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Developmental, Molecular, and Genetic Dissection of I_Na In Vivo in Embryonic Zebrafish Sensory Neurons

Department of Physiology and Biophysics, University of Colorado Health Sciences Center at Fitzsimons, Aurora, Colorado

Submitted 12 October 2004; accepted in final form 25 January 2005

	ABSTRACT

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The presence of multiple Na_v1 isotypes within a neuron and the lack of specific blockers hamper identification of the in vivo roles of sodium current (I_Na) components, especially during embryonic stages. To identify the functional properties of I_Na components in vivo in developing neurons, we took a molecular genetic approach. Embryonic zebrafish Rohon–Beard (RB) mechanosensory neurons express two different sodium channel isotypes: Na_v1.1 and Na_v1.6. To examine the properties of Na_v1.1- and Na_v1.6-encoded currents in RB cells at different developmental stages, we eliminated the contribution of Na_v1.6 and Na_v1.1 channels, respectively, using an antisense morpholino (MO) approach. MOs were injected into one-cell stage embryos, and RB sodium currents were recorded using patch-clamp techniques in both conventional whole cell mode as well from nucleated patches. Only a subset of RB cells appeared to be affected by the Na_v1.1MO. Overall, the effect of the Na_v1.1MO was a small 25% average reduction in current amplitude. Further, Na_v1.1MO effects were most pronounced in RB cells of younger embryos. In contrast, the effects of the Na_v1.6 MO were observed in all cells and increased as development proceeded. These results indicated that developmental upregulation of RB I_Na entailed an increase in the number of functional Na_v1.6 channels. In addition, analysis of voltage-dependent steady-state activation and inactivation parameters revealed that specific functional properties of channels were also developmentally regulated. Finally, analysis of macho mutants indicated that developmental upregulation of I_Na was absent in RB cells. These results indicate that MOs are a useful tool for the molecular dissection and analysis of ion channel function in vivo.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION GRANTS ACKNOWLEDGMENTS REFERENCES

 
During development, neurons display dynamically regulated changes in the number as well as the properties of ion channels (for reviews see Spitzer and Ribera 1998; Spitzer et al. 2002). Regulation of functional ion channel expression could occur at multiple steps, including transcriptional, posttranscriptional, translational, and posttranslational levels. Analysis of developmental regulation of electrical excitability needs to consider both molecular and functional characteristics of specific currents. Further, because neurons are electrically excitable before synapse formation, developmental analyses need to consider prenatal periods. Finally, the diversity of electrophysiological phenotypes displayed by neurons, either during development or in the adult, indicates that analysis of developmental mechanisms requires cell-type–specific study.

A classic system for multilevel analysis of neuronal excitability at early stages is the developing spinal cord of the Xenopus embryo. For this system, several lines of evidence implicate transcriptional mechanisms in developmentally regulated changes in excitability. For example, the duration of the action potential shortens as neurons mature (Baccaglini and Spitzer 1977; Spitzer and Lamborghini 1976). This developmentally regulated change in excitability requires upregulation of voltage-gated potassium current (I_Kv) that in turn requires new transcription (Ribera and Spitzer 1989). Even in this system, however, there is evidence for posttranslational control of I_Kv (Blaine et al. 2004).

Several different types of neurons increase sodium current density during early developmental stages (Alessandri-Haber et al. 1999; Baines and Bate 1998; Dourado and Dryer 1992; Gao and Ziskind-Conhaim 1998; Gottmann et al. 1991; Huguenard et al. 1988; MacDermott and Westbrook 1986; McCobb et al. 1990; Nerbonne and Gurney 1989; O'Dowd et al. 1988; Ribera and Nüsslein-Volhard 1998; Schmid and Guenther 1998; Skaliora et al. 1993). Despite the critical role that sodium current (I_Na) plays in generation of rapid signaling in the nervous system, little is known about mechanisms that regulate expression and function of this current during early stages of neuronal development. Some studies support the notion that increases in the numbers of early expressed sodium channels underlie the dramatic upregulation that is frequently observed in embryonic neurons (Dourado and Dryer 1992; Gao and Ziskind-Conhaim 1998; MacDermott and Westbrook 1986; Nerbonne and Gurney 1989). However, other studies indicate that changes in either time- or voltage-dependent properties of sodium channels occur (Huguenard et al. 1988; Schmid and Guenther 1998; Skaliora et al. 1993). At a molecular level, the latter findings raise the possibility that different channel classes appear and/or that channels undergo posttranslational modifications at later developmental stages.

Electrophysiological analysis of the zebrafish macho (mao) mutant indicates that several different classes of sensory neurons fail to undergo developmental upregulation of sodium current. Retinal ganglion cells have much reduced sodium current densities and consequently fail to show an activity-dependent pruning of connections in the tectum (Gneugge et al. 2001). Rohon–Beard (RB) mechanosensory spinal neurons fail to upregulate sodium current density and consequently do not fire overshooting action potentials that are required for generation of the behavioral response to touch (Ribera and Nüsslein-Volhard 1998). Recently, the identities of sodium channel -subunit (Na_v1) genes expressed in zebrafish RB cells have been elucidated (Novak and Ribera 2004). This molecular information combined with the mao mutant provide powerful tools for dissection of mechanisms regulating sodium current maturation in RB cells.

Our studies reveal that 2 different sodium -subunit genes contribute to RB I_Na. One of these, Na_v1.6, underlies current in all RB cells. In contrast, the other, Na_v1.1, contributes to current in only a subset of cells. Further, the consequences of elimination of each subunit vary during development. Elimination of Na_v1.1 has greater effects at earlier stages, whereas knockdown of Na_v1.6 eliminates more current in neurons of older embryos. Elimination of each subunit also selectively allows in vivo examination of developmental regulation of specific I_Na components. These latter studies indicate that both changes in the number of channels as well as their properties occur during differentiation of zebrafish RB cells.

