Characterization of Na⁺ currents regulating intrinsic excitability of optic tectal neurons

The intrinsic excitability of Xenopus tectal neurons is correlated with the amplitude of voltage-gated Na⁺ currents

Principal tectal neurons, which are the main recipients of retinal input in the developing visual circuit of tadpoles, must adapt their intrinsic excitability to maintain neuronal function in response to changes in synaptic input as a result of both developmental circuit reorganization and sensory experience (Aizenman et al, 2003; Pratt & Aizenman, 2007; Ciarleglio et al, 2015; Busch & Khakhalin, 2019). Although studies strongly implicate the regulation of Na⁺ currents with changes in tectal neuron excitability (Aizenman et al, 2003; Pratt & Aizenman, 2007; Ciarleglio et al, 2015), there is also evidence that the regulation of K⁺ currents plays a role in this process (Hamodi & Pratt, 2014; Ciarleglio et al, 2015). As such, the molecular mechanisms by which Na⁺ currents are regulated to control intrinsic excitability, and the degree to which regulation of K⁺ currents contributes to this control of excitability, are not fully understood.

To examine the relationship between intrinsic excitability of tectal neurons and the regulation of voltage-gated Na⁺ and K⁺ currents, we performed whole-cell recordings on 61 deep-layer principal tectal neurons from tadpoles at developmental stage 49 using a whole brain ex vivo preparation (Fig 1A). We selected stage 49 tadpoles because the biophysical properties of their tectal neurons is heterogenous at this developmental stage, with most neurons exhibiting low intrinsic excitability but with some more highly excitable neurons still observed (Ciarleglio et al, 2015). As expected, we observed a broad range of values in the maximum number of spikes that tectal neurons could generate (1–8), with most neurons spiking once or twice (Fig 1B). For each neuron, we also measured the peak current amplitudes for the voltage-gated Na⁺ current, the transient K⁺ current (K_T), and the steady-state K⁺ current (K_SS) (Fig 1C). Importantly, these biophysical properties matched previous reports for stage 49 tectal neurons (Aizenman et al, 2003; Pratt & Aizenman, 2007; Ciarleglio et al, 2015; James et al, 2015) and showed a wide range of excitability levels which allowed us to investigate the cellular basis for heterogeneity of intrinsic excitability within tectal neurons.

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Figure 1. Intrinsic excitability of Xenopus tectal neurons is correlated with the amplitude of voltage-gated Na⁺ currents.

(A) Diagram shows a Xenopus tadpole illustrating whole-cell recordings from a neuron of the optic tectum that receives innervation from visual and mechanosensory inputs. (B) Left: example current-clamp recordings showing the spiking response of two stage 49 tectal neurons held at −65 mV to a 50 pA current injection, illustrating the range of responses observed. Middle: plot shows the number of spikes elicited in response to 0–200 pA current injections for all cells analyzed (grey), and the average response (red, mean ± SD). Right: maximum number of spikes (median = 2 spikes, IQR 1–3 spikes, n = 61 cells). (C) Left: example voltage-clamp recording from a tectal neuron held at −65 mV in response to a series of depolarizing steps (−65 to +25 mV). Recording is leak subtracted to show only active currents. Middle: I–V plot shows average current amplitude for the Na⁺, the transient K⁺ (K_T) and the steady state K⁺ currents (K_SS). Right: peak current amplitudes (Na⁺: 341.4 ± 176.1 pA; K_T: 666.9 ± 281.0 pA; K_SS: 517.1 ± 251.2 pA; n = 61 cells). (D, E, F) Plots show Pearson correlations between intrinsic excitability (max. number of spikes) and (D) the amplitude of the Na⁺ current, (E) the transient K⁺ current, and (F) the ratio of the Na⁺ current to transient K⁺ current. r values are Pearson correlation coefficients (***P = 0.0002; ****P < 0.0001). The complete Pearson correlations matrix showing the relationships between all biophysical properties measured is presented in Fig S1.

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Source Data for Figure 1[LSA-2023-02232_SdataF1.xlsx]

To measure the relationship between intrinsic excitability and other intrinsic properties of tectal neurons, we performed a multivariate analysis and calculated Pearson pairwise correlations between each of the biophysical properties measured (the complete correlation matrix is shown in Fig S1). When comparing voltage clamp currents with the maximum number of spikes, we found a strong correlation with the peak amplitude of the Na⁺ current (R 0.47; Fig 1D), whereas the peak amplitude of the transient K⁺ current was weakly correlated with the maximum number of spikes (R −0.08; Fig 1E). Accordingly, we also found a strong correlation between the maximum number of spikes and the Na⁺ to transient K⁺ current ratio (R 0.59; Fig 1F). We restricted our measurements to peak current amplitude, and not activation and inactivation kinetics because of possible space-clamp issues, consistent with previous studies (Aizenman et al, 2003; Pratt & Aizenman, 2007). These data indicate that the regulation of Na⁺ currents is an important mechanism to control the intrinsic excitability of tectal neurons, with an increased ratio of Na⁺ to K⁺ currents present in more excitable tectal neurons. In further support of this idea, we measured significant correlations between the peak amplitude of the Na⁺ current and characteristics of the first spike including rate of rise and spike width; but little correlation between characteristics of the first spike and the amplitude of K⁺ currents (Fig S1). Taken together, these data provide further evidence that Na⁺ currents are a key determinate of the intrinsic excitability of tectal neurons.

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Figure S1. Relationship between biophysical properties of Xenopus tectal neurons at developmental stage 49.

Pearson correlation matrix showing the relationship between cell properties for 61 stage 49 tectal neurons. Properties analyzed included neuronal intrinsic excitability (Max # spikes), characteristics of the first spike (spike threshold, spike amplitude, the rate of rise [20–80%], and spike width at 50%), and peak amplitudes for the voltage-gated fast Na⁺ current (Na⁺), transient potassium current (K_T), and steady state potassium current (K_SS). For 40 cells, we measured spike accommodation (Spike accom. = amplitude spike 1/amplitude spike 2), interspike interval (interstimulus intervals [ISI] = time peak of spike 1 − time peak of spike 2). For 22 cells, we measured interspike interval accommodation (ISI accom. = ISI spike 2–3/ISI spike 1–2). Values are Pearson correlation values (r). Bolded values indicate significant Pearson correlations (Bonferroni-corrected P-value < 0.05).

