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(Circulation Research. 1996;78:244-252.)
© 1996 American Heart Association, Inc.

Articles

Multiple Domains Contribute to the Distinct Inactivation Properties of Human Heart and Skeletal Muscle Na⁺ Channels

Naomasa Makita, Paul B. Bennett, Jr, Alfred L. George, Jr

From the Departments of Medicine and Pharmacology, Vanderbilt University School of Medicine, Nashville, Tenn.

Correspondence to Dr Alfred L. George, Jr, S-3223 MCN, Vanderbilt University Medical Center, 21st Ave South at Garland Ave, Nashville, TN 37232-2372.

	Abstract

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Abstract Voltage-gated Na⁺ channels are essential for the normal electrical excitability of neuronal and striated muscle membranes. Distinct isoforms of the Na⁺ channel -subunit have been identified by molecular cloning, and their functional attributes have been defined by heterologous expression coupled with electrophysiological recording. Two closely related Na⁺ channel -subunit isoforms, hH1 (human heart) and hSkM1 (human skeletal muscle), exhibit differences in their inactivation properties and in their response to the coexpressed ß₁-subunit. To localize regions that contribute to inactivation and to ß₁-subunit response, we have exploited these functional differences by studying chimeric channels composed of segments from both hH1 and hSkM1. Chimeras in which one or more of the cytoplasmic interdomain regions (ID1-2, ID2-3, and ID3-4) were exchanged between hH1 and hSkM1 exhibit inactivation properties identical with the background channel isoform, suggesting that these regions are not sufficient to cause gating differences. In contrast, inactivation properties of chimeras composed of approximately equal halves of the two channel isoforms were intermediate between hH1 and hSkM1. Furthermore, the response to the coexpressed ß₁-subunit was dependent on structures located in the carboxy-terminal half of the -subunit, although domains D3, D4, and the carboxy terminal are not singularly responsible for this effect. These data indicate that inactivation differences between hH1 and hSkM1 are determined by multiple -subunit domains.

Key Words: ion channel gating • electrophysiology • Na⁺ channel

	Introduction

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Voltage-gated Na⁺ channels are responsible for the rapid membrane depolarization that characterizes the initial phase of the action potential in most excitable cells.¹ A vital property of Na⁺ channels is fast inactivation, an intrinsic mechanism that closes the channel during sustained membrane depolarizations. In myocardium, Na⁺ channel inactivation contributes to the control of membrane refractoriness during repetitive stimulation and is an important determinant of class I antiarrhythmic drug action.² Disturbances in Na⁺ channel inactivation are also responsible for one form of the congenital long-QT syndrome.^{3 4}

There is strong evidence implicating an essential role for the cytoplasmic linker region joining domains 3 and 4 (ID3-4) in fast inactivation.^{5 6 7 8} One critical subregion of ID3-4 required for fast inactivation has been localized by site-directed mutagenesis to a hydrophobic tripeptide sequence of Ile-Phe-Met.⁸ This sequence is highly conserved among virtually all known voltage-gated Na⁺ channels, even though substantial differences exist between channel isoforms in their inactivation kinetics. Thus, other regions of these proteins must also contribute to inactivation. The amino acid sequences of other cytoplasmic interdomain regions (ID1-2 and ID2-3) vary greatly among Na⁺ channel -subunit isoforms, and their role in inactivation is unknown.

One approach to define regions important for inactivation is to exploit known structural and functional differences among Na⁺ channel -subunit isoforms. Recombinant rat skeletal and cardiac Na⁺ channel -subunits exhibit clear differences in both kinetic and steady state properties of inactivation when expressed in Xenopus oocytes.^{9 10 11} The skeletal muscle isoform displays a time course of inactivation that is slower than that of the cardiac channel. The slow inactivation behavior of the rat skeletal muscle Na⁺ channel in oocytes appears in part to be due to the absence of a ß₁-subunit, because coexpression of this subunit greatly accelerates the time course of current decay exhibited by this channel.^{12 13} Similarly, the inactivation kinetics of recombinant hSkM1 Na⁺ channels are accelerated by coexpression with the ß₁-subunit,¹⁴ but cloned hH1 Na⁺ channels do not require this subunit for fast inactivation.¹⁵ These observations suggest that intrinsic structural differences between hH1 and hSkM1 are responsible for the distinct inactivation kinetics and modulatory effects of the ß₁-subunit.

