step is +30 mV at 10 ms. B, population data showing the current amplitude as a function of repolarizing membrane voltage (V MEMBRANE) in response to conditioning step of +30 mV at 10 ms. C, population data showing the decay time constant (tau) of I NaR as a function of repolarizing membrane voltage (V MEMBRANE). D, population data showing time to peak of I NaR at different embryonic ages. * P < 0.05. Numbers on the bars represent sample size. Error bar, SEM.

Next, we performed voltage‐clamp experiments on NM neurons at E11–12, when chickens are not able to respond to sound (i.e. before hearing onset; Jones et al. 2006). In 10 out of 14 recorded neurons, we observed small I NaR when using the conditioning step of +30 mV, 10 ms (Fig. 9 A, note scale). The maximal amplitude of I NaR was slightly above 100 pA, which is dramatically smaller than the current amplitude of the other two age groups (Fig. 9 B). In addition, the voltage dependence of I NaR at E11–12 was very different from the other two age groups, showing an obvious shift in peak voltage (i.e. at −20 mV, Fig. 9 B). Fig. 9 C shows the decay time constant of I NaR for NM neurons before hearing onset, which presented with an increasing trend with more depolarized membrane voltage. Time to peak was 9.54 ± 2.87 ms when measured at the maximal I NaR. This value is significantly larger than those of the other two age groups, indicating a stronger affinity between the open channel blocker and the α‐subunit of NaV channels, and thus slower kinetics of I NaR at E11–12 (Fig. 9 D). Taken together, during the prehearing period, I NaR was present in the majority of NM neurons with preliminary properties. With development, the I NaR amplitude increased, kinetics improved and its voltage dependence shifted towards negative direction. By the time of hearing onset, NM neurons obtained more mature‐like I NaR properties. Table 3 summarizes the developmental changes in amplitude of I NaT, I NaR and I NaP, when measured at membrane voltage of −30 mV. The amplitude of I NaP was measured at the end of 100 ms repolarization after the conditioning step (+30 mV, 10 ms). We observed significant increases in all three current amplitudes for NM neurons as a function of development.

Development of frequency–firing pattern in NM

Based on the functional role of I NaR and its development in NM, we predicted that neurons at the age of E14–16 would show similar frequency–firing pattern with late‐developing neurons (i.e. E19–21), whereas neurons at E11–12 would not be able to follow square pulse frequency as high as the other two age groups, due to their underdeveloped I NaR (see Fig. 9) and KV channels (Hong et al. 2016). To test this prediction, we first applied the aforementioned current commands of square pulse trains to NM neurons at E14–16. As expected, neurons at this age were able to follow square pulse trains of 150 Hz in a one‐to‐one manner (Fig. 10 Aa and c). Firing probability dropped continuously when stimulus frequency was higher than 150 Hz (Fig. 10 Ac). In response to square pulse trains of 200 and 250 Hz, firing probabilities were ∼0.5 and ∼0.25, respectively (Fig. 10 Ab and Ac), which are slightly lower than those of E19–21 NM neurons but these differences are not statistically significant (P = 0.49 and 0.32, respectively). When the stimulus frequency was 300 Hz, E14–16 NM neurons showed a single onset spike similar to late‐developing neurons (data not shown).

Figure 10

Frequency–firing pattern of NM neurons at E14–16 and E11–12

Aa and b, representative voltage responses recorded from an E16 NM neuron to current injections of square pulse trains at 150 and 200 Hz, respectively. Ac, population data showing firing probability of E14–16 NM neurons as a function of square pulse frequency. The strength of square pulse trains is 1 nA with a duration of 1 s. Individual square pulse width is 2 ms. Ba and b, representative voltage responses recorded from an E12 NM neuron to current injections of square pulse trains at 70 Hz and 150 Hz, respectively. Bc, population data showing firing probability of E11–12 NM neurons as a function of square pulse frequency. The strength of square pulse trains is 1 nA with a duration of 1 s. Individual square pulse width is 5 ms. Error bar, SEM.