Expression of Na⁺ channel subtypes are differentially regulated in the Xenopus optic tectum during key developmental time windows and in response to changes in network activity

How are voltage-gated Na⁺ currents regulated to control intrinsic excitability? Na⁺ channels are comprised of alpha and beta subunits, the alternate expression of which confers Na⁺ channels with distinct cellular expression profiles, subcellular localization, conduction properties, and responses to Na⁺ channel activators and inhibitors (Savio-Galimberti et al, 2012). The alpha subunits (Na_v1.1–Na_v1.9) are the pore-forming components of Na⁺ channels, whereas the accessory beta subunits (Na_v1β–Na_v4β) can regulate expression, cellular localization, and gating properties of Na⁺ channels (O’Malley & Isom, 2015). A search of the X. laevis genome (Xenbase: X. laevis version 9.2 on JBrowse) revealed that X. laevis expresses alpha subunit genes encoding for the brain-expressed voltage-gated Na⁺ channel subtypes Na_v1.1, Na_v1.2, and Na_v1.6. Xenopus also expresses the accessory beta subunit Na_v4β that has a well-described role in regulating neuronal excitability by mediating resurgent Na⁺ currents (Bant & Raman, 2010). If the regulation of Na⁺ channel expression is a mechanism to control voltage-gated Na⁺ currents and intrinsic excitability, then we would predict that the expression of individual Na⁺ channel genes would correlate with changes in the intrinsic excitability of tectal neurons. Because the intrinsic excitability of tectal neurons is regulated across tectal circuit development and increased in response to a short-term patterned sensory experience (Aizenman et al, 2003; Pratt & Aizenman, 2007; Ciarleglio et al, 2015), we used qRT-PCR to quantify the expression of Na_v1.1, Na_v1.2, Na_v1.6, and Na_v4β subunits in the optic tectum in these conditions.

As the tectal circuitry matures between developmental stages 42 and 49, the intrinsic excitability of tectal neurons peaks at stage 46 congruent with an overall increase in the level of excitatory synaptic input at this stage of circuit development (Fig 2A) (Pratt & Aizenman, 2007). If the expression levels of individual Na⁺ channel genes correlate with changes in neuronal intrinsic excitability, we would predict increased expression at developmental stage 46. Interestingly, we found that the expression levels of Na_v1.1 and Na_v1.6 in the optic tectum were regulated developmentally, but with differing developmental expression profiles (Fig 2B and Table S1). We found that the expression levels of Na_v1.1 increased with development. We measured a ∼twofold increase in Na_v1.1 expression levels between stage 42 and stage 49 (st 42: 1.00 ± 0.12, st 49: 1.84 ± 0.31; P = 0.0151), but no significant change between stages 42 and stage 46 (st 46: 1.57 ± 0.25; P = 0.0535) or between stage 46 and stage 49 (P = 0.2106). Although this increase in Na_v1.1 levels may signal maturation of neurons, it does not correlate with developmental changes in intrinsic excitability. In contrast, Na_v1.6 expression levels peaked at developmental stage 46, when intrinsic excitability is highest. Expression of Na_v1.6 was increased ∼twofold between stage 42 and stage 46 (st 42: 1.00 ± 0.09, st 46: 1.71 ± 0.22; P = 0.0350), with the level of expression at stage 49 being not significantly different from stages 46 (st 49: 1.40 ± 0.32; P = 0.1932) or stages 42 (P = 0.1932). Unlike the developmental regulation of Na_v1.1 and Na_v1.6 channels, the expression levels of Na_v1.2 and Na_v4β remained largely unchanged across development (Fig 2B). These data suggest that Na_v1.1 channel expression increases with the functional maturation of tectal neurons, whereas the expression of Na_v1.6 channels more closely matches developmental changes in intrinsic excitability.

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Figure 2. The expression of Na⁺ channel genes is regulated with developmental and homeostatic changes in neuronal intrinsic excitability.

(A) Schematic illustrates how tectal neurons homeostatically adapt intrinsic excitability in response to changing excitatory synaptic drive across development and in response to 4 h exposure to enhanced visual stimulation (EVS) to maintain a broad dynamic range and, thereby, conserve input–output function as the tectal circuitry changes (Aizenman et al, 2003; Pratt & Aizenman, 2007; Ciarleglio et al, 2015). (B) Expression levels of selected brain-expressed voltage-gated Na⁺ channel alpha and beta subunits in the optic tectum at key development stages. Developmental stages were selected to sample immature tectal neurons with low excitability (stage 42), highly excitable tectal neurons undergoing experience-dependent circuit remodeling (stage 46), and mature tectal neurons with low excitability (stage 49). The expression of Na⁺ channel alpha and beta subunits genes was normalized to a housekeeper (RSP13), before determining the fold change in expression from developmental stage 42 (Na_v1.1 stage 42 versus stage 49, P = 0.0151; Na_v1.6: stage 42 versus stage 46, P = 0.0350. N = 3 experiments with RNA isolated from 10 tadpoles). See Table S1 for comparisons of expression levels between all developmental stages. (C) Expression levels of selected Na⁺ channel alpha and beta subunits in the optic tectum of stage 49 tadpoles with or without 4 h exposure to EVS, which triggers a homeostatic increase in neuronal excitability. Expression is shown as the fold change in normalized Na⁺ channel alpha and beta subunit gene expression compared with naïve stage 49 control tadpoles (Ctrl versus EVS: Na_v1.1, P = 0.0.108; Na_v1.6, P = 0.0135; Na_v4β, P = 0.0434. N = 3 experiments with RNA isolated from 10 tadpoles).

To test whether observed changes in Na⁺ channel expression levels were simply a function of tectal neuron development, or whether changes in Na⁺ channel expression represents a mechanism for the homeostatic regulation of in tectal neuron excitability, we quantified the expression of Na⁺ channel genes in the optic tectum of stage 49 tadpoles exposed to 4 h of enhanced visual stimulation (EVS). EVS is a well-established method for triggering increased intrinsic excitability of tectal neurons as a homeostatic response to decreased excitatory synaptic drive (Fig 2A) (Aizenman et al, 2003; Ciarleglio et al, 2015). Critically, we found that Na_v1.6 expression levels, and to a lesser extent Na_v1.1 expression levels, were increased in response to EVS (Fig 2C). The level of Na_v1.6 expression was increased ∼fourfold in the optic tectum of tadpoles exposed to EVS compared with naïve stage 49 controls (Ctrl: 1.00 ± 0.18, EVS: 3.80 ± 1.13; P = 0.0135). We also found a ∼threefold increase in the expression of Na_v4β (Ctrl: 1.00 ± 0.07, EVS: 3.21 ± 1.31; P = 0.0434), whereas a more modest ∼twofold increase in the level of Na_v1.1 expression was observed in the optic tectum of EVS exposed tadpoles (Ctrl: 1.00 ± 0.23, EVS: 1.71 ± 0.13; P = 0.0108), whereas Na_v1.2 expression levels were unchanged between control tadpoles and EVS-exposed tadpoles (Ctrl: 1.00 ± 0.13, EVS: 1.00 ± 0.09; P = 0.9460). Given the well-established link between Na_v1.6 and Na_v4β expression and the regulation of resurgent Na⁺ currents and intrinsic excitability (Grieco et al, 2005; Bant & Raman, 2010; Lewis & Raman, 2011), these data provide evidence that tectal neurons homeostatically control their intrinsic excitability by regulating the amplitude of Na⁺ currents through changes in Na_v1.6 gene expression levels, and perhaps via the regulation of a resurgent Na⁺ current. Our data also show that up-regulation of Na_v1.1 expression contributes to the increased amplitude of Na⁺ currents as intrinsic excitability is homeostatically increased. However, the modest up-regulation of Na_v1.1 compared with Na_v1.6, and the fact that Na_v1.1 expression levels were most elevated at stage 49 when tectal cells are less excitable, would indicate that their role in regulating excitability may be less important compared with Na_v1.6 channels.