In the present study, we exploited the isoform-specific functional differences in Na⁺ channel inactivation to determine structural determinants of inactivation and ß₁-subunit response by studying chimeras composed of segments from hSkM1 and hH1. Understanding the structural basis for these critical differences will shed new light on the molecular basis for Na⁺ channel gating.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Construction of Chimeric Na⁺ Channels
Chimeric Na⁺ channel cDNAs were constructed from pSP64T-hSkM1^{16 17} and pSP64T-hH1¹⁸ using either site-directed mutagenesis as previously described¹⁹ or the overlap extension PCR technique.²⁰ Correct assembly was verified by restriction analysis and dideoxynucleotide sequencing of junction regions. Chimeras generated by the PCR were sequenced more extensively to identify clones without polymerase errors. In most cases, functional expression studies were performed on multiple independent recombinants. Oligonucleotides were made by ß-cyanoethylphosphoramidite chemistry using a Milligen Cyclone Plus DNA synthesizer.

Construction of ID3-4 Chimeras (hSkM1-C and hH1-C)
Site-directed mutagenesis was used to change all amino acid residues in the hSkM1 ID3-4 to match those of hH1. Initially, two cDNA fragments of WT hSkM1, Sac I–HindIII (nucleotides 2406 to 3973, including the 5' half of ID3-4) and HindIII–Sac I (nucleotides 3973 to 5557, including the 3' half of ID3-4), were subcloned into the plasmid pSELECT, and single-strand DNAs were rescued using R408 helper phage. The following seven mutations were made simultaneously in hSkM1: K1308Q, Q1339L, I1342Y, M1345F, V1346I, Y1347F, and L1349I. Mutagenesis was performed using two mutagenic primers, C1 and C2 (Table 1), to obtain mutant plasmids pSlD3B and pSlD4A, respectively. Full-length pSP64T–hSkM1-C constructs were assembled using a 510-bp Nar I–HindIII fragment from pSlD3B and a 413-bp HindIII–Sac II fragment from pSlD4A. The boundaries of the chimeric sequence coincided precisely with the junction of the ID3-4 region with D3 and D4 (hSkM1 amino acid positions 1295 and 1349).

View this table: [in this window] [in a new window]   Table 1. Oligonucleotides Used in Constructing Chimeric Na+ Channels

Overlap-extension PCR mutagenesis was used for constructing hH1-C with oligonucleotide primers C3, C4, C5, and C6 (Table). A 330-bp segment of hH1 (nucleotides 4092 to 4422) was amplified using primers C3 and C4. In a separate reaction, a 235-bp segment of hSkM1 encompassing the ID3-4 fragment (nucleotides 3877 to 4111) was amplified using primers C5 and C6. Subsequently, the two PCR products were purified from low-melting agarose gel and combined in a second round of PCR with the C3/C6 primer pair. A 545-bp PCR product was digested with Kpn I–BstEII and subcloned back into pSP64T-hH1 to assemble the hH1-C construct. The boundaries of the chimeric sequence coincided with the precise junction of ID3-4 with D3 (hH1 position 1470) and with the amino-terminal portion of D4/S1 (hH1 position 1536).

Construction of ID1-2 Chimeras (hSkM1-A and hH1-A)
Site-directed mutagenesis was used to introduce unique HindIII sites within the corresponding carboxy-terminal region of D1/S6 in both hSkM1 (nucleotide 1302) and hH1 (nucleotide 1200) without changing the amino acid sequence. Similarly, silent BamHI sites were engineered into the amino-terminal region of D2/S1 in both channel cDNAs (hSkM1, nucleotide 1728; hH1, nucleotide 2145) The following subfragments and primers (Table) were used for mutagenesis: hSkM1 Acc I–Sph I, primers A1 (HindIII) and A2 (BamHI); hH1 Xho I–EcoRI, primers A3 (HindIII) and A4 (BamHI). Functional expression studies in Xenopus oocytes were performed to ensure that these silent mutations did not alter WT function. Finally, chimeric ID1-2 channel constructs (hSkM1-A and hH1-A) were made by interchanging the corresponding HindIII-BamHI segment of each channel. Boundaries of the chimeric sequences coincided with the precise junctions of ID1-2 region with D2 (hSkM1 position 574, hH1 position 713) and with the carboxy-terminal portion of D1/S6 (hSkM1 position 439, hH1 position 405).