For NM neurons at E11–12, individual square pulse width was extended to 5 ms because neurons at this age are present with average AP half‐width of ∼4.6 ms (Hong et al. 2016). We found that NM neurons at E11–12 were able to follow square pulse trains with good fidelity up to 70 Hz (Fig. 10 Bc). Fig. 10 Ba shows a representative E12 neuron that was able to reliably generate an AP in response to each pulse of the 70 Hz stimulation. When stimulus frequency was increased to 100 Hz or higher, however, firing probability dropped steeply below 0.5 and in response to 150 Hz stimulation, NM neurons at E11–12 only generated a single onset spike followed by membrane oscillations at a depolarized level (Fig. 10 Bb and c). This is in stark contrast to the other two age groups, which could follow 150 Hz square pulse trains in a reliable manner (see Figs 2 D and 10 Ac). Taken together, NM neurons at E14–16 showed a frequency–firing pattern closely resembling their late‐developing counterparts, while the ability of NM neurons at E11–12 to follow high‐frequency stimulations was markedly limited. These results are similar to our previous data, which show that E11–12 NM neurons are most responsive to low‐frequency (<40 Hz) sinusoidal current injections (Hong et al. 2016).

Expression of NaV1.6 channels

NaV1.6 channels are extensively expressed in the mammalian central nervous system and act as a predominant carrier for I NaR (Eijkelkamp et al. 2012; Lewis & Raman, 2014). Therefore, we explored whether NaV1.6 channels are expressed in developing NM, using an antibody specifically recognizing chicken NaV1.6 (Kuba et al. 2006, 2010, 2014). In late‐developing NM neurons (i.e. E21), strong NaV1.6 immunoreactivity was observed as bright punctate segments (Fig. 11 Aa and b). Because E21 NM neurons are mostly adendritic (Jhaveri & Morest, 1982a), double staining of NaV1.6 and neurofilament further demonstrated that NaV1.6 is localized in neurofilament‐stained NM axons that can be traced back to the cell bodies (Fig. 12). These NaV1.6‐containing segments closely resembled the characterized distribution pattern of NaV1.6 in chicken auditory brainstem as reported by Kuba et al. (2010, 2014) and possibly represented the axon initial segments (AIS) and nodes of Ranvier. In contrast, the NaV1.6‐containing segments were absent in NM at E15 (Fig. 11 Ba and b) and E11 (Fig. 11 Ca and b). In addition, NM cell bodies contained a low level of NaV1.6 immunoreactivity at E21, which became increasingly less distinct from E21 to E15 and from E15 to E11.

Figure 11

NaV1.6 distribution in developing NM

NaV1.6 immunoreactivity at E21 (A), E15 (B) and E11 (C). Left (Aa, Ba and Ca) and right (Ab, Bb and Cb) columns are low‐ and high‐magnification confocal images, respectively. Dashed lines indicate the boundary of NM. NM, nucleus magnocellularis. Scale bars: 50 μm in Ca (left column), 10 μm in Cb (right column).

Figure 12

Axonal localization of NaV1.6 immunoreactivity at E21

Double staining of NaV1.6 and neurofilament in E21 NM. Arrows indicate a distinct segment containing NaV1.6, which overlaps with neurofilament‐stained axon that can be traced back to the cell body. NM, nucleus magnocellularis. Scale bars: 20 μm for all panels.

Discussion

Auditory brainstem neurons of vertebrates fire phase‐locked APs at high rates with remarkable fidelity. We show here that late‐developing NM neurons fire reliable APs to square‐pulse current injections up to 200 Hz. In addition to the recognized role of KHVA channels in this process, we also report that I NaR of NaV channels is an equally important component that operates synergistically with KHVA channels to enable rapid AP firing in NM, an evolutionarily conserved process between birds and mammals likely promoting similar hearing functions. To our surprise, small I NaR responses were present during a prehearing period (E11–12). At hearing onset (E14–16), I NaR properties closely resemble late‐developing NM neurons (E19–21), despite developmental refinement in NaV channel protein expression patterns. In line with these results, NM neurons at E14–16 showed comparable frequency–firing ability to their late‐developing counterparts, whereas NM neurons at E11–12 are most responsive to lower‐frequency stimulations.