Tectal neurons express distinct fast and persistent Na⁺ currents that are regulated with homeostatic changes in intrinsic excitability across development and in response to EVS

Voltage-gated Na⁺ channels carry both fast and persistent Na⁺ currents, which have distinct roles in action potential generation and the repetitive firing of neurons (French et al, 1990; Raman et al, 1997; Magistretti et al, 2006). The fast Na⁺ current mediates the upstroke of action potentials before becoming rapidly inactivated (Hodgkin & Huxley, 1952; Stuart & Sakmann, 1994; Martina & Jonas, 1997). When Na⁺ channels fail to fully inactivate, even with prolonged depolarization, the resulting sustained current is termed a persistent Na⁺ current (French et al, 1990). Because the persistent Na⁺ current is a depolarizing current that occurs at a subthreshold voltage range, it can amplify a neuron’s response to synaptic input and enhance the neurons capacity to fire repetitively (Bant & Raman, 2010; Chen et al, 2011). We therefore hypothesized that tectal neurons modulate fast and persistent Na⁺ currents to achieve homeostatic changes in intrinsic excitability. Although fast Na⁺ currents are known to be modulated with changes in excitability in Xenopus tectal neurons (Aizenman et al, 2003; Pratt & Aizenman, 2007), the relative contribution of fast and persistent Na⁺ currents to changes in excitability has not been directly examined.

To determine whether fast and persistent Na⁺ currents are regulated with changes in intrinsic excitability that occur across development (refer to diagram in Fig 2A), we performed whole-cell recordings on tectal neurons from tadpoles at developmental stages 42, 46, and 49. Na⁺ currents were isolated in voltage-clamp recordings using a Tris-based internal saline solution to block outward K⁺ currents (Aizenman et al, 2003), which revealed distinct fast and persistent voltage-gated Na⁺ currents (Fig 3A). When we compared peak amplitudes of the fast and persistent Na⁺ currents across development, we observed a transient peak for fast and persistent Na⁺ currents at stage 46 (Fig 3B and C), congruent with the increased intrinsic excitability at this developmental stage (Pratt & Aizenman, 2007), and with changes in Na⁺ channel expression observed in this study.

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Figure 3. Tectal neurons express fast and persistent voltage-gated Na⁺ currents that are regulated with developmental and homeostatic changes in intrinsic excitability.

(A) Example voltage-clamp recordings from a tectal neuron held at −65 mV in response to a series of 150 ms depolarizing steps (−65 to 25 mV) using a Tris-based internal saline solution, which blocks outward K⁺ currents to reveal distinct fast and persistent Na⁺ currents. Recordings are leak subtracted to show only active currents. Fast (I_NaF) and persistent (I_NaP) Na⁺ currents are indicated on the recording. (B, C) Quantifications of peak amplitudes for the fast and persistent Na⁺ currents. (B) Fast Na⁺ current ([in pA] Stage 42: 157.7 ± 32.5; Stage 46: 197.3 ± 64.4; Stage 49: 143.1 ± 60.3; Stage 49 + enhanced visual stimulation [EVS]: 197.4 ± 50.9). (C) Persistent Na⁺ current ([in pA] Stage 42: 59.8 ± 12.5; Stage 46: 80.7 ± 37.3; Stage 49: 56.3 ± 22.3; Stage 49 + EVS: 73.4 ± 29.0). (D) Example resurgent current recording from a tectal neuron held at −65 mV that was hyperpolarized to −90 mV for 500 ms before stepping to +30 mV for 15 ms to open voltage-gated Na⁺ channels. Resurgent Na⁺ currents were then recorded by a repolarizing step to −30 mV for 200 ms, which revealed distinct tail resurgent (R_Tail), resurgent (R) and persistent resurgent (R_Persistent) currents. Recordings were obtained using a Tris-based internal saline solution to block outward K⁺ currents, and leak subtracted to show only active currents. (E) Averaged resurgent current traces obtained from tectal neurons between developmental stages 42–49, and at stage 49 after exposure to 4 h of EVS. (F) Peak resurgent current amplitude across development and in response to 4 h of EVS ([in pA] Stage 42: 55.9 ± 18.6; Stage 46: 74.3 ± 22.8; Stage 49: 60.0 ± 25.7; Stage 49 + EVS: 78.4 ± 26.9). Groups were compared using a Welch’s ANOVA test with Dunnett T3 test for multiple comparisons. (D) n values are shown in (D).

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Source Data for Figure 3[LSA-2023-02232_SdataF3.xlsx]

We also recorded Na⁺ currents from stage 49 tectal neurons after the exposure of tadpoles to 4 h of EVS to trigger a homeostatic increase in the excitability of tectal neurons. When we compared the amplitude of the fast and persistent Na⁺ currents in tectal neurons of EVS-exposed stage 49 tadpoles with naïve stage 49 tadpoles, we found a significant increase in both currents (Fig 3B and C). Thus, we find a correlation between changes in excitability, Na⁺ current amplitude, and changes in Na⁺ channel expression levels.

Tectal neurons express resurgent Na⁺ currents that are regulated with homeostatic changes in intrinsic excitability across development and in response to EVS

Voltage-gated Na⁺ channels can also generate a resurgent Na⁺ current that is known to facilitate repetitive firing in many types of neurons (Raman et al, 1997; Cummins et al, 2005; Theile & Cummins, 2011; Browne et al, 2017). The resurgent Na⁺ current is a subthreshold depolarizing current that results from a distinctive gating mechanism, typically by open-channel block mediated by accessory proteins including Na_v4β (Grieco et al, 2005; Bant & Raman, 2010). Open-channel block is relieved at subthreshold potentials, generating a resurgent current that promotes firing. Because we had observed that exposure to EVS triggered an up-regulation of Na_v4β in the optic tectum, we asked if tectal neurons express a resurgent Na⁺ current and whether this resurgent Na⁺ current is regulated with homeostatic changes in the excitability of tectal neurons.

We followed established protocols for measuring resurgent currents (Raman et al, 1997). We first hyperpolarized the neuron to −90 mV to shift inactivated Na⁺ channels into a closed state, and then briefly depolarized to 30 mV for 15 ms to open Na⁺ channels, before performing a repolarizing step to −30 mV for 200 ms to generate the resurgent Na⁺ currents (Fig 3D). Significantly, we detected a resurgent Na⁺ current with distinct components (see labelled example trace in Fig 3D). We observed a fast tail current (R_Tail) that occurred ∼1 ms after cells were repolarized to −30 mV (rise time [0–100%] = 0.92 ± 0.39 ms; n = 156 cells across all developmental stages). We termed this current a tail resurgent Na⁺ current as the amplitude of the tail current had a near linear voltage dependency as the voltage of the resurgent step was changed from −80 to 20 mV (Fig S2A and B), and was attenuated as the time of the depolarizing step increased (Fig S2C and D), which is consistent with characteristics of tail Na⁺ currents (Raman et al, 1997). We also observed a resurgent Na⁺ current (R) that peaked at ∼7.5 ms (rise time [0–100%] = 7.50 ± 9.64 ms) before decaying to a persistent state with a half-life of ∼24 ms (decay time [100–50%] = 23.99 ± 13.64 ms). Significantly, the resurgent Na⁺ current peaked at −60 mV when the repolarization voltage was stepped from −90 to +30 mV (Fig S2A and B), decreased as the time of the depolarizing step increased (Fig S2C and D), and increased as the voltage of the depolarizing step was increased from 0 to +30 mV (Fig S2E and F), consistent with what has previously been described for resurgent Na⁺ currents (Patel et al, 2016). Lastly, we identified a persistent component of the resurgent Na⁺ current (R_Persistent) that showed voltage kinetics similar to the persistent Na⁺ current (Fig S2A and B). The resurgent Na⁺ current most closely resembled resurgent currents initially described in rodent Purkinje neurons (Raman et al, 1997; Raman & Bean, 2001), and it is the first description of a resurgent Na⁺ current in Xenopus tectal neurons.