	METHODS

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Animals

Three different wild-type strains were used: Tü, WIK, and PS (the latter strain was from a local pet store). Similar results were obtained with all strains. Adult fish were housed at 28°C in the fish facility of the UCHSC Center for Animal Laboratory Care. Embryos were obtained by pairwise crossing of adults. The mao line was kindly provided by Dr. Hans-Georg Frohnhöfer of the Tübingen Stockcenter. The ethynitrosourea-induced mutation was created in the Tü strain (Granato et al. 1996; Haffter et al. 1996). Macho (mao) mutant zebrafish embryos were obtained from matings of identified heterozygous carriers. Typically, a subset of embryos in a clutch was sorted at the earliest time the phenotype appeared [about 30 h postfertilization (hpf)]. The identification of an embryo as mutant or sibling was reconfirmed just before dissection and recording. Staging was done according to the criteria of external morphology as described by Kimmel et al. (1995).

Morpholino design and microinjection

Antisense oligonucleotide morpholinos (MOs) for sodium channels Na_v1.6 (MO1.6) and Na_v1.1 (MO1.1) were designed and synthesized by GeneTools (Corvalis, OR). The MO1.6 and MO1.1 sequences were 5'-gggTgCAgCCATgTTTTCATCCTgC-3' and 5'-CgCCATT-TTCTCATCCTgAAgCT-3', respectively. Similar results were obtained with MOs that targeted sequences that were upstream or downstream of those above (MO1.6: 5'-gCAAgAAggggTgCAgCCATgTTTT-3'; MO1.1: 5'-TCAgAgTgATCCgCTACAC-3'). For each Na_v1 MO, control MOs were synthesized by introducing mismatches at 5 positions. Stock MO solutions were prepared by resuspending the oligonucleotide in sterile filtered water (5 mM), and aliquots were stored at –80°C. Working MO solutions (1.00–3.30 mg/ml) were prepared by dilution with 1 x Danieau solution [58 mM NaCl, 0.7 mM KCl, 0.4 mM MgSO₄, 0.6 mM Ca(NO₃)₂, 5.0 mM HEPES, pH 7.6] containing 1% Fast Green or 300 mM rhodamine-conjugated dextran (10,000 MW; Molecular Probes, Eugene OR). MOs were microinjected using a gas-driven PLI-100 injection apparatus (Medical Systems, Greenvale, NY) into the yolk sac of one-cell–stage wild-type zebrafish embryos (Nasevicius and Ekker 2000). At 15 min after injection, the embryos were examined and those that had Fast Green or rhodamine–dextran within animal cells were transferred to a petri dish containing embryo medium (130 mM NaCl, 0.5 mM KCl, 0.02 mM Na₂HPO₄, 0.04 mM KH₂PO₄, 1.3 mM CaCl₂, 1.0 mM MgSO₄, 0.4 mM NaH₂CO₃) and then raised at 28°C until the desired developmental stage. For all experiments, embryos were manually dechorionated at 16–24 hpf. Embryos that were injected with MO1.6 or MO1.1 are referred to as 1.6 or 1.1 morphants, respectively.

Semi-intact preparations of spinal cord from zebrafish embryos

Semi-intact preparations of zebrafish embryos were prepared as described previously (Ribera and Nüsslein-Volhard 1998). Briefly, in the presence of Ringer solution (145 mM NaCl, 3 mM KCl, 1.8 mM CaCl₂, 10 mM HEPES, pH 7.2) containing 0.02% tricaine, embryos were mounted on glass coverslips using Vetbond tissue adhesive (3M Animal Care Products, St Paul, MN). Once mounted, embryos were killed by transection at the level of the hindbrain. The skin was removed using dissection needles. Tricaine was then removed by washing the preparation with 40 ml of recording solution over the course of 15 min. Preparations were viewed with differential interference contrast optics on an Axioskop FS microscope (Zeiss) at a magnification of 640x.

Conventional whole cell and nucleated-patch recordings

Current- and voltage-clamp recordings were obtained using conventional whole cell and nucleated-patch configurations. Experiments were performed using an Axopatch 200B patch-clamp amplifier (Axon Instruments, Foster City, CA) in conjunction with a Digidata 1322A (Axon Instruments) analog-to-digital (A/D) interface. The pCLAMP8 software package (Axon Instruments) was used for data acquisition and analysis. Recordings were conducted at room temperature (20–22°C). Unpolished electrodes with tip resistances ranging between 2.5 and 3.5 M were fabricated from borosilicate glass using a P-97 Flaming–Brown micropipette puller (Sutter Instruments, Novato, CA). Cell capacitance and series resistance were routinely compensated by 75–80% with a lag of 10 µs using the electronic features of the amplifier. Currents were filtered at 5 kHz and digitized at 25 kHz. Passive leak and capacitative transients were subtracted on-line using a P/8 protocol (Armstrong and Bezanilla 1977).

After establishment of the whole cell configuration, nucleated patches were formed (Sather et al. 1992). Briefly, mild suction was applied to the recording pipette. Then, the pipette was slowly raised, thereby pulling the Rohon–Beard soma out of the neural tube. Seal resistance was closely monitored as the pipette was raised. Electronic compensation of series resistance and cell capacitance were adjusted as necessary. Typically, after achieving the nucleated-patch configuration, seal resistances and cell capacitances decreased by approximately 5–20% versus the whole cell configuration. Nucleated-patch recordings were continued when the following criteria were met on-line: 1) input resistances >1 G and 2) monoexponential decay of the whole cell capacitance transient.

