| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
1Centre de Recherche, Université Laval Robert-Giffard, and Department of Medicine, Université Laval, Quebec City, Quebec, Canada; and 2Molecular Pharmacology Department, AstraZeneca R&D Södertälje, Södertälje, Sweden
Submitted 14 December 2007; accepted in final form 4 March 2008
 |
ABSTRACT |
|---|
|
|
|
INTRODUCTION |
|---|
|
-subunit (260 kDa), which forms the core of the channel and is responsible for the voltage-dependent gating and ion permeation (Armstrong and Hille 1998
; Catterall 1986; Fozzard and Hanck 1996) and auxiliary β-subunits. The -subunit is composed of four homologous domains (DI–DIV), each with six -helical transmembrane-spanning segments (S1–S6). To date, ten mammalian -subunit isoforms have been identified, at least seven of which are expressed in the nervous system. Nav1.1, Nav1.2, Nav1.3, and Nav1.6 are predominantly expressed in the CNS, whereas Nav1.7, Nav1.8, and Nav1.9 are mainly found in the peripheral nervous system (Ogata and Ohishi 2002). Two of these Na+ channels predominate in small nociceptive sensory neurons, a fast inactivating tetrodotoxin (TTX)-sensitive channel (Nav1.7) and a slow inactivating TTX-insensitive channel (Nav1.8), and have been cloned from peripheral sensory nerves.
Nav1.7 may play an important role in nociceptive pain processing (Waxman 2007). Numerous distinct homozygous nonsense mutations in Nav1.7 have been identified in patients with a rare congenital disease that causes a complete inability to sense pain in otherwise healthy individuals (Ahmad et al. 2007; Cox et al. 2006). All mutations described thus far cause a loss of function of this channel. Moreover, a chronic inflammatory dominant human disease, primary erythromelalgia, has been linked to Nav1.7 mutations (Dib-Hajj et al. 2005; Harty et al. 2006; Lampert et al. 2006; Yang et al. 2004), indicating that this channel may play an important role in inflammatory pain. These studies strongly suggest that Nav1.7 is essential for nociception in humans.
Alternative splicing is a mechanism for generating a versatile repertoire of functionally different proteins from a common gene. This mechanism has been observed to occur at five sites for several -subunits encoding voltage-gated Na+ channels from different species (Diss et al. 2004; Kerr et al. 2004; Oh and Waxman 1998; Tan et al. 2002). More specifically, we focused on alternative splicing at two positions of mammalian sodium channel -subunit genes. The first position is in exon 5, which encodes part of segment S3 and all of S4 of domain I in neuronal -subunits. This splicing event produces the 5N or 5A splice forms, which have been described for Nav1.2 (Gustafson et al. 1993; Sarao et al. 1991), Nav1.3 (Gustafson et al. 1993), and Nav1.6 (Plummer et al. 1998; Raymond et al. 2004). The two splice variants differ by either one or two amino acids. Transcripts containing exon 5N predominate in fetal and neonatal brain tissue, whereas in adult brain tissue, the predominant transcripts contain exon 5A. The second position for alternative splicing is in exon 11, which encodes the cytoplasmic loop connecting sodium channel transmembrane domains I and II. This splicing event produces a long (11L) and a short version (11S). Variations in the length of the encoded loop have been observed in Nav1.1 (Schaller et al. 1992), Nav1.3 (Schaller et al. 1992; Thimmapaya et al. 2005), and Nav1.6 (Plummer et al. 1997, 1998; Raymond et al. 2004).
Both alternative splicing events have also been described for Nav1.7 in human dorsal root ganglia (DRG) neurons (Raymond et al. 2004). For this channel, exons 5A and 5N differ by two amino acids in the S3 membrane-spanning segment of domain I (exons 5A and 5N; D1S3:L201V and D1S3:N206D, respectively). In exon 11, two other splice variants are characterized by the presence (11L) or the absence (11S) of an 11 amino acid sequence (VIIDKATSDDS) in the DI–DII linker (see Fig. 1, A and B). The two splicing events can produce four different Nav1.7 isoforms: 5N11S, 5A11L, 5N11L, and 5A11S. Interestingly, Raymond et al. (2004), who also monitored alternatively spliced sodium channel transcripts of Nav1.7 in a rat model of neuropathic pain, observed different levels of expression, with a selective enrichment of transcripts containing exon 11S. This suggests that different splice variants may play a role in different pain states. However, the functional significance of this splicing is not known.
|
|
|
METHODS |
|---|
|
A functional human Nav1.7 cDNA clone (5N11S splice variant) assembled from overlapping PCR fragments (amino acid sequence corresponding to NM_002977 [GenBank] ) and inserted in a cRNA in vitro transcription vector, pKGEM (J. Aiyar, unpublished) was used as the starting material to generate the 5A11S, 5N11L, and 5N11S splice variants. Unique Kpn I and Xho I restriction sites were inserted into the multiple cloning site of pKGEM to allow the transfer of Nav constructs to other vectors (L. Dahllund, unpublished).