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Figure S2. Characterizing the voltage and time dependency of the resurgent Na⁺ current.

Resurgent Na⁺ currents result from a distinctive gating mechanism that traps Na⁺ channels in a blocked open-state, which is then relieved at depolarized potentials to generate a subthreshold depolarizing current. As a result, resurgent currents display distinct voltage dependency to repolarizing voltage steps, and are affected by the probability of Na⁺ channels entering the open state (voltage of the depolarizing step) or the inactivated/closed state (time of the depolarizing step). (A, B) The effect of altering the voltage of the repolarizing resurgent step on the identified components of the resurgent current. (A) Example voltage clamp recording showing the response of a single cell when the voltage of the repolarization step was increased from −90 to 20 mV. (B) Plot shows the average voltage dependency of each component of the resurgent current in tectal neurons. (C, D) The effect of altering the time of the depolarizing step on each component of the resurgent current. (C) Example voltage-clamp recording illustrating the response of a single cell when the depolarizing step was increased from 5–35 ms. (D) Plot shows the average voltage response for each component of the resurgent current as the time of the depolarizing step increases. (E, F) The effect of altering the voltage of the resurgent step on each component of the resurgent current. (E) Example voltage-clamp recording showing the response of a single cell when the voltage of the depolarizing step was changed from −20 to 30 mV. (F) Plot illustrates how altering the voltage dependency of the resurgent currents. Plots shown in (B, D, F) are mean ± SD; n = 27 tectal neurons. Resurgent currents are plotted on the left y-axis. Total resurgent current, calculated by measuring the area under the curve or charge (pA*s) over the 200 ms resurgent step, is plotted on the right y-axis.

If the regulation of a resurgent Na⁺ current is a mechanism used by tectal neurons to regulate intrinsic excitability, then we would predict that the resurgent current would be regulated with homeostatic changes in intrinsic excitability across development and with exposure to visual stimulation. Consistent with what we had observed for the fast and persistent Na⁺ currents, we found that the resurgent Na⁺ current transiently peaked at developmental stage 46 and was increased in response to EVS (Fig 3E and F). Crucially, this regulation of resurgent Na⁺ current with changes in excitability was not observed for the tail or persistent resurgent Na⁺ current (Fig S3A–C), providing evidence for our identification of the resurgent Na⁺ current. These data show that changes in the resurgent Na⁺ current correlate with changes in intrinsic excitability, suggesting that tectal neurons regulate fast, persistent, and resurgent Na⁺ currents together to control homeostatic changes in intrinsic excitability.

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Figure S3. Quantification of changes in identified components of the resurgent Na+ current in Xenopus tectal neurons with changes in intrinsic excitability.

Quantification of changes in identified components of the resurgent current across development and in response to 4 h of enhanced visual stimulation (EVS). (A) Peak amplitude of the tail resurgent Na⁺ current (R_Tail) ([in pA] Stage 42: 75.4 ± 23.7; Stage 46: 84.5 ± 36.7; Stage 49: 73.1 ± 33.5; Stage 49 + EVS: 87.4 ± 40.9). (B) Peak amplitude of the persistent resurgent Na⁺ current (R_Persistent) ([in pA] Stage 42: 22.3 ± 10.8; Stage 46: 34.2 ± 16.7; Stage 49: 27.2 ± 19.7; Stage 49 + EVS: 39.5 ± 27.2). (C) The total resurgent Na⁺ current (R_Total) ([in pA*s] Stage 42: 5.99 ± 2.55; Stage 46: 9.02 ± 3.54; Stage 49: 6.82 ± 4.07; Stage 49 + EVS: 9.91 ± 5.31). Groups were compared using a Welch’s ANOVA test with Dunnett T3 test for multiple comparisons. n = 24–87 cells.

Persistent and resurgent Na⁺ currents, but not the fast Na⁺ current, are insensitive to tetrodotoxin (TTX)

We next performed a series of experiments utilizing either ion substitution or specific channel inhibitors to further characterize the ionic permeability and channel types that contribute to the fast, persistent, and resurgent Na⁺ currents observed in tectal neurons.

To confirm that persistent and resurgent Na⁺ currents identified in this study were indeed the result of Na⁺ influx, we performed an ion substitution experiment by recording voltage-clamp currents when extracellular Na⁺ was replaced with NMDG. NMDG is an impermeant organic monovalent cation that abolishes inward Na⁺ currents (Blair & Bean, 2002) (Fig 4A–D). When we recorded from tectal neurons in NMDG external using a Tris-based internal solution (compare black and grey traces in Fig 4A–C), we found that blocking Na⁺ influx abolished the fast, persistent and resurgent Na⁺ currents (Fig 4D–F and Table S2). Abolishing Na⁺ currents revealed a small presumptive Ca²⁺ current as previously described (Hamodi & Pratt, 2014). To show that the effect of NMDG external was specific to Na⁺ currents, we also recorded voltage-clamp currents using a K⁺-based internal (Fig S4), which confirmed that NMDG specifically abolishes Na⁺ currents. Furthermore, to exclude the possibility that the observed persistent and resurgent currents are the result of inward flux of K⁺ ions, we measured Na⁺ currents when K⁺ channels were blocked by performing recordings with a TEA-based external saline. Not unsurprisingly, we observed no effect of blocking K⁺ influx on Na⁺ currents (Table S2). These data suggest that the persistent and resurgent Na⁺ currents identified in this study are carried by voltage-gated Na⁺ channels.

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Figure S4. Removal of external Na⁺ specifically inhibits Na⁺ influx without affecting K⁺ currents.

(A) Example voltage-clamp recordings with a K⁺-based internal saline in control and Na⁺ deficient (NMDG) external saline solutions. Magnifications of the initial 30 ms of each trace illustrate how Na⁺ and K⁺ currents are temporally isolated in mixed current recordings. (B) I–V plot illustrates the specific effect of removing external Na⁺ ions on the Na⁺ currents (circles), sparing the K⁺ current (squares). (C, D) Quantification of the effect of NMDG external on Na⁺ currents and K⁺ currents (****P < 0.0001, n = 7–9 cells).

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Figure 4. The persistent and resurgent Na⁺ currents, but not the fast Na⁺ current, are insensitive to tetrodotoxin (TTX).