Factors regulating AP firing patterns in NM

KHVA channels are a common regulator of AP kinetics (Rudy & McBain, 2001; Johnston et al. 2010). In the auditory brainstem of both birds and mammals, KV3‐containing channels are abundantly expressed (Wang et al. 1998; Parameshwaran et al. 2001; Parameshwaran‐Iyer et al. 2003). Blockade or knockout of KV3 channels results in wider APs that undermine the ability to generate APs at high rates (Wang et al. 1998; Klug & Trussell, 2006). We report a similar result here (Fig. 2). In addition, a recent study on Xenopus oocytes shows that KV3.1 subunits are able to produce resurgent potassium current during repolarization, which provides an additional repolarizing drive to the membrane (Labro et al. 2015). This property is not due to the open channel blocker but to the unique gating kinetics of KV3.1 subunits, and it facilitates the termination of APs and thus likely promotes high‐frequency repetitive firing. Nevertheless, it should be noted that 1 mm TEA used in the current study can also block a small portion of KV7‐containing channels and calcium‐activated BK channels (Johnston et al. 2010). KV7‐containing channels are one of the KLVA channels with slow activation kinetics (Johnston et al. 2010). They contribute minimally in NM and their immunoreactivity is minimal compared to KV3 and KV1 (Kuba et al. 2015). Due to their weak expression, blockade of KV7‐containing channels in NM neurons does not induce significant changes in AP properties (Kuba et al. 2015). In contrast, BK channels play an important role in regulating AP kinetics in many other neurons (Kimm et al. 2015). However, a previous study in NM reported no change in KV current when external calcium was replaced by cobalt ions, suggesting minimal contribution of calcium‐activated potassium current to the total current (Koyano et al. 1996). Therefore, KV3‐containing channels are probably the primary targets of 1 mm TEA applied in our experiments.

Another factor that regulates AP firing pattern is fast inactivation kinetics of NaV channels (i.e. the steep decay slope of I NaT) observed in auditory brainstem neurons (Ming & Wang, 2003; Hong et al. 2016). As demonstrated by double‐pulse experiments, a second I NaT can fully recover within 5 ms following the initial pulse (Leao et al. 2005; unpublished observation in NM), which explains in the current study why NM neurons are able to reliably follow the stimulus frequency at 150 Hz (interspike interval = 6.7 ms, Fig. 2 D) after blockade of KHVA channels. Finally, KLVA channels with fast activation kinetics, such as KV1‐containing channels, work as another regulator of AP firing by controlling the time constant of passive membrane properties, shaping the speed of membrane voltage changes (Klug & Trussell, 2006).

I NaR is an additional factor in regulating AP firing rates of NM neurons. The open channel block promotes NaV channel availability and recovery immediately after brief depolarization in NM, a result consistent with previous studies (Raman & Bean, 2001; Patel et al. 2015). However, we report two major differences. First, NaV channel availability is much higher in our study at any given recovery time period (Fig. 5 C). This is likely because our recordings were made from intact neurons in brainstem slices that have significantly more neuron‐specific NaV channels. Previously reported data were obtained from the soma of dissociated Purkinje cells or from HEK cells transfected with NaV channels. Second, the distinction in recovery time constant between the open channel block and classic inactivation state is smaller in our study (∼1 ms versus > 5 ms, Fig. 5 D). This may be attributed to relatively fast kinetics of I NaT in auditory brainstem neurons. However, the extent to which the open channel block or the classic inactivation gate contributes to rapid I NaT kinetics in NM is unclear.