Construction of ID2-3 Chimeras (hSkM1-B and hH1-B)
A double-overlap extension PCR method was used to construct hSkM-B and hH1-B. Initially, six overlapping PCR primers (Table) were used to amplify the following fragments: hSkM1-SB1 (nucleotides 2119 to 2412, primer set B1/B2), hSkM1-SB2 (nucleotides 2392 to 3100, primer set B9/B10), and hSkM1-SB3 (nucleotides 3079 to 3504, primer set B5/B6). The hH1 fragments hH1-HB1 (nucleotides 2157 to 2823), hH1-HB2 (nucleotides 2804 to 3621), and hH1-HB3 (nucleotides 3601 to 4129) were also amplified by primer sets B7/B8, B3/B4, and B11/B12, respectively. Subsequently, hSkM1-SB1 and hH1-HB2 were annealed, 5'-extended, and then reamplified with the flanking primers B1 and B4 to obtain the 1.0-kb chimeric PCR fragment SB1/HB2. Similarly, a 1.2-kb HB2/SB3 chimeric PCR product was amplified by B3 and B6. Last, a 1.5-kb SB1/HB2/SB3 product was obtained by another round of overlap-extension PCR using SB1/HB2 and HB2/SB2 as overlapping templates with primers B1 and B6. Final PCR products were digested with Sph I and Avr II and subcloned back into the full-length hSkM1 cDNA. Boundaries for the chimeric sequence coincided precisely with the junctions of ID2-3 with D2 (hSkM1 position 802) and D3 (hSkM1 position 1027).

For construction of hH1-B, two chimeric PCR products, HB1/SB2 (1.3 kb) and SB2/HB3 (1.2 kb), were amplified by primer sets B7/B10 and B9/B12, respectively. Each fragment was digested with EcoRI–Xho I or Xho I–Acc I, respectively, and subcloned into the full-length hH1 cDNA to obtain hH1-B. Boundaries for the chimeric sequence coincided precisely with the junctions of ID2-3 with D2 (hH1 position 939) and D3 (hH1 position 1201).

Construction of Triple Interdomain Chimeras (hSkM1-ABC and hH1-ABC)
A 2.8-kb Not I–Nsi I fragment of pSP64T-hSkM1-A and a 1.3-kb Nsi I–Avr II fragment of pSP64T-hSkM1-B were subcloned back into the 5.2-kb Avr II–Not I digestion product of the pSP64T-hSkM1-C construct to obtain the triple interdomain chimera hSkM1-ABC. Similarly, a 1.2-kb Xho I–EcoRI fragment of hH1-A and a 1.5-kb EcoRI–Acc I fragment of hH1-B were subcloned back into the 8.4-kb Acc I–Xho I digestion product of pSP64-hH1-C to construct hH1-ABC.

Construction of Split Chimeras (HS and SH)
The ID2-3 chimeras described above were further used to construct channels that consisted of the amino terminal through ID2-3 of one isoform and D3 through the carboxy terminal of the other isoform. These "split" chimeras were designated as HS and SH with the relative position of hH1 sequences noted by H and that of hSkM1 by S. Unique restriction sites contained within the ID2-3 region (hSkM1 ID2-3, Xho I, nucleotide 2719; hH1 ID2-3, BamHI, nucleotide 3120) were used to assemble cDNA fragments derived from WT and ID2-3 chimeric channels.

Construction of D3, D4, and Carboxy-Terminal Chimeras
A chimera consisting of the hH1 D3 region inserted into the background sequence of hSkM1 (designated as S124H3) was constructed by ligating a 1.2-kb HindIII–Xho I fragment from hH1-ABC into the corresponding region of WT-hSkM1. The HindIII and Xho I sites are located within the D1/S6 and ID2-3 regions, respectively, of the hH1-ABC chimera.