For recording of voltage-gated sodium currents (I_Na), the bath solution consisted of 127 mM NaCl, 20 mM TEA-Cl, 3 mM KCl, 10 mM CoCl₂, and 10 mM HEPES, pH 7.2. [To determine the extent to which series resistance errors might be affecting currents recorded from RBs with large sodium current (e.g., those from 48 hpf wild-type embryos as in Fig. 2), some recordings were done with reduced extracellular sodium concentration of 30 and 80 mM using N-methyl-glucamine to maintain osmolarity.] The pipette solution contained: 125 mM CsCl, 12 mM NaCl, 10 mM EGTA, and 10 mM HEPES, pH 7.2. Inward sodium currents were evoked from a holding potential of –80 mV by 160-ms depolarizing voltage steps to potentials ranging between –60 and +60 mV in 5- to 10-mV increments.

View larger version (26K): [in this window] [in a new window]   FIG. 2. Amplitude of the sodium current (INa) of RB cells increased during development. A: recordings of INa were obtained from nucleated patches made from RB neurons at 3 different developmental stages: 16–18 (16; top), 24–26 (24; middle), and 48–50 (48; bottom) hours postfertilization (hpf). Sodium currents were evoked by a series of voltage steps to potentials ranging between –60 and +40 mV in 10-mV increments from a holding potential of –80 mV. For illustrative purposes current traces in 20-mV increments are shown. Voltage command protocol used is shown at the bottom of the panel. B: peak current amplitude increased 6-fold between 16 and 48 hpf (***P < 0.001 vs. 16 hpf; n ranged between 9 and 14 for each developmental stage). C: current–voltage (I–V) relationship revealed that peak current was elicited at more negative voltages when recording from RB cells of 24 vs. 16 hpf embryos [–10.0 ± 0.01 mV (24 hpf) vs. –0.01 ± 0.00 (16 hpf); P < 0.001]. Difference in the voltage errors attributed to series resistance (about 2 mV) did not fully account for the observed 10-mV shift in peak current.

For recording of voltage-gated potassium current I_Kv, the bath solution contained 80 mM NaCl, 3 mM KCl, 5 mM MgCl₂, 10 mM CoCl₂, 5 mM HEPES, 1 µM tetrodotoxin (TTX; Calbiochem, La Jolla, CA), pH 7.4. The pipette solution consisted of (in mM): 135 mM KCl, 10 mM EGTA, 10 mM HEPES, pH 7.4. Neurons were held at –80 mV and stepped to potentials ranging between –60 and +40 mV in 10-mV increments for 60 ms. I_Kv amplitudes were measured by averaging values during a 10-ms interval at the end of the depolarizing pulse.

Analysis of data

Statistical analysis was performed using JMP v5.0 software (SAS Institute). Results are presented as means ± SE. Statistical comparisons were done using either the Student's t-test or, for comparisons of multiple groups, ANOVA. Statistical significance was identified by values of P 0.05.

	RESULTS

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION GRANTS ACKNOWLEDGMENTS REFERENCES

 
In this study, we used genetic, molecular, biophysical, and developmental criteria to identify I_Na components in zebrafish RB cells at different developmental stages. A genetic approach was allowed by the mao mutant, in which RB I_Na is substantially reduced in vivo (Ribera and Nüsslein-Volhard 1998). By recording from neurons in vivo, we avoided effects that might arise in vitro if the mao gene had noncell autonomous actions. We used MOs to reveal the contributions of specific molecular isotypes to RB I_Na at each developmental stage.

Previously, recordings were obtained from RB cells in situ in preparations of the developing spinal cord (Ribera and Nüsslein-Volhard 1998). However, the long axonal processes of RB cells led frequently to space-clamp problems that precluded detailed biophysical examination of the voltage-dependent properties of I_Na. In this study, we avoided this difficulty by creating nucleated patches from RB cells in intact preparations. To form a nucleated patch, we first established the conventional whole cell configuration (Hamill et al. 1981). Next, we isolated the soma from the processes by lifting the cell body out of the spinal cord (Fig. 1; Sather et al. 1992). The formation of the nucleated patch resulted in a more compact spherical cell body geometry that avoided space-clamp problems and improved control of membrane voltage. In addition, the use of nucleated patches focused our study on currents present in or near the vicinity of the soma.

View larger version (64K): [in this window] [in a new window]   FIG. 1. Formation of nucleated patches from Rohon–Beard (RB) neurons in vivo. A: RB neurons were visualized with DIC/Nomarski optics in the dorsal region of a semi-intact preparation of the zebrafish spinal cord (48 hpf, left panel). Recording pipette indicates one RB cell in the preparation; asterisk marks a second RB cell. Procedure for formation of nucleated patches consisted of first establishing a whole cell recording configuration. In this configuration, the capacitative transient decayed slowly and its fit often required more than one exponential. Consistent with this, the sodium currents recorded in this configuration were prone to poor voltage control (right panel). B: by raising the pipette (left panel), the RB soma was physically disconnected from its processes, allowing the formation of a nucleated patch (Sather et al. 1992). In the nucleated-patch configuration, the capacitative transient decayed more quickly and had a slightly smaller peak amplitude than that obtained from the same cell in the conventional whole cell configuration. Importantly, voltage control of the membrane was improved (right panel). Once established, nucleated patches provided stable recordings for as long as >5 min.

We recorded I_Na from RB nucleated patches excised from embryos at 3 developmental stages: 16–18 (16), 24–26 (24), and 48–50 (48) hpf (Fig. 2). At all stages, the reversal potential (E_rev) was about 46 mV, close to the calculated value of E_Na.