‘Real‐time’ I NaR appears to be a relatively small inward NaV current during repolarization from an initial I NaT, as shown by our model NM neuron (Fig. 7 Ad). This observation closely resembles experimental data when AP waveforms are used as voltage commands and the dynamic changes in NaV current are documented (i.e. AP‐clamp method; Raman & Bean, 1997, 1999). As seen from studies that utilized this technique, the I NaR provided a small depolarizing drive immediately after an AP, important for subsequent AP firing (Raman & Bean, 1997). However, AP‐clamp is susceptible to various technical limitations (e.g. space clamp errors), and as a result the majority of these studies were conducted on dissociated neurons (Raman & Bean, 1997; Do & Bean, 2003). Nevertheless, there are successful slice recordings with AP clamp from adendritic mesencephalic trigeminal neurons (Enomoto et al. 2006) Given the fact that the majority of late‐developing NM neurons are also adendritic, AP‐clamp experiments could provide valuable insight into the biological relevance of I NaR. There are still technical limitations (e.g. sample rate, kinetics, immature dendritic NM neurons, etc.) and therefore we employed a computational model to better address its functional significance. Indeed, removal of I NaR reduced AP firing rate at 200 Hz for our model NM neuron (Fig. 7). Yet, the effects on NaV channels after the knockdown of open channel blocker remain to be determined, especially regarding the kinetics of I NaT under experimental condition. These effects are dependent on the types of β‐subunits expressed in NM neurons and their specific interactions with NaV α‐subunits (Qu et al. 2001; Aman et al. 2009; Bant & Raman, 2010). Therefore, our computational model of NM neuron is subject to future improvements when more information about molecular substrates for the open channel blocker in NM is obtained.

The inner ear of chickens only responds to loud sound (>80 dB) and afferent ganglion neurons present with poor frequency selectivity at E14–16 (Jones et al. 2006). It is surprising at this age that I NaR properties are relatively established, except for decay kinetics, which are due in part to underdeveloped NaV α‐subunit (Fig. 8, see below). I NaR properties at E11–12 – a developmental period when the auditory system does not respond to sound and is considered ‘prehearing’ – differ greatly from those of the other age groups (Fig. 9). I NaR development in NM parallels the development of frequency–firing pattern (Fig. 10), along with maturation of intrinsic AP properties and KV channel conductances as we previously reported (Hong et al. 2016). NM neurons at E14–16 generate APs as fast and reliably as late‐developing neurons. Additionally, properties of KLVA and KHVA currents are underdeveloped at E11–12 but become more mature‐like around hearing onset. Taken together, NM neurons have near‐mature ion channel properties around hearing onset, suggesting a priming period during hearing development for establishing mature auditory functions.

I NaR at E11–12, though small, may be important for shaping NM's AP firing pattern. In response to low‐frequency (5–10 Hz) sinusoidal current injections, NM neurons at this age generate short bursts of APs per cycle, despite the fact that their overall low KV channel conductances are not sufficient to work against the largely depolarized membrane voltage during bursts (Hong et al. 2016). I NaR also contributes to burst firing in other non‐auditory neurons (Enomoto et al. 2006, 2007). We speculate that I NaR at E11–12, albeit reduced, helps generate burst firing for immature NM neurons, but its functional significance at this age is unclear.

Anatomically, we detected strong NaV1.6 expression at a late‐developing stage (E21), which was dramatically reduced at hearing onset (E14–16) and prehearing (E11–12) periods (Fig. 11). Additionally, the majority of NaV1.6 channels in late‐developing neurons are located along the axons, while this distribution pattern is not evident for the other age groups. The NaV1.6‐positive punctate segments in late‐developing NM neurons are likely to be the AIS and the nodes of Ranvier, respectively, though future immunocytochemistry with double staining of ankyrin G, a scaffold protein that marks the AIS, is needed to confirm this speculation (Kuba et al. 2014). The presence of I NaR at E11–16 suggests that additional NaV subtypes may contribute to I NaR during development of NM neurons. The following evidence supports this suggestion. First, strong expression of NaV1.2 channels is observed at E11–15 in NM's postsynaptic target nuclei (i.e. nucleus laminaris) (Kuba et al. 2014). Second, studies in spinal sensory neurons report the generation of I NaR from different NaV subtypes, including NaV1.2 channels (Rush et al. 2005; Jarecki et al. 2010).