The 5A splice variant was generated by replacing two codons in 5N (corresponding to D1S3:L201V and D1S3:N206D) by site-directed mutagenesis (QuikChange II Site-Directed Mutagenesis Kit, Stratagene) using the following two oligonucleotides: 5'-GTT TTT GCG TAT GTA ACA GAA TTT GTA GAC CTA GGC AAT GTT TCA GC-3' and 5'-GCT GAA ACA TTG CCT AGG TCT ACA AAT TCT GTT ACA TAC GCA AAA AC-3'.
The 33-bp 11L insertion was generated by assembling two PCR fragments containing the extra bases in the oligonucleotide sequence. One of the fragments contained the 11L insertion at the 3' end, whereas the other fragment contained the insertion at the 5' end. A unique Spe I site was inserted into the 33-bp fragment, without altering the amino acid sequence, to join the two PCR fragments. The PCR oligonucleotides were 5'-AGC ACA TGA AAA GAG GTT GTC TACC-3' and 5'-GCT GTC ATC ACT AGT TGC CTT ATC TAT TAT CAC CTC TGG CAG AAG CTG TCC ATTG-3' for fragment 1 and 5'-GTG ATA ATA GAT AAG GCA ACT AGT GAT GAC AGC GGC ACG ACC AAT CAA ATA CAC AAG-3' and 5'-AAA TCT GTA CCA CCA AGG TGG ACAT-3' for fragment 2. The assembled PCR fragments were transferred to the 5A and 5N splice variants using Acc I and Bst XI restriction enzymes. The resulting human Nav1.7 splice variants (5A11S, 5N11S, 5A11L, and 5N11L) were then transferred to the pcDNA4/TO (Invitrogen, Carlsbad, CA) expression vector using Kpn I and Xho I restriction sites. All intermediate clones and the final constructs were verified by DNA sequencing of both strands.
Transfection of tsA201 cell line
tsA201 is a mammalian cell line derived from human embryonic kidney HEK 293 cells by stable transfection with SV40 large T antigen (Margolskee et al. 1993). Cells were grown in high glucose DMEM supplemented with fetal bovine serum (10%), L-glutamine (2 mM), penicillin (100 U/ml), and streptomycin (10 mg/ml) (Gibco BRL Life Technologies, Burlington, ON, Canada). The cells were incubated in a 5% CO2 humidified atmosphere. The tsA201 cells were transfected using the calcium phosphate method as previously described (Margolskee et al. 1993).
In all experiments, the human β1-subunit was coexpressed with the human Nav1.7 splice variants. The human Na+ channel β1-subunit and CD8 were constructed in the piRES vector (piRES/CD8/β1). Using this strategy, transfected cells that bind beads (Invitrogen) also express the β1-subunit and are visually distinguishable from nontransfected cells by light microscopy.
For the patch-clamp experiments, 1- to 2-day posttransfection cells were incubated for 5 min in a medium containing anti-CD8-A coated beads (Dynabeads M-450 CD8-a), as previously described (Margolskee et al. 1993). The unattached beads were removed by washing with extracellular solution.
Patch-clamp experiments
Macroscopic currents from transfected tsA201 cells were recorded using the whole cell configuration of the patch-clamp technique. The measurements were carried out at room temperature (
22°C). Fire-polished, low-resistance patch electrodes (1 M
) were pulled from Corning 8161 glass (Dow Corning, Midland, MI) using a model P-97 Flaming/Brown micropipette puller (Sutter Instrument, Novato, CA). The pipettes were coated with Sylgard (Dow Corning) to minimize capacitance. Voltage-clamp experiments were performed using an Axopatch 200B amplifier with a CV 203BU headstage (Molecular Devices, Sunnyvale, CA). Series resistance compensation was performed to values >80% to minimize voltage errors. Voltage command pulses were generated by a personal computer equipped with an AD converter (Digidata 1322A, Molecular Devices) using pCLAMP software v9.0 (Molecular Devices). When appropriate, linear leak currents and capacitance artifacts were removed using P/N leak subtraction. Sodium currents were filtered at 5 kHz and digitized at 200 kHz. The digitized currents were stored on a computer for later off-line analysis.