(A, B, C) Example voltage-clamp recordings from tectal neurons with a Tris-based internal solution to isolate Na⁺ currents from stage 49 tectal neurons in control conditions (black), in zero Na⁺ external solution (NMDG) to block all inward Na⁺ currents (grey), in 1 µM TTX to block TTX-sensitive Na⁺ currents (magenta), or in 100 nM Cd²⁺ to block Ca²⁺ currents (cyan). (A) Example recordings show fast and persistent from a single depolarizing step to −15 mV for each experimental group. (A, B) Magnification of the initial 15 ms of the recordings shown in (A) to a series of voltage steps (−65 to +25 mV) to highlight the effect of each treatment on the fast Na⁺ current. Note that TTX attenuates fast but not persistent Na⁺ currents, whereas blocking all Na⁺ influx by replacing external Na⁺ with NMDG attenuates both fast and persistent currents to reveal a small, presumptive Ca²⁺ current. (C) Example recordings illustrating the effect of each condition on the resurgent Na⁺ current. (D, E, F) Quantification of peak amplitudes for the (D) fast, (E) persistent, and (F) resurgent Na⁺ currents for each experimental group. Groups were compared using a Welch’s ANOVA test with Dunnett T3 test for multiple comparisons. n values for (D, E, F) were 87 stage 49 controls, 15 NMDG, 27 TTX, 11 TTX + Cd²⁺, and 12 Cd²⁺. Values and comparisons are shown in Table S2.

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Source Data for Figure 4[LSA-2023-02232_SdataF4.xlsx]

TTX-resistant Na⁺ currents have been observed in neurons of anuran species (Campbell, 1992a, 1992b; Kobayashi et al, 1993, 1996). Although the fast Na⁺ current in Xenopus tectal neurons is sensitive to the Na⁺ channel blocker TTX (Aizenman et al, 2003), it remained to be determined whether persistent and resurgent Na⁺ currents are sensitive to TTX. Because mammalian Na_v1.1, Na_v1.2, and Na_v1.6 channel subtypes are TTX-sensitive, we predicted that TTX would also abolish the persistent and resurgent currents in tectal neurons. However, when we measured Na⁺ currents in the presence of 1 µM TTX (magenta traces in Fig 4A–C), we found that TTX attenuated the fast Na⁺ current as expected; however, surprisingly, there was no significant effect of TTX on the amplitude of persistent or resurgent Na⁺ currents (Fig 4D–F and Table S2). Moreover, the persistent and resurgent Na⁺ currents remained after the concentration of TTX was increased to 30 μM (Table S2), suggesting that the persistent and resurgent Na⁺ currents are largely TTX-insensitive in Xenopus tectal neurons.

As we had observed no effect of TTX on the persistent and resurgent Na⁺ currents, we next tested whether the less-specific voltage-gated Na⁺ channels blocker lidocaine attenuates persistent and resurgent Na⁺ currents (Fig S5A and B). Because lidocaine binds Na⁺ channels in the inactivated state, we measured Na⁺ currents after a 10 s depolarizing step to 0 mV. When we measured Na⁺ currents in the presence of 1 µM lidocaine, we found a near abolishment of fast, persistent, and resurgent Na⁺ currents (Table S2). These data provide further evidence to suggest that persistent and resurgent Na⁺ currents in Xenopus tectal neurons are carried by TTX-resistant voltage-gated Na⁺ channels.

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Figure S5. Effect of lidocaine on Na⁺ currents in tectal neurons.

(A) Example voltage-clamp recordings showing the effect of 1 µM lidocaine on the Na⁺ currents after a 10 s depolarization step to 0 mV, which allows for lidocaine block of open Na⁺ channels. Recordings were made with a Tris-based internal saline in stage 49 control tectal neurons (black) or tectal neurons exposed to lidocaine (blue). (B) Box indicates portion of each trace shown in (B). (B) Magnification of each trace illustrates how lidocaine affects the fast, persistent, and resurgent Na⁺ currents. Quantification is shown in Table S2.

We had observed a small, presumptive Ca²⁺ current in the absence of extracellular Na⁺. Therefore, we examined whether the TTX-insensitive persistent and resurgent Na⁺ currents could be mediated by Ca²⁺ influx via voltage-gated Ca²⁺ channels. To test this, we recorded currents in the nonspecific voltage-gated Ca²⁺ channels blocker Cd²⁺ in the presence or absence of TTX (cyan traces in in Fig 4A–C). Crucially, we observed no effect of blocking voltage-gated Ca²⁺ channels on the amplitude of the fast, persistent or resurgent Na⁺ current (Fig 4D–F and Table S2). These findings show that Ca²⁺ influx contributes little to the persistent and resurgent current, consistent with the idea that the TTX-insensitive component of the persistent and resurgent Na⁺ currents being mediated by Na⁺ influx via voltage-gated Na⁺ channels. Taken together, these experiments confirm that tectal neurons express a TTX-sensitive fast Na⁺ current and extend this finding to demonstrate the presence of a TTX-insensitive component of the persistent and resurgent Na⁺ currents.

Inhibition of Na⁺ channel subtype Na_v1.6 attenuates resurgent Na⁺ currents and decreases the intrinsic excitability of tectal neurons

To understand the mechanisms by which Na⁺ currents are dynamically regulated with changes in the excitability of tectal neurons, we sought to determine the molecular composition of the fast, persistent, and resurgent Na⁺ currents. Our expression analysis had suggested that Na_v1.1 and Na_v1.6 channels may contribute to the regulation of Na⁺ currents and intrinsic excitability, whereas Na_v1.2 channels s did not appear to be regulated with intrinsic excitability. Although Na_v1.6 is TTX-sensitive in mammalian neurons (Lee & Ruben, 2008), it is evolutionarily distinct from Na_v1.1 and Na_v1.2 (Zakon, 2012), and has been shown to mediate persistent and resurgent Na⁺ currents (Raman et al, 1997; Bant & Raman, 2010; Patel et al, 2015). Furthermore, Xenopus and other anuran species have a tyrosine to phenylalanine substitution at the same residue within the P-loop of domain I of Na_v1.6 that has been shown to be critical for TTX binding in the channel pore (Herzog et al, 2003), which may confer Xenopus Na_v1.6 with distinct TTX sensitivity. We therefore hypothesized that Na_v1.6 could be a candidate to mediate TTX-insensitive components of the persistent and resurgent Na⁺ currents in Xenopus tectal neurons.

To examine the contribution of Na_v1.6 channels to the regulation of persistent and resurgent Na⁺ currents and intrinsic excitability in tectal neurons, we performed whole-cell recordings that measured Na⁺ currents and intrinsic excitability in the presence of the specific Na_v1.6 inhibitor MV1312 (Fig 5). MV1312 shows five to sixfold sensitivity for Na_v1.6 over Na_v1.1, and has been shown to rescue seizure behavior in a zebrafish model of Dravet syndrome where a loss-of-function Na_v1.1 function results in overexpression of Na_v1.6 and epileptogenesis (Weuring et al, 2020). MV1312 is predicted to bind Na⁺ channels similar to lidocaine, therefore we performed whole-cell recordings with Tris-based internal solution and acutely washed in 5 µM MV1312 with recordings preceded by a 5-s depolarizing step to 0 mV to open Na⁺ channels (Fig 5A). We found that blocking Na_v1.6 channels caused a significant attenuation of all Na⁺ currents, with the largest effect observed for the resurgent Na⁺ current (Fig 5B–D). These findings suggest that MV1312 blocks Na⁺ channels in the inactivated state with slow kinetics, and provide evidence that Na_v1.6 channels are a major contributor to fast, persistent, and resurgent Na⁺ currents in Xenopus tectal neurons.