Peak I_Na amplitude increased almost 6-fold between 16 and 48 hpf (16 = 519 ± 64, n = 9; 24 = 1,080 ± 123, n = 14; 48 = 2,930 ± 235, n = 14; 24 or 48 vs. 16 hpf: P < 0.001; Fig. 2B). In RB cells of older embryos, I_Na peak amplitude was elicited at more negative voltages (Fig. 2C). Between 16 and 24 hpf, there was a –10-mV shift in the observed voltage that elicited peak amplitude (16 = –0.01 ± 0.001 mV; 24 = –10.0 ± 0.01 mV). Because current amplitudes also increased during this period, the shift may have reflected voltage errors arising from series resistance. To examine this possibility, we carried out some recordings in the presence of lower concentrations of external sodium to reduce current amplitudes and resultant series resistance voltage errors. Even under these conditions, developmental shifts in voltage activation were observed (Table 1; Fig. 4B).

View larger version (15K): [in this window] [in a new window]   FIG. 4. Developmental regulation of INa activation properties. A: conductance–voltage plots indicated that the midpoint for activation of INa (V1/2) steadily shifted to more negative potentials as development progressed. Data presented are for the cells shown in Fig. 2; "n" ranged between 9 and 14 for each developmental stage. B: regardless of the external sodium concentration level, INa recorded from RB cells of 48 hpf embryos displayed the negative shift in the activation V1/2. These data indicate that voltage errors arising from series resistance are unlikely to account for the developmental changes noted for the voltage dependency of steady-state activation. C: at each developmental stage examined, V1/2 became significantly more negative (***P < 0.001 vs. 16 hpf). Similarly, the values of k steadily decreased, reflecting the steeper dependency on voltage for activation of INa at later stages of development (**P < 0.01 vs. 16 hpf). D: time-to-peak current decreased between 16 and 24 hpf but showed no further changes at 48 hpf (***P < 0.001 vs. 16 hpf). Inset: time to peak at –30 mV.

In anamniote vertebrates, RB cells constitute a transient population of mechanosensory neurons whose function is later assumed by dorsal root ganglion neurons (Clarke et al. 1984). Mammalian dorsal root ganglion neurons display a substantial I_Na component that is resistant to the blocker TTX (Kostyuk et al. 1981; for review, see McCleskey and Gold 1999). However, RB I_Na was completely abolished at 300 nM TTX, a concentration that spares TTX-resistant current (Blair and Bean 2002; Fig. 3).

View larger version (13K): [in this window] [in a new window]   FIG. 3. Zebrafish RB cells lacked tetrodotoxin (TTX)-resistant currents. TTX-resistant current was not present in RB cells. Current traces shown were evoked by a depolarizing pulse to –10 mV before and after the application of 300 nM TTX. In the presence of 300 nM TTX, INa was completely blocked (n = 16).

To examine voltage-dependent properties of activation more directly, we calculated conductance–voltage relationships (Fig. 4). Statistically significant 5- to 6-mV negative shifts in the midpoint of activation were observed between each developmental stage (16 = –16.2 ± 0.9 mV; 24 = –21.9 ± 1.0 mV; 48 = –27.4 ± 1.1 mV; P < 0.001; Fig. 4, A and B). To minimize voltage errors arising from series resistance that would be most significant for RB cells from 48 hpf embryos, we also performed some recordings under conditions of reduced extracellular sodium (30 and 80 mM; see METHODS). In the presence of 30 mM external sodium, I_Na recorded from RB neurons of 48 hpf embryos had peak amplitudes (433 ± 71 pA, n = 4) similar to those recorded at 16 hpf (519 ± 64 pA, n = 9). Further, we found that, even under conditions of reduced external sodium, the V_1/2 of activation was shifted to more positive values (80 mM Na⁺: –26.5 ± 3.0 mV, n = 4; 30 mM Na⁺: –27.6 ± 0.3 mV, n = 4) and thus not different from that obtained under control conditions (Table 1).

In addition, the slope factor k also decreased significantly at each developmental stage, reflecting the steeper dependency on voltage for I_Na activation in nucleated patches formed from RB cells in older embryos (16 = –6.0 ± 0.2 mV; 24 = –4.8 ± 0.3 mV; 48 = –3.8 ± 0.4 mV; Fig. 4, A and B; P < 0.01).

Between 16 and 24 hpf, the time to peak current decreased. However, the time to peak did not show further changes at 48 hpf (in milliseconds at –30 mV: 16 = 0.96 ± 0.01; 24 = 0.42 ± 0.01; 48 = 0.44 ± 0.02; P < 0.001; Fig, 4C). The changes in time to peak that occurred between 16 and 24 hpf could have resulted from changes in either activation or inactivation, as discussed in the following text.

Developmental regulation of voltage-dependent properties of inactivation of RB I_Na

We used a standard protocol to examine steady-state inactivation (Fig. 5). In contrast to V_1/2 of activation, the half-inactivation potential for I_Na remained relatively constant during the developmental period examined (16 = –49.7 ± 0.9 mV; 24 = –51.9 ± 1.0 mV; 48 = –52.8 ± 0.8 mV; Fig. 5, A and B). However, the absolute value of the slope factor k increased between 16 and 24 hpf (2.3 ± 0.4 vs. 7.2 ± 0.2 ms, P < 0.0001). No further changes were noted at 48 hpf (7.1 ± 0.2 ms). This change in k reflected the less-steep dependency on voltage of steady-state inactivation of I_Na recorded from RB cells of older embryos (Fig. 5, A and B).