Sodium currents were generated from a holding potential of –140 mV and depolarized to potentials ranging from –90 to 50 mV for 50 ms in 5-mV increments with 5-s stimulus intervals. The voltage dependence of activation was determined from the relative membrane conductance as a function of potential using the formula gNa = INa/(Vm – Vrev), where gNa is peak conductance, INa is peak sodium current for the test potential Vm, and Vrev is the apparent reversal potential of the sodium current. The resulting sodium ion conductance was normalized to the maximum response for each cell. The voltage dependence of inactivation was obtained by measuring the peak Na+ current during a 20-ms test pulse to –20 mV, which followed a 500-ms prepulse to membrane potentials between –150 and –10 mV from a holding potential of –150 mV. Peak inward currents were measured and normalized to the maximum response of each cell. The activation and inactivation data were fitted with a Boltzman equation, where V1/2 and K represent, respectively, the half-maximum voltage of activation and inactivation and the slope factor: I/Imax = 1/{1 + exp[(V – V1/2)/K]}, and Imax represents the maximum current. The decay time constants were obtained by fitting a single exponential to the current traces. Deactivation was measured using a short (0.5-ms) depolarizing pulse to –20 mV followed by a 20-ms repolarizing pulse to potentials ranging from –100 to –40 mV (
5 mV). Decaying current was then fitted with a single-exponential function. Recovery from inactivation was determined using a paired-pulse protocol in which the time between pulses varied. First and second pulses were a step depolarization from a holding potential of –140 to –20 mV with duration of, respectively, 50 and 20 ms. Interpulse potential durations were increased in length from 0.5 to 1,500 ms at a holding potential of –140 mV. Second pulse peak currents were then normalized to first pulse peak currents and plotted as a function of interpulse time. Data were fitted with a simple exponential. To measure frequency-dependent inhibition, a train of fifty –20-mV pulses was applied at frequencies of 5, 10, 20, 50, and 100 Hz. The peak current elicited by the 50th test pulse was normalized to the current of the first pulse (P50/P1) and was plotted versus frequency. The pulse duration was 8 ms for all frequencies. The holding and interpulse potentials were –140 mV. To measure the time course for the development of inactivation, a prepulse was applied to the cells from a holding potential of –140 mV to the inactivating potentials (from –100 to –60 mV, 10 mV) for increasing durations from 1 to 2,000 ms. Cells were then stepped to the test potential of –20 mV for 20 ms to measure the fraction of current remaining available. Values were then plotted as fraction inactivated versus time of the prepulse and fitted with a single exponential to determine the inactivation time constant. For the ramp current, the cells were held at –140 mV and stimulated with a depolarizing voltage ramp that increased from –140 to 20 mV within 500 ms (0.32 mV/ms). All voltage-clamp protocols were made 10 min after obtaining the whole cell configuration to allow the current to stabilize and ensure adequate diffusion of the contents of the patch electrode.
Solutions and reagents
The patch pipettes were filled with (mmol L–1): 35 NaCl, 105 CsF, 10 EGTA, and 10 HEPES. The pH was adjusted to 7.4 using CsOH. The bath solution contained (mmol L–1): 150 NaCl, 2 KCl, 1.5 CaCl2, 1 MgCl2, 10 glucose, and 10 HEPES. The pH was adjusted to 7.4 using NaOH. A –7-mV correction of the liquid junction potential between the patch pipette and the bath solutions was performed. For the PKA phosphorylation studies, the cells were held in the bath solution containing 1 mM 8-bromo-cyclic adenosine monophosphate (8Br-cAMP, Sigma) for 15 min prior to the experiments. To monitor the effect of 8Br-cAMP on current amplitude, the cells were held in the standard bath solution and perfused with a control solution (standard bath solution) to measure the control current–voltage (I-V) curve. Then, the same solution containing 1 mM 8Br-cAMP was applied to the cell using a microperfusion system. Recordings were taken on a single cell before and during exposure to 8Br-cAMP.
Data analysis
The data were analyzed using a combination of pCLAMP software v9.0 (Molecular Devices), Microsoft Excel, and with either Student's t-test or a one-way ANOVA (SigmaPlot 8.0, SSPS, Chicago, IL). Normality and variance assumptions based on tests and graphs were used to validate the statistical analyses. All reported P values are two-tailed. P < 0.05 values were considered significant. Statistical data are given as means ± SE. The data were analyzed using the statistical package program SAS v9.1.3 (SAS Institute, Cary, NC).
|
|
RESULTS |
|---|
|
Nav1.7 splice variants display similar I-V curve and activation and inactivation parameters.
To study biophysical properties of the four different splice variants, each isoform was transfected with the β1-subunit into tsA201 cells. Whole cell patch-clamp recordings of the four Nav1.7 splice variants (Fig. 1, A, and B) using voltage steps from a holding potential of –140 mV produced typical fast activating and fast inactivating currents for each construct (Fig. 1C). The expression levels of the different variants were not statistically different with a current density of 439.44 ± 63.55, 366.90 ± 85.4, 439.42 ± 66.95, and 316.41 ± 58.80 pA/pF for 5N11S (n = 19), 5A11L (n = 9), 5N11L (n = 15), and 5A11S (n = 15), respectively. The I-V relationships obtained for 5N11S, 5A11L, 5N11L, and 5A11S splice variants are shown in Fig. 2A. No significant difference between the four splice variants was detected. Sodium current activation began at a potential of approximately –65 mV and the peak current was observed at –25 mV for variants with exon 5A and –20 mV for variants with exon 5N.
|
|
The time courses of current decay of the splice variants elicited at depolarized voltages were fitted using a single-exponential function. The resulting time constants were plotted versus voltage (Fig. 3A). No significant differences in inactivation time constants were observed for potentials ranging from –40 to 30 mV. The kinetics of deactivation of the Nav1.7 splice variants, which reflect the transition from the open state to the closed state, were examined by eliciting tail currents at a range of potentials after briefly activating the channel (–20 mV for 0.5 ms). No significant differences in time constants for deactivation (measured with single-exponential fits of tail currents) were observed for potentials ranging from –100 to –40 mV (Fig. 3B).
|
The time course of recovery from inactivation of the splice variants (Fig. 4A) was investigated using a double-pulse protocol. Averaged normalized currents evoked by the second depolarization were plotted as a function of the interpulse interval and fit with a single-exponential equation. There was no significant difference among the four isoforms in terms of time constants (Table 1). Maximum recovery was obtained after 100 ms for all the splice variants.
|
Nav1.7 alternative splicing affects the time course for the development of inactivation and slow ramp current amplitude.