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Figure 5. The Na_v1.6 specific inhibitor MV1312 decreases intrinsic excitability by reducing fast, persistent, and resurgent Na⁺ currents in a use-dependent manner.

(A) To measure the effect of Na_v1.6 channel inhibition on Na⁺ currents, we performed whole-cell recordings in the presence or absence of 5 µM of the specific Na_v1.6 channel inhibitor MV1312. We recorded Na⁺ currents before and after a 5 s depolarizing step to 0 mV to promote channel opening and drug binding, and then calculated the fraction of each Na⁺ current that remained. (B, C, D) Quantification of the fraction of initial peak current amplitude for the (B) fast Na⁺ current (Control: 0.82 ± 0.07, n = 8, MV1312: 0.70 ± 0.04, n = 5; P = 0.0038), (C) persistent Na⁺ current (Control: 0.73 ± 0.12, MV1312: 0.58 ± 0.10; P = 0.0439), and (D) the resurgent Na⁺ current (Control: 0.74 ± 0.10, MV1312: 0.38 ± 0.08; P < 0.0001). Groups were compared with an unpaired t test. (E, F) MV1312 decreases intrinsic excitability of tectal neurons by attenuating Na⁺ current amplitude. (E, F, G) The effect of MV1312 on intrinsic excitability was observed by measuring (E) spikes generated by current injection, (F) the maximum number of spikes generated by current injection (Control: 2.88 ± 1.32, n = 17; MV1312: 1.75 ± 0.72, n = 20; P = 0.0021), and (G) the capacity of cells to spike in response to 200 ms cosine current injections at 30 Hz with increasing amplitudes from 40 to 120 pA (response frequency [% of current injections resulting in a spike]. 40 pA: Control 0.93 ± 0.02, n = 16, MV1312 0.72 ± 0.07, n = 19; P = 0.0002; 80 pA: Control 1.00 ± 0.00, n = 16, MV1312 0.97 ± 0.02, n = 19; P = 0.7429; 120 pA: Control 1.00 ± 0.00, n = 16, MV1312 0.99 ± 0.01, n = 19; P = 0.9712. Groups were compared using a two-way ANOVA with a Holm–Sidak test for multiple comparisons).

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Source Data for Figure 5[LSA-2023-02232_SdataF5.xlsx]

If Na_v1.6-mediated Na⁺ currents are important regulators of intrinsic excitability in tectal neurons, then inhibition of Na_v1.6 channels would be predicted to reduce neuronal excitability. To measure the effect of Na_v1.6 channel inhibition on the intrinsic excitability of tectal neurons, we performed whole-cell recordings with a K⁺-based internal solution in the presence of 5 µM MV1312. Because tectal neurons at stage 49 inherently exhibit lower intrinsic excitability, it would be difficult to observe a decrease in excitability. As such, we performed recordings on neurons at developmental stage 47/48, when intrinsic excitability of tectal neurons is higher (Pratt et al, 2008; Ciarleglio et al, 2015). As expected, the intrinsic excitability of control stage 47/48 tectal neurons was increased compared with stage 49 neurons (compare Fig 5E–G with Fig 1B). Crucially, we found that the intrinsic excitability of stage 47/48 tectal neurons exposed to MV1312 was significantly decreased compared with controls (Fig 5E and F), which shows that Na⁺ currents carried by Na_v1.6 channels are important regulators of intrinsic excitability in Xenopus tectal neurons.

Tectal neurons, as the major recipient of sensory inputs in the optic tectum, play a crucial role in integrating multisensory information to produce an appropriate behavioral response (Deeg et al, 2009; Felch et al, 2016; Truszkowski et al, 2017; Busch & Khakhalin, 2019). As such, it is vital that tectal neurons can faithfully generate an action potential in response to graded sensory inputs. Therefore, we tested whether inhibition of Na_v1.6 channels with MV1312 alters the capacity of tectal neurons to repeatedly generate action potentials in response to a cosine-shaped injection of current. For these experiments, cells were exposed to repeated 200 ms cosine current injections at 30 Hz, with an increasing current amplitude from 40 to 120 pA (Fig 5G; top). The response frequency for each cosine current amplitude was calculated by measuring whether a spike was generated to each individual cosine wave. Significantly, whereas control cells could faithfully generate spikes at all current amplitudes, MV1312-exposed tectal neurons showed a lower response frequency at 40 pA (Fig 5G), which is consistent with Na_v1.6-mediated Na⁺ currents being important for the response properties of tectal neurons. Taken together, these data suggest that Na_v1.6 channels contribute to all phases of the Na⁺ currents and are important for determining tectal neuron action potential firing.

Regulation of Na⁺ channel subtype Na_v1.6 mediates homeostatic changes in Na⁺ currents to control the intrinsic excitability of tectal neurons

We observed changes in the levels of expression of Na_v1.1 and Na_v1.6 channels in the optic tectum across tectal circuit development and in response to EVS in a similar manner to the regulation of Na⁺ currents and intrinsic excitability. We also found that inhibition of Na_v1.6 channels reduces intrinsic excitability. We therefore hypothesized that these Na⁺ channel subtypes may play an important role in the homeostatic regulation of intrinsic excitability. Knockout of Na_v1.1, Na_v1.2 or Na_v1.6 channels is lethal, resulting in prenatal death in rodent models (Catterall et al, 2010), whereas heterozygous loss of function of individual Na⁺ channel subtypes causes compensatory changes that often leads to neuronal and circuit hyperexcitability and seizures (Meisler et al, 2021). To determine the contribution of individual Na⁺ channel subtypes to homeostatic changes in Na⁺ currents and intrinsic excitability, we tested whether antisense morpholino RNA technology targeted to individual Na_v channel subtypes could suppress the EVS-mediated up-regulation of Na⁺ currents and intrinsic excitability. To achieve this, we bulk electroporated the tectum of stage 49 tadpoles with lissamine-tagged, translation-blocking antisense morpholino oligonucleotides (MO) specific for Na_v1.1, Na_v1.2 or Na_v1.6 channels, or a control MO. After 24 h, we performed whole-cell recordings from lissamine-positive tectal neurons and measured Na⁺ currents and intrinsic excitability from control tadpoles or tadpoles exposed to 4 h of EVS (Fig 6A).

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Figure 6. Knockdown of Na_v1.6 attenuates network activity-dependent homeostatic increase in Na⁺ currents.

(A) Schematic illustrates how exposure of stage 49 tadpoles to 4 h of enhanced visual stimulation (EVS) decreases excitatory synaptic drive and triggers a compensatory increase in intrinsic excitability via an increase in Na⁺ currents. (B, C, D) Quantification showing the effect of acute knockdown of expression of specific Na_v channel subtypes on the EVS-triggered increases in the amplitude of the fast, persistent, and resurgent Na⁺ currents. Comparisons between experimental groups are presented in Table S3 (n = 16–22 cells). (E, F, G, H) Exposure of stage 49 tadpoles to 4 h of EVS triggers an increase intrinsic excitability, which is attenuated by the suppression of Na_v1.6 channel expression, but not Na_v1.1 channel expression (n = 16–31 cells). (I) K⁺ currents are not regulated with EVS-induced increases in intrinsic excitability. Comparisons between experimental groups are presented in Table S4. Groups were compared using a Welch’s ANOVA test with Dunnett T3 test for multiple comparisons.