View larger version (16K): [in this window] [in a new window]   FIG. 5. Developmental regulation of INa inactivation properties. A: conductance–voltage plots indicated that the voltage dependency of inactivation displayed a steeper relationship for data obtained at younger stages of development. Data presented are for the cells shown in Fig. 2. Inactivation protocol is on top of the panel. B: values of k increased between 16 and 24 hpf (***P < 0.001 vs. 16 hpf), but showed no further change at 48 hpf. Values of V1/2 remained relatively constant during the developmental period studied. C: time constant for inactivation of the transient current showed different developmental regulation between 16 and 24 hpf vs. 24 and 48 hpf (–10 mV: P < 0.0001 for 24 or 48 vs. 16 hpf; 0 mV: P < 0.001 48 vs. 16 hpf; 10 mV: P < 0.05 for 24 vs. 48 hpf). D: time for recovery from inactivation did not differ for INa recorded at any of the 3 developmental stages examined. Inset: voltage protocol used to assay recovery from inactivation.

These data suggested that changes in fast inactivation could have contributed to decreases in the time to peak observed between 16 and 24 hpf at voltages –10 mV, potentials at which the largest changes were observed. However, at voltages positive to 0 mV, the increase in between 16 and 24 hpf opposes the observed decrease in time to peak (Fig. 4C). Thus the data support changes in both activation and inactivation rates of I_Na between 16 and 24 hpf.

The time course of recovery from inactivation was examined at each developmental stage using a standard double-pulse protocol (Fig. 5D). At the 3 stages examined, recovery from inactivation occurred with similar time courses. The current recovered 50% of its initial amplitude within about 2 ms (n = 6, for each stage).

Molecular dissection of RB I_Na

The large changes observed in I_Na amplitude suggested that the density of functional sodium channels increased during this developmental period. If the upregulated channel type had properties different from those of preexisting ones, changes in voltage-dependent properties of activation and inactivation would also result. Alternatively, the observed changes in channel properties could result from posttranslational modification of channels. We used an antisense approach to explore these possibilities.

RB cells express 2 different sodium channel -subunit genes: Na_v1.1 and Na_v1.6 (Novak and Ribera 2004; Tsai et al. 2001). We injected MO1.6, MO1.1, and control MOs and then assayed sodium currents as described above for RB cells in wild-type (WT) uninjected embryos. Control MOs had no effect on sodium current amplitude, regardless of the dose (Fig. 6A). In contrast, injection of MO1.6 or MO1.1 led to dose-dependent decreases in the amplitude of RB I_Na. Further, only a subset of neurons responded to injection of MO1.1 (Fig. 6B, columns in black). In contrast, all RB neurons of MO1.6-injected embryos displayed reduced sodium-current amplitudes (Fig. 6B, gray column) when compared with data obtained from control embryos (white columns).

View larger version (16K): [in this window] [in a new window]   FIG. 6. Morpholinos (MOs) inhibited sodium current components selectively and in a dose-dependent manner. A: dose–response curve for inhibition of RB INa at 48 hpf indicated that control MOs did not affect INa amplitude. However, MO1.1 and MO1.6 reduced INa amplitude in a dose-dependent manner. B: in the majority of neurons, INa amplitude was affected by injection of MO1.6 (gray column). In contrast, INa amplitude was unaffected in many cells by injection of MO1.1 (black columns). White columns show the control INa amplitude values. Frequency range is given in 250-pA increments. C: potassium current amplitude was unaffected by injection of either MO1.1 or MO1.6.

View larger version (14K): [in this window] [in a new window]   FIG. 7. Injection of MO1.6 reduced the behavioral response to tactile stimulation. MO1.6 and control-MO embryos were scored behaviorally to test for the effects on touch sensitivity. Touch sensitivity was assayed by touching the dorsal trunk of an embryo with a probe and scoring the behavioral response as follows: 0 = no response, 1 = normal response, and 0.5 = abnormal response. Each embryo was tested 10 times and the scores for each trial were summed. Thus behavioral scores ranged between 0 and 10. Touch sensitivity was reduced by injection of the MO1.6 but not the control-MO. With time, the behavioral score of MO1.6-injected embryos improved. *P < 0.03 vs. stage-matched control; ***P < 0.001 vs. stage-matched control.

To get an idea of the current components that were sensitive or not to MO1.1, we compared exemplar sodium current recordings from 48 hpf control embryos to those obtained from 48 hpf MO1.1-injected embryos. Further, for this comparison, the y-axis (amplitude) was scaled so that the exemplars reflected the mean amplitudes obtained either for controls (Fig. 8A, black trace) or MO1.1-injected (Fig. 8A, green trace) embryos. For example, at 48 hpf, RB neurons of 1.1 morphant embryos displayed an average 25% reduction in sodium current amplitudes when compared with stage-matched WT embryos (1. 8 ± 0.2 vs. 2.9 ± 0.2 nA; P < 0.001; Fig. 8A). Thus the MO1.1-resistant current trace (Fig. 8A, green trace) was scaled to have an amplitude that was about 60% of that obtained from control (Fig. 8A, black trace). Then, we subtracted these 2 exemplar traces from each other to obtain a difference current that reflects the properties of MO1.1-sensitive current (Fig. 8A, red trace).