To investigate the development of inactivation, the cells were stimulated with inactivating prepulse potentials of varying durations followed by a test pulse to –20 mV to measure the remaining current (see METHODS). Figure 5A shows the average time course for the development of inactivation fitted with a single exponential and obtained for a prepulse potential of –80 mV. At this potential, all variants reached steady-state inactivation after 500 ms. However, the development of inactivation was significantly (P < 0.01, one-way ANOVA) slower for 5A11L and 5A11S than for 5N11S and 5N11L. 5A11L and 5A11S displayed slower inactivation time constants (59.66 ± 6.32 ms, n = 7 and 61.16 ± 4.52 ms, n = 9, respectively) than 5N11S and 5N11L (41.33 ± 3.65 ms, n = 7 and 37.38 ± 4.44 ms, n = 6, respectively). Figure 5B shows the inactivation time constants obtained for each prepulse potential tested. As at –80 mV, at –90 mV the inactivation time constant was significantly slower (P < 0.05, one-way ANOVA) for 5A11L and 5A11S splice variants (71.80 ± 6.70 ms, n = 7 and 73.50 ± 6.22 ms, n = 9, respectively) than for 5N11S and 5N11L splice variants (55.12 ± 4.16 ms, n = 7 and 49.79 ± 6.77 ms, n = 6, respectively). For more depolarized potentials (–70 and –60 mV), there were no significant differences between exons 5A and 5N.
|
). To test this, we used a slow ramp protocol ranging from –140 to +20 mV in 500-ms increments to depolarize the cells. Peak ramp currents were expressed as a percentage of the transient peak current amplitudes obtained from the initial I-V curves of single cells (Fig. 5C). Interestingly, there were significant exon 5–dependent differences in ramp current amplitudes (P < 0.05, one-way ANOVA). 5A11L and 5A11S displayed significantly higher ramp currents (1.22 ± 0.24%, n = 7 and 1.25 ± 0.21%, n = 9, respectively) than 5N11S and 5N11L (0.63 ± 0.11%, n = 10 and 0.73 ± 0.09%, n = 10, respectively), indicating that alternative splicing of exon 5 modulates the response of Nav1.7 to slow ramp current depolarization, with exon 5A inducing larger ramp currents than exon 5N. Figure 5D shows representative traces obtained during slow depolarization (0.32 mV/ms) of the membrane potentials of 5N11S and 5A11S. The ramp current activated near –75 mV for all variants and no significant differences in the peak ramp current potentials of 5N11S, 5A11L, 5N11L, and 5A11S (–55.02 ± 3.28, –50.65 ± 3.91, –52.54 ± 1.87, and –56.43 ± 2.07 mV, respectively) were observed. Most of the biophysical properties of the four splice variants were similar. However, alternative splicing of exon 5 appeared to modulate the development of inactivation for negative potentials. As a consequence, current amplitudes evoked with slow ramp depolarization were also modulated.
Nav1.7 splice variants exhibit different sensitivities to 8Br-cAMP.
Exon 11 is located in the intracellular loop between domains I and II of Nav1.7, a region of importance for phosphorylation by protein kinase A (PKA). We thus investigated the effects of 8Br-cAMP, a PKA activator, on the Nav1.7 splice variants by treating the cells for 15 min with 1 mM 8Br-cAMP.
Figure 6A shows the effect of 1 mM 8Br-cAMP on the I-V relationships obtained for 5N11S and 5N11L. Although 8Br-cAMP had no significant effect on 5N11L, the 5N11S isoform was significantly affected (P < 0.01, Student's t-test), undergoing a 10-mV shift in peak current to a more hyperpolarized potential. 8Br-cAMP induced a significant 8-mV shift in the midpoint of activation (V1/2) of 5N11S to more negative potentials compared with the control without 8Br-cAMP (P < 0.01, Student's t-test) but had no effect on the slope factor (Kv) (Fig. 6B, Table 1). No significant difference was observed in the V1/2 and Kv of 5N11L in the presence or absence of 8Br-cAMP (Table 1). However, it caused a significant (P < 0.01, Student's t-test) 5-mV negative shift in the V1/2 of activation of 5A11S in the presence of 8Br-cAMP compared with the control, but had no effect on the Kv of activation (Table 1). As with 5N11L, the V1/2 and Kv of activation of 5A11L was not significantly affected by the presence or absence of 8Br-cAMP (Table 1).
|
To determine whether the shift in activation was associated with a variation in current amplitude, 8Br-cAMP was applied to cells with a perfusion system. This allowed us to monitor changes in current amplitudes over time. The current variation, after 15 min of 8Br-cAMP perfusion, expressed in percentage of the initial control current amplitude recorded before 8Br-cAMP perfusion was –4.7 ± 4.5% (5N11S, n = 4) and –2.9 ± 2.1% (5N11L, n = 3) and was not significant. Therefore 8Br-cAMP treatments did not significantly change maximum current amplitude of 5N11S and 5N11L splice variants. Figure 6D shows typical I-V relationships obtained for 5N11S before (perfusion of control solution) and after 15 min of 1 mM of 8Br-cAMP perfusion.
|
|
DISCUSSION |
|---|
|
; Kerr et al. 2004; Oh and Waxman 1998; Schaller et al. 1992; Tan et al. 2002). In the present study, we showed that alternatively spliced variants of the human Nav1.7 channel (5N11S, 5A11L, 5N11L, and 5A11S) can have different functional properties.