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Source Data for Figure 6[LSA-2023-02232_SdataF6.xlsx]

We found that suppressing up-regulation of Na_v1.6 expression prevented the EVS-mediated increase in the fast, persistent, and resurgent Na⁺ currents; whereas suppressing up-regulation of Na_v1.1 expression prevented the EVS-mediated increase in the persistent and resurgent Na⁺ currents, but did not prevent an increase in the fast Na⁺ current (Fig 6B–D and Table S3). In addition, we observed that knockdown of Na_v1.6 triggers a compensatory increase in the fast Na⁺ current compared with control MO (Fig 6B and Table S3), providing further evidence for a crucial role for Na_v1.6 channels in regulating excitability of tectal neurons. In contrast, we observed that up-regulation of Na_v1.2 expression was not required for EVS-mediated increase in the fast, persistent or resurgent Na⁺ currents (Fig 6B–D and Table S3). Taken together these data provide evidence that increased expression of Na_v1.6, and to a lesser extent Na_v1.1, is the molecular mechanism leading to for EVS-mediated increases in Na⁺ currents.

Our data indicated that the up-regulation of Na_v1.6 expression is required for EVS-mediated increases in Na⁺ currents, and that the specific Na_v1.6 channel blocker MV1312 decreases intrinsic excitability, suggesting that Na_v1.6 is a key regulator of Na⁺ currents that control homeostatic changes in intrinsic excitability. However, we had also observed that suppression of Na_v1.1 can reduce EVS-mediated increases in persistent and resurgent Na⁺ currents. Therefore, we next tested whether suppressing Na_v1.6 or Na_v1.1 expression attenuates the EVS-mediated homeostatic increase in intrinsic excitability. We found that suppressing up-regulation of Na_v1.6 expression prevented the EVS-mediated increases in intrinsic excitability, as shown by its effects on the input–output curves and the maximum number of spikes generated by current step injection compared with control MO tadpoles (Fig 6E, F, and H and Table S4). In contrast, we found that inhibiting the up-regulation of Na_v1.1 expression has no effect on the EVS-mediated increases in intrinsic excitability (Fig 6E, G, and H and Table S4). As expected, there was no effect of EVS or Na_v1.6 MO on the peak amplitude of K⁺ currents (Fig 6I and Table S4), providing further evidence that regulation of Na⁺ currents is key to the control of intrinsic excitability in tectal neurons. Taken together, these data indicate that although EVS-induced increases in Na⁺ currents are mediated by both Na_v1.6 and Na_v1.1, but that it is Na_v1.6 up-regulation that is critical for mediating the increase in excitability as measured by spike output.

Perturbing expression of Na_v1.1 and Na_v1.6 channels during tectal circuit development causes deficits in sensorimotor behaviors

What are the functional consequences of Na⁺ channel subunit-mediated regulation of intrinsic excitability during development? The optic tectum, the primary visual area in the tadpole brain, undergoes activity-dependent refinement during development, which is necessary for the accurate performance of visually guided behaviors (Dong & Aizenman, 2012; Khakhalin et al, 2014; Shen et al, 2014; James et al, 2015; Hamodi et al, 2016). Activity-dependent refinement of tectal circuitry from developmental stage 46 leads to better visual acuity as receptive field size decreases, and the temporal window for multisensory integration becomes narrower, whereas interventions that alter this activity-dependent process cause perturbed performance in tests of visual acuity, multisensory integration, and schooling behaviors (Schwartz et al, 2011; James et al, 2015; Truszkowski et al, 2016). One hypothesis is that the increase in excitability mediated by elevated Na_v1.1 and Na_v1.6 levels during developmental stage 46 is important for creating a permissive environment for activity-dependent plasticity required for proper tectal circuit development. Thus, we asked whether perturbing expression of Na⁺ channel genes Na_v1.1 and Na_v1.6 during tectal circuit development causes behavioral deficits in tasks that require sensorimotor transformations such as visual acuity, multisensory integration, and schooling.

For these experiments, tadpoles at developmental stages 44–45 were co-electroporated with Na_v1.1 MO and Na_v1.6 MO, or with a control MO, to suppress Na⁺ channel expression during a critical window of circuit development, with the behavioral tasks performed at stage 49. Importantly, electroporation of tadpoles with Na_v channel-specific MOs or with Na_v1.1 MO + Na_v1.6 MO at stage 44–45 had no effect on the amplitude of the fast or persistent Na⁺ currents at stage 49 (Table S5), which suggests that the effect of the channel expression knockdown on Na⁺ current amplitude does not persist into the period of behavioral assessment. This is most likely because of the tectal neurons having compensated for the effect of the MOs on Na⁺ channel expression (Van Wart & Matthews, 2006; Vega et al, 2008).

Visual acuity behavior correlates with the capacity of tectal neurons to tune visual spatial frequency sensitivity (Schwartz et al, 2011), providing a measure of tectal circuitry development. When presented with counterphasing gratings tadpoles change their swimming speed proportional to the spatial frequency of the gratings (Schwartz et al, 2011). We therefore examined whether perturbing expression of Na⁺ channel genes Na_v1.1 and Na_v1.6 during tectal circuit development affects visual acuity behavior. Visual acuity responses were measured by calculating the change in velocity of tadpoles in response to the onset of a series of counterphasing sine wave gratings of different spatial frequencies presented (3, 4.5, 9, and 18 cycles/cm; Fig 7A). To analyze responses, we calculated a Z-score for individual tadpole to each spatial frequency, which describes how many standard deviations the mean response to each stimulus was from the mean response to no stimulus. To determine if tadpoles had responded to the visual stimuli, we tested whether the mean Z-score for control MO and Na_v MO tadpoles to each spatial frequency was different from the no stimulus control. Control MO tadpoles showed responsiveness to gratings with low spatial frequency (Fig 6B), consistent with what has previously described (Schwartz et al, 2011). In contrast, Na_v MO tadpoles showed decreased responsiveness to low spatial frequencies (Fig 7B), indicating a decreased visual acuity response. Importantly, this decreased the response in Na_v MO tadpoles was not the result of decreased motility, as baseline motility was unchanged between control and Na_v MO tadpoles (Fig 7C). These data suggest that regulation of Na_v1.1 and Na_v1.6 expression levels during a critical period of tectal circuit development is necessary for the normal development of visual acuity response properties.

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Figure 7. Tadpoles in which expression of Na_v1.1 and 1.6 was perturbed during tectal circuit development show impairments in visual acuity, multisensory integration, and schooling behaviors.