View larger version (26K): [in this window] [in a new window]   FIG. 8. MO1.1 removed a slowly activating and inactivating INa component. A: at 48 hpf, MO1.1-resistant INa displayed faster activation, inactivation, and smaller current amplitude than did INa recorded from stage-matched control MO-injected embryos. Superimposed currents were elicited at –10 mV. MO1.1-"sensitive" current, obtained by subtracting the resistant from wild-type (WT) current, showed slow activation and inactivation. B: conductance–voltage plots revealed that MO1.1-resistant INa displayed a negative shift (about 10 mV) in the activation midpoint between 24 and 48 hpf. C: shift of about 10 mV in the V1/2 for inactivation for MO1.1-resistant INa occurred between 24 and 48 hpf. D: transient MO1.1-resistant INa inactivated more quickly in neurons of older embryos (P < 0.001, between –10 and 0 mV).

The activation V_1/2 of the MO1.1-resistant current shifted to more negative values between 24 and 48 hpf (–18.1 ± 0.1 mV vs. 29.1 ± 1.6; P < 0.001; Fig. 8B). The slope value k did not show any statistically significant change (5.8 ± 0.1 vs. 5.4 ± 0.8, respectively).

The V_1/2 and k for steady-state inactivation also displayed developmental differences (Fig. 8C). For example, V_1/2 shifted to more negative values (24 = –50.2 ± 0.8 mV; 48 = –59.4 ± 0.6 mV; P < 0.001). The absolute value of k revealed a slight change diminishing the voltage sensitivity (6.4 ± 0.3 at 24 hpf vs. 8.2 ± 0.8 at 48 hpf; P < 0.05). In comparison to these properties, the time constant of fast inactivation of the MO1.1-resistant current displayed smaller differences between 24 and 48 hpf (Fig. 8D).

Injection of MO1.6 removed a large-amplitude, rapidly activating I_Na component

In contrast to the effects of MO1.1, RB neurons of 1.6 morphant embryos displayed dramatically reduced sodium-current amplitudes (Fig. 9A). The exemplar MO-resistant and MO-sensitive traces were obtained as described for Fig. 8. The MO1.6-resistant current seems to activate more slowly than did currents recorded from wild-type uninjected or control-injected embryos (Fig. 9A). For comparison, we subtracted the MO1.6-resistant sodium current from stage-matching WT I_Na at –10 mV. The difference current, reflecting the I_Na component that was sensitive to MO1.6, seemed to reach time to peak more rapidly than did control currents (e.g., Fig. 2). Further, although the majority of the resistant current inactivated rapidly, a small persistent current was observed similar to what was present in control cells (e.g., Fig. 3). In these respects, the MO1.6-difference current was similar to the MO1.1-resistant current.

View larger version (26K): [in this window] [in a new window]   FIG. 9. MO1.6 targeted a rapidly activating and inactivating, large-amplitude INa component. A: at 48 hpf, MO1.6-resistant current showed both slow activation and inactivation and was small in amplitude vs. control. Difference current (MO1.6-sensitive component) showed a large-amplitude current with fast activation and inactivation. B: conductance–voltage plots revealed that MO1.6-resistant INa displayed a positive shift in the activation midpoint for data obtained at 24 vs. 48 hpf. C: similar behavior was observed in the conductance–voltage plots of inactivation for the MO1.6-resistant INa. D: transient current showed slower inactivation in older embryos (P < 0.001 at all voltages).

The voltage midpoint for steady-state inactivation of the MO1.6-resistant current displayed a shift toward less-negative values as development proceeded (–52.1 ± 0.8 vs. –38.9 ± 1.1 mV; P < 0.05; Fig. 9C). Further, the absolute value of k changed, reflecting the observed developmental reduction in voltage sensitivity (6.8 ± 0.4 vs. 9.2 ± 0.5 mV; P < 0.05, 24 vs. 48 hpf, respectively). Additionally, a decrease in the time constant for fast inactivation was observed (Fig. 9D).

Developmentally regulated effects of MO1.1 and MO1.6

Injection of MO1.6 had substantial effects on current amplitude at all stages examined, suggesting that Na_v1.6 channels underlie a large component of RB I_Na between 16 and 48 hpf. In contrast to MO1.6, the MO1.1 affected only a subset of RB neurons at each developmental stage examined (Fig. 10A), indicating that, at all stages examined, Na_v1.1 channels were present in only a subset of RB cells.

View larger version (20K): [in this window] [in a new window]   FIG. 10. Effects of MOs 1.1 and 1.6 differed. A: effects of MOs 1.1 and 1.6 differed as development proceeded. At 16 hpf, MOs 1.1 and 1.6 produced a similar, 60–70% average reduction in peak INa amplitude. However, at 48 hpf, MO1.1 produced a reduction of only about 25% in current amplitude in contrast to the nearly 85% reduction achieved by MO1.6. B: time to peak was a property of RB INa that showed developmental regulation. Control MOs had no effect on time to peak. In contrast, MO1.1 and MO1.6 had significant effects on time to peak (P < 0.001). Moreover, injection of MO1.1 reduced time to peak values, revealing a resistant INa component that activated more rapidly. Conversely, injection of MO1.6 increased time to peak values, revealing a resistant INa component that activated more slowly. Inset: time to peak values obtained at –30 mV (open triangle, control; black diamond, MO1.1-mis; gray diamond, MO1.6-mis; black triangle, MO1.1; gray triangle, MO1.6; ***P < 0.001 vs. control).

The time to peak of sodium current was a key developmentally regulated property of RB I_Na in WT embryos (Fig. 4C). Injection of MO1.6 prevented the normal developmental decrease in time to peak and the MO1.6-resistant current had times to peak that were significantly greater than those of I_Na in either WT or control-injected neurons (Fig. 10B). Further, the time to peak of MO1.6-resistant current resembled that of I_Na recorded from 16 hpf WT embryos (at –30 mV: 1.4 ± 0.1 vs. 1.0 ± 0.01 ms). In contrast, injection of MO1.1 significantly reduced the time to peak (Fig. 10B). Control MOs had no effect on the time to peak when compared with wild-type (WT = 0.44 ± 0.02; MO1.1-mis = 0.48 ± 0.08 and MO1.6-mis = 0.42 ± 0.08 ms at –10 mV). Overall, the results supported the possibility that Na_v1.6 channels underlie the developmentally regulated increase in I_Na amplitude observed for RB neurons between 16 and 48 hpf (Fig. 2).