In our study, most of the biophysical properties of Nav1.7 splice variants were very similar. The change of two amino acids in the S3 domain I (exons 5A and 5N; D1S3:L201V and D1S3:N206D, respectively) induced a small 3-mV shift of activation to more hyperpolarized potentials for exon 5A compared with exon 5N. However, this shift was significant only between 5N11S and 5A11S splice variants. Since this difference was not significant for other variants, it is possible to say that alternative splicing of exon 5 has no effect on activation parameters of human Nav1.7. Variants of exon 11 had no significant differences in activation parameters. Alternative splicing of exon 5 and exon 11 had no significant effects on the steady- state inactivation, inactivation kinetics for depolarized potentials, and deactivation time constants. These results are in agreement with the study of Thimmapaya et al. (2005) on the Nav1.3 channel. Indeed, these authors investigated the functional properties of Nav1.3 exon 12 splice variants, which are very similar to those of Nav1.7 exon 11 splice variants. They observed minimal differences (2 mV) in the activation parameters of the isoforms and no differences in inactivation parameters. Our results are also in agreement with those of Dietrich et al. (1998), who studied exon 12 (which corresponds to exon 11 of Nav1.7) splice variants of Nav1.6. They observed no significant differences in steady-state activation and steady-state inactivation parameters. However, the insertion of amino acids (corresponding to exon 11L of Nav1.7) resulted in a faster time constant of inactivation for depolarized potentials and a faster recovery from inactivation than the other isoform. In our study, we did not see these differences. In fact, the splice variants in our study with an 11 amino acid insertion in exon 11 displayed similar recoveries from inactivation, which is in agreement with the observations of Thimmapaya et al. (2005). Moreover, in our experiments, there were no significant differences in the frequency-dependent inhibition of the Nav1.7 splice variants. Although alternative splicing did not seem to affect most of the biophysical properties of Nav1.7, the development of inactivation at polarized potentials was significantly slower for variants with exon 5A than those with exon 5N. This difference was related to the higher current elicited by slow ramp depolarization in splice variants containing exon 5A. The link between a slower development of inactivation for negative potentials and a larger slow ramp current amplitude has also been reported (Cummins et al. 1998). In this paper on Nav1.4 and Nav1.7, Cummins et al. (1998) showed that Nav1.7 produced a larger slow ramp current than Nav1.4. They proposed that this difference was a consequence of a slower development of inactivation for Nav1.7. This hypothesis was confirmed by the use of divalent cations such as Cd2+ and Zn2+ that slowed the development of inactivation and also increased ramp current for Nav1.7 but not for Nav1.4. In our study, channels with exon 5A inactivate more slowly at negative potentials than channels with exon 5N, causing a delay in inactivation for 5A variants compared with 5N during a slow depolarization ramp protocol. As a consequence, fewer channels are inactivated at the potential range where the inward current is present. This increase in channel availability will therefore induce larger inward current amplitudes. In our study, peak ramp currents were approximately –55 mV, which is close to the resting potential of DRG neurons (Zheng et al. 2006). Larger ramp current amplitudes might thus increase cell excitability by boosting small stimuli in DRG neurons. Indeed, several studies on Nav1.7 mutations in erythermalgia have correlated a larger ramp current amplitude with an increase in DRG excitability (Cummins et al. 2004; Han et al. 2006; Harty et al. 2006; Lampert et al. 2006; Rush et al. 2006). In our study, Nav1.7 channels containing exon 5A displayed a larger ramp current than those containing exon 5N. Mutations involved in erythermalgia that increase the ramp current might thus be more harmful in the presence of exon 5A. Interestingly, symptoms of primary erythermalgia arise in childhood or adolescence and may progress and become constant with age (Drenth et al. 1996; Lee et al. 2007). In Nav1.2, 1.3, 1.5, exon 5N is the neonatal isoform and is replaced progressively with age by exon 5A (Chioni et al. 2005; Gustafson et al. 1993; Ou et al. 2005; Sarao et al. 1991). If, like Nav1.2, 1.3, and 1.5, exon 5 variants of Nav1.7 are developmentally regulated, the increased symptoms of erythermalgia with age might be related to an increase in the proportion of exon 5A. Similarly, changes in the proportions of exons 5N and 5A with age might also be related to differences in pain sensitivity. However, the developmental regulation of exon 5 variants of Nav1.7 has never been studied and would be of great interest.