Resurgent NaV current is conserved across species and structures

I NaR has been reported as a conserved property in numerous neuronal types in the mammalian cerebellum, cortex, brainstem and spinal cord (Afshari et al. 2004; Cummins et al. 2009; Lewis & Raman, 2014). As for the mammalian auditory system, I NaR has been found at the calyx of Held and its postsynaptic target – the medial nucleus of trapezoid body (Leao et al. 2005; Kim et al. 2010). The only report of I NaR beyond the scope of mammals comes from chicken Purkinje cells (Lewis & Raman, 2011). Our results from the current study further confirm its contribution in the chicken auditory brainstem. Here, the AP firing pattern of NM neurons (and the analogous bushy cells of the mammalian anteroventral cochlear nucleus) is essential for the encoding of behaviourally relevant acoustic cues. Central to this are established and specialized structural and functional features shared across species (Carr & Soares, 2002; Koppl, 2009; Grothe et al. 2010). The report of I NaR in the current study strongly suggests yet another physiological feature shared across species within the central nervous system.

Despite the distribution of I NaR in the central nervous system (Lewis & Raman, 2014), its underlying molecular substrates are unclear. The NaV β4‐subunit has been identified as one important open channel blocker in cerebellar Purkinje cells, granule cells and dorsal root ganglia neurons (Grieco et al. 2005; Bant & Raman, 2010; Barbosa et al. 2015). Interestingly, the amino acid sequence of the β4‐subunit is also conserved across species, from frogs to mammals, raising the possibility of its being the open channel blocker for avian NM neurons (Lewis & Raman, 2011). Nevertheless, questions are raised of whether the β4‐subunit is the only possible open channel blocker. For example, both perirhinal and entorhinal pyramidal neurons show robust I NaR but not β4 expression at high levels, possibly indicating multiple molecular substrates of I NaR (Castelli et al. 2007; Nigro et al. 2012). In addition, FGF14, an intracellular protein and a member of the fibroblast growth factor homologous factors (FHFs), has been proposed as another key player in open channel block in Purkinje cells (Yan et al. 2014). A recent study on dorsal root ganglia neurons demonstrated that another member of the FHF family, FHF2, plays an important regulatory role for I NaR (Barbosa et al. 2017). Interestingly, two isoforms of FHF2 have opposite effects on the amplitude of I NaR. As for NM neurons, it should be noted that differences in I NaR across development, especially at E11–12, might be attributed not only to different expression of NaV α‐subunits, but also to changes in the open channel blocker molecular substrates themselves. Therefore, what the underlying open channel blockers for NM neurons are and how the developmental expression of these blockers shapes I NaR maturation are interesting questions for future studies.

Additional information

Competing interests

The authors declare no competing financial interests.

Author contributions

All authors have approved the final version of the manuscript and agree to be accountable for all aspects of the work. All persons designated as authors qualify for authorship, and all those who qualify for authorship are listed. HH, TL, XW, YW and JTS designed the study. HH performed electrophysiology experiments at Northwestern University. TL performed computational modeling experiments at Northwestern University. XW and YW performed immunocytochemical experiments at Florida State University. HH, TL, XW, YW and JTS analyzed/interpreted data and wrote the manuscript.

Funding

This research was supported by the National Institute on Deafness and Other Communication Disorders (NIDCD) DC013841 (J.T.S.) and the Hugh Knowles Hearing Research Center (J.T.S.).