I_Na of mao mutants resembled that of 1.6 morphants and 16 hpf embryos

During development, wild-type zebrafish embryos acquire the ability to respond to tactile stimuli by 24–27 hpf, displaying a stereotypical escape response (Kimmel et al. 1995; Saint-Amant and Drapeau 2000). However, mao homozygous mutant embryos (Granato et al. 1996) do not acquire a touch response at 27 hpf and remain touch-insensitive thereafter. The gene harboring the mao mutation has not yet been identified. Ribera and Nüsslein-Volhard (1998) demonstrated that RB I_Na of mao mutants does not undergo the developmentally regulated increase in sodium current, suggesting that a larger sodium current density was critical for acquisition of the touch response. On the basis of results we obtained using MO1.1 and MO1.6, one possibility is that mao mutants lack Na_v1.6 channels. To test this possibility, we characterized the biophysical properties of RB I_Na in mao mutants and compared them to those of MO1.1- and MO1.6-resistant currents.

We recorded RB I_Na from mao mutants (–/–) and behaviorally unaffected siblings (+/?). RB cells of mao mutants displayed sodium currents of much smaller amplitude (–0.5 ± 0.1 nA; n = 12) versus those recorded from siblings (–2.0 ± 0.2 nA, n = 20; P < 0.001; Fig. 11, A and C). Further, the amplitude of I_Na in RB cells of mao mutants was similar to that recorded from RB cells of 16 hpf WT embryos (–0.5 ± 0.1 nA; Fig. 2) and of 48 hpf MO1.6-injected embryos (–0.9 ± 0.2 nA; Fig. 8). Interestingly, the average peak current amplitude recorded from RB cells of mao siblings was smaller than that recorded from WT embryos at the same developmental stage (–1.8 ± 0.1 vs. –2.9 ± 0.2 nA; n = 14; P < 0.001; Fig. 11C; cf. Figs. 2 and 10). This result suggested a heterozygous phenotype that was not detected at the behavioral level. If so, because siblings represent a mixture of wild-type and heterozygous embryos, some of the current amplitudes recorded from sibling embryos (about 1/3) would be in the range of wild-type values, whereas the remainder (about 2/3) would be reduced. In fact, analysis of the I_Na amplitudes in mao unaffected siblings supported the existence of a heterozygous phenotype (Fig. 11D).

View larger version (22K): [in this window] [in a new window]   FIG. 11. Voltage-dependent properties of RB INa recorded from mao mutant (–/–) and unaffected sibling embryos (mao +/?). A: RB sodium currents had significantly smaller amplitudes in 48 hpf mao mutant (right) vs. sibling embryos (left). B: I–V relationships for INa recorded from mao (black hexagons) and sibling (gray hexagons) RB cells. Peak INa amplitude recorded from mao embryos was similar to that of 16 hpf wild-type embryos (Fig. 2). INa of RB cells in sibling embryos had significantly larger amplitude vs. that of mao mutants (P < 0.001 vs. wild-type). C: peak current amplitude of mao mutants was similar to that recorded from wild-type embryos at 16 hpf and to 48 hpf MO1.6-injected embryos. Wild-type data are from Fig. 2. D: distribution of peak INa amplitudes recorded from RB neurons in mao +/? embryos were suggestive of a heterozygous phenotype.

View larger version (12K): [in this window] [in a new window]   FIG. 12. Activation and inactivation properties of INa of RB cells in mao mutant and sibling embryos. A: activation conductance–voltage relationship revealed that the midpoint of activation of INa of mao mutant (black hexagons) vs. sibling embryos (gray hexagons) differed by about 14 mV (V1/2 = –18.8 ± 1.4 vs. –32.5 ± 2.9, mutants and siblings, respectively; P < 0.001). Slope factors did not differ (k = 5.6 ± 0.7 mutants vs. 4.4 ± 0.9 siblings). "n" was 9 for each group. B: time to peak values indicated that RB INa of mao –/– embryos resembled that of 1.6 morphants (–30 mV). Morphant data are from Fig. 10. INa of mao mutants and 1.6 morphants showed larger time to peak values vs. wild-type embryos (P < 0.001 vs. WT). Interestingly, the time to peak obtained from MO1.6 morphants was not significantly different from that obtained from mao homozygous mutants. C: inactivation conductance–voltage relationships for INa of mao mutant vs. sibling embryos were similar. "n" was 10 for each group.

Thus on the basis of current amplitude and time to peak, the possibility that mao mutants lack developmental upregulation of Na_v1.6 channels is supported. Further, voltage-dependent properties of activation were similar for I_Na recorded from mao mutants and 1.6 morphants. However, steady-state properties of inactivation differed between mao mutants and 1.6 morphants, raising the possibility that the mao mutation has effects that cannot be accounted for by a lack of Na_v1.6 channels in RB cells (Table 1).