The biophysical properties of Nav1.7 were not affected by the alternative splicing of exon 11, which is located in the intracellular loop between domains I and II. This region possesses several PKA phosphorylation sites and is therefore important in sodium channel modulation (Cantrell et al. 2002; Chahine et al. 2005; Murphy et al. 1993; Smith and Goldin 2000). cAMP-dependent phosphorylation of the four Nav1.7 isoforms revealed that the activation and steady-state inactivation of the exon 11L splice variants were not affected by 8Br-cAMP, whereas the activation of the exon 11S isoforms was strongly affected, with a significant shift to hyperpolarized potentials (5 to 8 mV). Steady-state inactivation, however, was not affected. The insertion of the 11 amino acid sequence (VIIDKATSDDS) in the intracellular loop between domains I and II thus suppressed the effects of 8Br-cAMP. The cAMP-mediated PKA phosphorylation of rat Nav1.7 in Xenopus oocytes decreases the peak current without changing the gating properties of the channel (Vijayaragavan et al. 2004). However, the rat Nav1.7 isoform used by Vijayaragavan et al. (2004) contained the 11 amino acid insertion, which might explain why there was no cAMP-mediated effect on gating. In our study, we studied the effect of 8Br-cAMP on current amplitude using a perfusion system. The shift in activation was not associated with a change in the maximum current amplitude at peak potential. Similarly, there was no significant change in current amplitude for the 11L splice variants. The mechanism involved in the disruption of the cAMP effect by the insertion of these 11 amino acid residues remains to be elucidated. It is possible that the insertion has a major effect on the structure of the intracellular loop and thus significantly changes the availability of potential phosphorylation sites. A second possibility is that another protein is involved in the cAMP effect. One candidate could be an A kinase-anchoring protein like AKAP-15. Indeed, AKAP-15 has been shown to be essential for PKA-mediated sodium channel phosphorylations and to interact with the cytoplasmic loop connecting transmembrane domains I and II (Cantrell et al. 1999, 2002; Few et al. 2007; Tibbs et al. 1998). In a recent study, the binding site of AKAP-15 was identified to be a modified leucine zipper motif localized at the beginning of the intracellular loop between transmembrane domains I and II of Nav1.2 (Few et al. 2007). Whereas this binding motif is not localized within exon 11, the incorporation of the 11 amino acid residues might change the conformation of the intracellular loop and thus disrupt the binding site of AKAP-15. In these conditions, PKA-mediated phosphorylation could be unsettled. This might explain the absence of effects of 8Br-cAMP observed on splice variants that contain exon 11L.
There is increasing evidence in the literature that Nav1.7 plays a critical role in pain, as in erythromelalgia (Cummins et al. 1998, 2004; Dib-Hajj et al. 2005; Han et al. 2006; Lampert et al. 2006; Rush et al. 2006; Yang et al. 2004) and in diseases involving a complete inability to sense pain (Ahmad et al. 2007; Cox et al. 2006), indicating that changes in the activity of this channel might have a significant impact on pain sensitivity. The splice variants of Nav1.7 have been detected in human DRG neurons (Raymond et al. 2004). In this study, the authors investigated the relative proportions of these variants in a rat model of neuropathic pain. They observed a significant change in the expression pattern of alternatively spliced isoforms. Exon 11S–containing transcripts increased in relative abundance in response to neuropathic injury, indicating that the exon 11S isoform may be involved in pain generation. Although we observed no biophysical differences between the exon 11S and 11L splice variants, the activation of the exon 11S isoforms was significantly and negatively shifted in the presence of cAMP, whereas this messenger had no effect on channels containing exon 11L. This might explain the correlation between neuronal hyperexcitability and an increase in expression of exon 11S–containing channels. Interestingly, the cAMP–PKA pathway has little effect on the excitability of intact DRG neurons in uncompressed ganglia (Song et al. 2006; Zheng et al. 2006). However, these authors also reported that cAMP significantly increases neuronal hyperexcitability by decreasing the action potential threshold in a rat pain model created by the chronic compression of DRG neurons. This is in agreement with the findings of the present study and with the observations of Raymond et al. (2004). During neuropathic pain, an increase in cAMP combined with an increase in exon 11S Nav1.7 splice variants might induce a negative shift of the activation of sodium currents in DRG neurons. The action potential threshold would thus decrease and neuronal hyperexcitability would increase. However, since there are higher levels of exon 11L in physiological conditions and since this variant is not sensitive to cAMP, this might explain why the generation of action potentials in intact DRG neurons is not affected by cAMP (Song et al. 2006; Zheng et al. 2006).
In conclusion, the present study showed that alternative splicing of human Nav1.7 could alter the biophysical properties and the cAMP-mediated modulation of this channel. Changes in the proportions of the alternative splice variants may facilitate the generation of action potentials, thus contributing to neuropathic pain. Investigating the alternative splicing and cAMP modulation of Nav1.7 may thus be an interesting avenue to pursue in the search for therapies to treat neuropathic pain.
|
|
GRANTS |
|---|
|
|
|
FOOTNOTES |
|---|
Address for reprint requests and other correspondence: M. Chahine, Centre de Recherche, Université Laval Robert-Giffard, Local F-6539, 2601 chemin de la Canardière, Quebec City, QC, Canada G1J 2G3 (E-mail: mohamed.chahine{at}phc.ulaval.ca)
|
|
REFERENCES |
|---|
|
Armstrong CM, Hille B. Voltage-gated ion channels and electrical excitability. Neuron 20: 371–380, 1998.[CrossRef][Web of Science][Medline]
Cantrell AR, Tibbs VC, Westenbroek RE, Scheuer T, Catterall WA. Dopaminergic modulation of voltage-gated Na+ current in rat hippocampal neurons requires anchoring of cAMP-dependent protein kinase. J Neurosci 19: RC21, 1999.