Effect of knocking down of Na_v1.1 and Na_v1.6 expression at stages 44–46 on tadpole behavior at stage 49 (comparisons showing that knockdown of Na_v1.1 and Na_v1.6 expression treatment had no effect on Na⁺ currents at stage 49 are presented in Table S5). (A, B, C) Effect of developmental knockdown of Na_v expression on visual acuity behavior. (A) Control morpholino (Ctrl MO) or Na_v1.1/Na_v1.6 morpholino (Na_v MO) tadpoles were exposed to gratings counterphasing at 4 Hz over a range of spatial frequencies (3, 4.5, 9, and 18 cycles/cm). (B) Knockdown of Na_v1.1 and Na_v1.6 during the critical period of tectal development impaired responses to low spatial frequencies. Ctrl MO. 3 cycles/cm: 7.0 ± 3.9 (P < 0.0001), 4.5 cycles/cm: 3.5 ± 2.7 (P < 0.0001), 9 cycles/cm: 1.3 ± 1.8 (P = 0.1238), 18 cycles/cm: 0.1 ± 1.4 (P = 0.8330). Na_v MO. 3 cycles/cm: 2.6 ± 3.4 (P = 0.0107), 4.5 cycles/cm: 1.0 ± 2.0 (P = 0.9059), 9 cycles/cm: 0.5 ± 1.6 (P = 0.9056), 18 cycles/cm: 0.7 ± 1.8 (P = 0.5767). N = 7 experiments of 3–4 animals. Groups were compared using a two-way ANOVA test with Holm–Sidak test for multiple comparisons. (C) This effect of Na_v knockdown on visual acuity was not caused by a change in motility (Ctrl MO: 1.34, IQR = 1.00–1.67; Na_v MO: 1.44, IQR = 0.87–1.72; P = 0.8886, n = 25–27 animals). Groups compared using a Mann–Whiney U test. (D, E, F, G, H) Effect of developmental knockdown of Na_v expression on multisensory integration behavior. (D) Experimental paradigm illustrating the presentation of visual and mechanosensory stimuli, or multisensory stimuli with interstimulus intervals ranging from 0–500 ms. (E) Mean normalized change in velocity (cm/s) in response to unisensory visual or mechanosensory stimuli, or multisensory stimuli, for Control MO and Na_v MO tadpoles. (F) Multisensory (MS) indexes calculated for individual tadpoles at 0, 250, and 500 ms. Values for individual tadpoles are connected by a line. (G) Quantification of MS index for interstimulus intervals of 0 ms (Ctrl MO: 1.1 ± 0.5, Na_v MO: 0.4 ± 0.5, P < 0.0001), 250 ms (Ctrl MO: −0.4 ± 0.2, Na_v MO: −0.2 ± 0.6, P = 0.0713) and 500 ms (Ctrl MO: 0.6 ± 0.5, Na_v MO: 0.3 ± 0.4, P = 0.0438). Groups were compared using Welch’s t test. (H) Histogram of preferred interstimulus intervals for each tadpole. N = 8 experiments of 3–4 animals. (I, J, K) Effect of developmental knockdown of Na_v expression on schooling behaviour. (I) Schematic illustrates aggregated schooling behavior observed in control MO and Na_v MO tadpoles, with Na_v MO tadpoles observed to be less likely to be swimming in the same direction, without a change in inter-tadpole distance. (J, K) These observations are quantified by observing (J) inter-tadpole distance (Ctrl MO: 2.4 cm, IQR = 1.4–3.6 cm; Na_v MO: 2.4 cm, IQR = 1.3–3.6 cm; P = 0.4088), and (K) inter-tadpole angles (Ctrl MO: 45.7°, IQR = 19.7–93.9°; Na_v MO: 65.6°, IQR = 29.2–119.1°; P < 0.0001). N = 6 experiments of 20 animals per experimental group. Groups were compared using a Kolmogorov–Smirnov test.

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Source Data for Figure 7[LSA-2023-02232_SdataF7.xlsx]

Multisensory integration (MSI), one of the primary functions of the tectum, is a highly conserved property of both neuronal output and behavior whereby the response to a stimulus of a single sensory modality is modulated by the coincident presentation of a stimulus from a different sensory modality. MSI depends on the strength of the neuronal response to each individual unimodal stimulus, the overlap between spatial receptive fields for the two sensory modalities, and the time window between presentations of the cross-modal pair (Wallace et al, 1998, 2006). As tectal neurons mature, they become more narrowly tuned to a more diverse range of interstimulus intervals, which occurs congruent with activity-dependent strengthening and refinement of synaptic connections (Felch et al, 2016). Because of its complex nature, MSI is a robust readout of connectivity deficits within the optic tectum. We therefore asked whether perturbing expression of Na⁺ channel genes Na_v1.1 and Na_v1.6 during tectal circuit development affects MSI. Subthreshold visual or mechanosensory stimuli, or multisensory stimuli with interstimulus intervals of 0, 250, and 500 ms were presented to each tadpole (Fig 7D). Responses were determined by measuring normalized change in velocity of tadpoles to the stimulus onset (Fig 7E), from which we then calculated the multisensory (MS) index (Fig 7F). We observed that control tadpoles robustly respond to multisensory stimuli, with tadpoles increasing swimming to interstimulus intervals of 0 or 500 ms, and slowing down to 250 ms interstimulus intervals (Fig 7G). When we examined the preferred interstimulus intervals (ISI) of control tadpoles, we observed that tadpoles most strongly respond to multisensory stimuli with an ISI of 0 ms, with a few tadpoles preferring an ISI of 500 ms (Fig 7H). In contrast, Na_v MO tadpoles exhibited a less robust response to multisensory stimuli and showed less temporal preference. Na_v MO tadpoles exhibited a decreased MS index for both 0 and 500 ms (Fig 7F and G), with a shift in the preferred ISI that reflected how Na_v MO tadpoles broadly responded to a wider range of ISI (Fig 7H), indicating a maturation of temporal tuning of multisensory responses.

Tadpoles perform a social aggregation behavior known as schooling, where tadpoles in close proximity engage in a coordinated unidirectional group swimming, which requires the integration of sensory cues (Wassersug & Hessler, 1971; Katz et al, 1981). Schooling tadpoles display short inter-tadpole distances and inter-tadpole angles less than 45° with neighboring tadpoles (depicted in Fig 7I), which is altered when tectal circuitry development is perturbed (James et al, 2015; Truszkowski et al, 2016). Because we had observed that perturbing Na⁺ channel gene expression during tectal circuit maturation causes defects in visual acuity and multisensory integration behaviors, we predicted that perturbing expression of Na_v1.1 and Na_v1.6 during tectal circuit development would affect schooling behavior. Consistent with previous reports, the inter-tadpole distance of control tadpoles was ∼2–3 cm, with control tadpoles most often found to be swimming in the same direction as their neighbors (Fig 7J and K). When we measured schooling behavior in Na_v MO tadpoles, we observed that the inter-tadpole angles of Na_v MO tadpoles was significantly altered, with fewer tadpoles swimming in the same direction (Fig 7K), but with no effect on the inter-tadpole distance (Fig 7J and illustrated in Fig 7I). These data show how perturbing the regulation of intrinsic excitability by impairing regulation of Na⁺ channel gene expression during circuit development alters circuit function and causes impaired schooling behavior, most likely by perturbing multisensory integration in the tectum.

All these behavior tasks require proper activity-dependent refinement of tectal circuits. We observed that tadpoles in which the expression of Na_v1.1 and Na_v1.6 was perturbed during tectal circuit development shows reduced visual acuity responsiveness, broader multisensory tuning, and abnormal schooling behavior. These data illustrate the vital importance of the correct regulation of neuronal excitability by Na⁺ channel gene expression for normal tectal circuit development.