	DISCUSSION

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The goal of this study was to identify the molecular basis of in vivo developmental regulation of sodium current in neurons of the embryonic vertebrate nervous system. Previous studies examined embryonic neurons developing in culture or after limited periods of time after dissociation achieved by enzymatic treatment (Dourado and Dryer 1992; Gao and Ziskind-Conhaim 1998; Huguenard et al. 1988; MacDermott and Westbrook 1986; Nerbonne and Gurney 1989; Schmid and Guenther 1998; Skaliora et al. 1993). To avoid the possibility that growth in culture or enzymatic treatment altered currents, we recorded from neurons present in semi-intact preparations of the developing spinal cord of the zebrafish embryo. The zebrafish embryo also allowed genetic and molecular approaches that allowed a multilevel analysis of current components in an identified cell type, sensory RB cells. Finally, to ensure adequate control of membrane voltage we formed nucleated patches and restricted our studies to currents present in or near the RB soma.

Our results reveal significant changes in the density and properties of voltage-gated sodium current in RB cells between 16 and 48 hpf (Fig. 2). Peak I_Na amplitude increased almost 6-fold during this period. Fast inactivation also accelerated (Fig. 5), probably contributing to the decrease in time to peak (Fig. 4). Moreover, the voltage of half-activation became more negative (Fig. 4). These changes would contribute to a developmentally regulated increase in excitability of RB cells.

To examine the molecular basis of developmental changes in I_Na, we used antisense morpholinos to knock down expression of protein coded for by Na_v1 genes expressed in RB cells, i.e., Na_v1.1 and Na_v1.6 (Novak et al. 2004). Morpholinos have proven to be highly effective and specific tools for gene inactivation (Nasevicius and Ekker 2000). MOs have been used successfully to knock down the function of specific ion channels (Brent and Drapeau 2002; Sidi et al. 2003). Nonetheless, nonspecific effects can occur with their use (Nasevicius and Ekker 2000). Our tests of specificity, however, indicated that, as used here, the Nav1.1 and Nav1.6 MOs targeted their intended sodium channel isotypes selectively and without nonspecific effects. First, we found that effects were dependent on dose (Fig. 6). For each targeted gene, we obtained similar results with 2 different MOs. Further, effects on I_Na but not I_Kv were observed. Control MOs consisting of 5 base mismatches did not affect I_Na. Moreover, we obtained different results both at the cellular (i.e., I_Na) and behavioral (Fig. 7) levels with MO1.6 versus MO1.1.

Our studies revealed that knockdown of Na_v1.6 prevented the developmentally regulated increase in sodium current density, the decrease in time to peak, and negative shift in the activation V_1/2 (cf. Fig. 4 and Figs. 9 and 10). These data suggest that increased expression of Na_v1.6 channels constitutes a molecular basis for the developmental changes in RB I_Na. However, the MO1.1-resistant current, to which Na_v1.6 channels presumably make the major contribution, displayed substantial changes in voltage-dependent properties between 24 and 48 hpf. These latter results suggest that in addition to increasing the number of Na_v1.6 channels, developmental regulation also entails posttranslational modifications of specific channel properties, including activation and inactivation V_1/2 values and the inactivation (Fig. 9). Posttranslational modifications, such as coassembly with Na_v subunits, glycosylation, or phosphorylation of Nav1 -subunits, can modify these properties for mammalian sodium channels (Tyrrell et al. 2001; for review, see Cantrell and Catterall 2001; Chen et al. 2002, 2004). Whether these modifications occur in a developmentally regulated manner has yet to be addressed.

Our data also indicated that the relative contributions of Na_v1.1 and Na_v1.6 channels to RB I_Na change during development (Fig. 11). The effects of knockdown of Na_v1.1 channels were greater in neurons of 16 hpf embryos. In contrast, MO1.6 had greater effects as development progressed. A straightforward explanation for these findings is that Na_v1.1 channels are gradually replaced by Na_v1.6 channels. However, because our studies were restricted to the vicinity of the soma, it is also possible that both channels are expressed to the same extent in RB neurons at all stages examined, but their subcellular distribution changes.

Between 16 and 48 hpf, Rohon–Beard cells provide sensory function to the embryo because dorsal root ganglion cells do not yet function. Mammalian dorsal root ganglion cells subserve different functional modalities and express multiple sodium channel genes (Benn et al. 2001; Felts et al. 1997). Expression of some Na_v1 isotypes is indicative of a functional class of dorsal root ganglion cells (e.g., Friedel et al. 1997). In contrast to dorsal root ganglion cells, RB cells are often considered to be a homogeneous neuronal population. Consistent with this notion, many molecular markers are expressed in all RB cells (e.g., Hu, HNK-1, P2X, acetylated tubulin; Boue-Grabot et al. 2000; Metcalfe et al. 1990; Svoboda et al. 2001). However, a few molecular markers are expressed in only a subset of RB cells (TrkC, Nav1.1; Martin et al. 1998; Novak and Ribera 2004; Williams et al. 2000). Between 16 and 48 hpf, we found that 2 different sodium channel types, Na_v1.1 and Na_v1.6, underlie sodium current in Rohon–Beard cells. However, knockdown of Na_v1.6 affected all RB cells, whereas only a subset of RB cells were sensitive to the Na_v1.1 MO. These results revealed a functional heterogeneity within the zebrafish RB cell population. These data raise the possibility that, like dorsal root ganglia, the functional diversity of RB cells allows them to subserve additional sensory modalities.

	GRANTS

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This work was supported by National Institute of Neurological Disorders and Stroke Grant NS-38937 to A. B. Ribera.

	ACKNOWLEDGMENTS

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We thank members of the Ribera laboratory for comments on the manuscript and especially B. Hultgren for editorial comments.

	FOOTNOTES

 
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Address for reprint requests and other correspondence: A. B. Ribera, Department of Physiology and Biophysics, Mail Stop 8307, University of Colorado Health Sciences Center at Fitzsimons, P.O. Box 6511, Aurora, CO 80045 (E-mail: angie.ribera{at}uchsc.edu)

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