Cantrell AR, Tibbs VC, Yu FH, Murphy BJ, Sharp EM, Qu Y, Catterall WA, Scheuer T. Molecular mechanism of convergent regulation of brain Na+ channels by protein kinase C and protein kinase A anchored to AKAP-15. Mol Cell Neurosci 21: 63–80, 2002.[CrossRef][Web of Science][Medline]
Catterall WA. Molecular properties of voltage-sensitive sodium channels. Annu Rev Biochem 55: 953–985, 1986.[CrossRef][Web of Science][Medline]
Chahine M, Ziane R, Vijayaragavan K, Okamura Y. Regulation of Na(v) channels in sensory neurons. Trends Pharmacol Sci 26: 496–502, 2005.[CrossRef][Medline]
Chioni AM, Fraser SP, Pani F, Foran P, Wilkin GP, Diss JK, Djamgoz MB. A novel polyclonal antibody specific for the Na(v)1.5 voltage-gated Na(+) channel "neonatal" splice form. J Neurosci Methods 147: 88–98, 2005.[CrossRef][Web of Science][Medline]
Cox JJ, Reimann F, Nicholas AK, Thornton G, Roberts E, Springell K, Karbani G, Jafri H, Mannan J, Raashid Y, Al-Gazali L, Hamamy H, Valente EM, Gorman S, Williams R, McHale DP, Wood JN, Gribble FM, Woods CG. An SCN9A channelopathy causes congenital inability to experience pain. Nature 444: 894–898, 2006.[CrossRef][Medline]
Cummins TR, Dib-Hajj SD, Waxman SG. Electrophysiological properties of mutant Nav1.7 sodium channels in a painful inherited neuropathy. J Neurosci 24: 8232–8236, 2004.
Cummins TR, Howe JR, Waxman SG. Slow closed-state inactivation: a novel mechanism underlying ramp currents in cells expressing the hNE/PN1 sodium channel. J Neurosci 18: 9607–9619, 1998.
Dib-Hajj SD, Rush AM, Cummins TR, Hisama FM, Novella S, Tyrrell L, Marshall L, Waxman SG. Gain-of-function mutation in Nav1.7 in familial erythromelalgia induces bursting of sensory neurons. Brain 128: 1847–1854, 2005.
Dietrich PS, McGivern JG, Delgado SG, Koch BD, Eglen RM, Hunter JC, Sangameswaran L. Functional analysis of a voltage-gated sodium channel and its splice variant from rat dorsal root ganglia. J Neurochem 70: 2262–2272, 1998.[Web of Science][Medline]
Diss JK, Fraser SP, Djamgoz MB. Voltage-gated Na+ channels: multiplicity of expression, plasticity, functional implications and pathophysiological aspects. Eur Biophys J 33: 180–193, 2004.[Web of Science][Medline]
Drenth JP, Vuzevski V, Van JT, Casteels-Van DM, Vermylen J, Michiels JJ. Cutaneous pathology in primary erythermalgia. Am J Dermatopathol 18: 30–34, 1996.[CrossRef][Web of Science][Medline]
Few WP, Scheuer T, Catterall WA. Dopamine modulation of neuronal Na(+) channels requires binding of A kinase-anchoring protein 15 and PKA by a modified leucine zipper motif. Proc Natl Acad Sci USA 104: 5187–5192, 2007.
Fozzard HA, Hanck DA. Structure and function of voltage-dependent sodium channels: comparison of brain II and cardiac isoforms. Physiol Rev 76: 887–926, 1996.
Gustafson TA, Clevinger EC, O'Neill TJ, Yarowsky PJ, Krueger BK. Mutually exclusive exon splicing of type III brain sodium channel subunit RNA generates developmentally regulated isoforms in rat brain. J Biol Chem 268: 18648–18653, 1993.
Han C, Rush AM, Dib-Hajj SD, Li S, Xu Z, Wang Y, Tyrrell L, Wang X, Yang Y, Waxman SG. Sporadic onset of erythermalgia: a gain-of-function mutation in Nav1.7. Ann Neurol 59: 553–558, 2006.[CrossRef][Web of Science][Medline]
Harty TP, Dib-Hajj SD, Tyrrell L, Blackman R, Hisama FM, Rose JB, Waxman SG. Na(V)1.7 mutant A863P in erythromelalgia: effects of altered activation and steady-state inactivation on excitability of nociceptive dorsal root ganglion neurons. J Neurosci 26: 12566–12575, 2006.
Kerr NCH, Holmes FE, Wynick D. Novel isoforms of the sodium channels Nav1.8 and Nav1.5 are produced by a conserved mechanism in mouse and rat. J Biol Chem 279: 24826–24833, 2004.
Lampert A, Dib-Hajj SD, Tyrrell L, Waxman SG. Size matters: erythromelalgia mutation S241T in Nav1.7 alters channel gating. J Biol Chem 281: 36029–36035, 2006.
Lee MJ, Yu HS, Hsieh ST, Stephenson DA, Lu CJ, Yang CC. Characterization of a familial case with primary erythromelalgia from Taiwan. J Neurol 254: 210–214, 2007.[CrossRef][Web of Science][Medline]
Margolskee RF, McHendry-Rinde B, Horn R. Panning transfected cells for electrophysiological studies. Biotechniques 15: 906–911, 1993.[Web of Science][Medline]
Murphy BJ, Rossie S, De Jongh KS, Catterall WA. Identification of the sites of selective phosphorylation and dephosphorylation of the rat brain Na+ channel subunit by cAMP-dependent protein kinase and phosphoprotein phosphatases. J Biol Chem 268: 27355–27362, 1993.
Ogata N, Ohishi Y. Molecular diversity of structure and function of the voltage-gated Na+ channels. Jpn J Pharmacol 88: 365–377, 2002.[CrossRef][Medline]
Oh Y, Waxman SG. Novel splice variants of the voltage-sensitive sodium channel alpha subunit. Neuroreport 9: 1267–1272, 1998.[Web of Science][Medline]
Ou SW, Kameyama A, Hao LY, Horiuchi M, Minobe E, Wang WY, Makita N, Kameyama M. Tetrodotoxin-resistant Na+ channels in human neuroblastoma cells are encoded by new variants of Nav1.5/SCN5A. Eur J Neurosci 22: 793–801, 2005.[CrossRef][Web of Science][Medline]
Plummer NW, Galt J, Jones JM, Burgess DL, Sprunger LK, Kohrman DC, Meisler MH. Exon organization, coding sequence, physical mapping, and polymorphic intragenic markers for the human neuronal sodium channel gene SCN8A. Genomics 54: 287–296, 1998.[CrossRef][Web of Science][Medline]
Plummer NW, McBurney MW, Meisler MH. Alternative splicing of the sodium channel SCN8A predicts a truncated two-domain protein in fetal brain and non-neuronal cells. J Biol Chem 272: 24008–24015, 1997.
Raymond CK, Castle J, Garrett-Engele P, Armour CD, Kan Z, Tsinoremas N, Johnson JM. Expression of alternatively spliced sodium channel -subnit genes: unique splicing patterns are observed in dorsal root ganglia. J Biol Chem 279: 46234–46241, 2004.
Rush AM, Dib-Hajj SD, Liu S, Cummins TR, Black JA, Waxman SG. A single sodium channel mutation produces hyper- or hypoexcitability in different types of neurons. Proc Natl Acad Sci USA 103: 8245–8250, 2006.
Sarao R, Gupta SK, Auld VJ, Dunn RJ. Developmentally regulated alternative RNA splicing of rat brain sodium channel mRNAs. Nucleic Acids Res 19: 5673–5679, 1991.
Schaller KL, Krzemien DM, McKenna NM, Caldwell JH. Alternatively spliced sodium channel transcripts in brain and muscle. J Neurosci 12: 1370–1381, 1992.[Abstract]
Smith RD, Goldin AL. Potentiation of rat brain sodium channel currents by PKA in Xenopus oocytes involves the I–II linker. Am J Physiol Cell Physiol 278: C638–C645, 2000.
Song XJ, Wang ZB, Gan Q, Walters ET. cAMP and cGMP contribute to sensory neuron hyperexcitability and hyperalgesia in rats with dorsal root ganglia compression. J Neurophysiol 95: 479–492, 2006.
Tan J, Liu Z, Nomura Y, Goldin AL, Dong K. Alternative splicing of an insect sodium channel gene generates pharmacologically distinct sodium channels. J Neurosci 22: 5300–5309, 2002.
Thimmapaya R, Neelands T, Niforatos W, vis-Taber RA, Choi W, Putman CB, Kroeger PE, Packer J, Gopalakrishnan M, Faltynek CR, Surowy CS, Scott VE. Distribution and functional characterization of human Nav1.3 splice variants. Eur J Neurosci 22: 1–9, 2005.[CrossRef][Web of Science][Medline]
Tibbs VC, Gray PC, Catterall WA, Murphy BJ. AKAP15 anchors cAMP-dependent protein kinase to brain sodium channels. J Biol Chem 273: 25783–25788, 1998.
Vijayaragavan K, Boutjdir M, Chahine M. Modulation of Nav1.7 and Nav1.8 peripheral nerve sodium channels by protein kinase A and protein kinase C. J Neurophysiol 91: 1556–1569, 2004.
Waxman SG. Nav1.7, its mutations, and the syndromes that they cause. Neurology 69: 505–507, 2007.
Yang Y, Wang Y, Li S, Xu Z, Li H, Ma L, Fan J, Bu D, Liu B, Fan Z, Wu G, Jin J, Ding B, Zhu X, Shen Y. Mutations in SCN9A, encoding a sodium channel alpha subunit, in patients with primary erythermalgia. J Med Genet 41: 171–174, 2004.
Zheng JH, Walters ET, Song XJ. Dissociation of dorsal root ganglion neurons induces hyperexcitability that is maintained by increased responsiveness to cAMP and cGMP. J Neurophysiol 97: 15–25, 2006.[Web of Science][Medline]
This article has been cited by other articles:
|
|
 |
|
  C. Han, S. D. Dib-Hajj, Z. Lin, Y. Li, E. M. Eastman, L. Tyrrell, X. Cao, Y. Yang, and S. G. Waxman Early- and late-onset inherited erythromelalgia: genotype-phenotype correlation Brain, July 1, 2009; 132(7): 1711 - 1722. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
TOP
ABSTRACT
INTRODUCTION