A separate construct consisting of the hH1 D4 region inserted into the hSkM1 background was assembled by ligating a 0.8-kb HindIII–HincII fragment from hH1-ABC into the corresponding region of WT-hSkM1. The HindIII and HincII sites are located within the ID3-4 and D4/S6 regions, respectively, of hH1-ABC.

A carboxy-terminal chimera was constructed by exchanging a HincII–Xba I fragment (hSkM1, 1.3 kb; hH1, 2.2 kb) between the two isoforms. This restriction fragment contains the entire carboxy terminal along with a small region of D4/S6 having identical amino acid sequences in WT-hH1 and WT-hSkM1.

In Vitro Transcription and Oocyte Injection
Both WT and chimeric human Na⁺ channel -subunit cDNAs engineered in the oocyte expression vector pSP64T²¹ were used for in vitro transcriptions. Sense cRNAs were transcribed in vitro in the presence of the methylated 5'-cap analogue (m⁷GpppG) from EcoRI-linearized (hSkM1-background chimeras) or Xba I–linearized (hH1-background chimeras) cDNAs using SP6 RNA polymerase (Boehringer). Similarly, the human ß₁-subunit cDNA in the pSP64T vector was transcribed as previously described.¹⁵ Defolliculated Xenopus oocytes were injected with 40 nL (4 to 20 ng) of 5'-capped cRNAs encoding Na⁺ channels and then incubated at room temperature in ND-96 solution (mmol/L: NaCl 96, KCl 2, CaCl₂ 1.8 , MgCl₂ 1, and HEPES 5 [pH 7.50 with NaOH]) for 1 to 2 days.

Voltage-Clamp Protocols
Whole-cell currents were recorded in oocytes using a two-microelectrode voltage clamp as previously described¹⁴ except that a Warner Instrument voltage-clamp amplifier with virtual bath ground was used. Electrodes were pulled from Radnoti Starbore glass and filled with 3 mol/L KCl. Electrode resistances were 1 to 3 M for voltage-recording electrodes and 0.5 to 1.5 M for current-passing electrodes. Voltage commands were generated by a 16-bit D/A converter driven by custom software running under Microsoft Windows. Currents were filtered at 5 kHz (-3 dB, four-pole Bessel filter) and sampled at 50 kHz by a 16-bit A/D converter.

To assess steady state channel inactivation and recovery, three pulse protocols were employed (single test pulse, double pulse–steady state inactivation, and double pulse–recovery from inactivation). Unless specified otherwise, the holding potential was set to -120 mV, and Na⁺ currents were recorded during test potentials to -20 mV. This test voltage was selected because it is a membrane potential at which most Na⁺ channels will open if available, and it is not on the negative limb of the current-voltage relationship for either hSkM1 or hH1. Analyses were restricted to cells in which the voltage control appeared adequate, as indicated by a narrow (<1 millisecond) capacitative transient and the absence of discontinuities in the current tracings. For assessing steady state inactivation, the membrane potential was stepped to a voltage between -120 and -20 mV for 1 second, and then peak Na⁺ current was measured during a -20-mV test potential. The prepulse duration used in these experiments was chosen on the basis of preliminary experiments demonstrating that 1 second was sufficient to allow steady state inactivation to occur in both Na⁺ channel isoforms before the test pulse. Recovery from inactivation was assessed by a double-pulse protocol consisting of a 500-millisecond prepulse to +20 mV and designed to fully inactivate both hSkM1 and hH1, followed by a variable-duration pulse to -120 mV and a test potential to -20 mV. The pulse-protocol cycle time was 30 seconds unless otherwise stated.

Data Analysis
All measurements were made with custom analysis programs designed to read and analyze binary data files. To determine the membrane potential for V_1/2 and the slope factor k, steady state inactivation data were fit with the Boltzmann equation as follows: I/I_max={1+exp[(V-V_1/2)/k]}^-1, where I is current and V is voltage. Recovery from inactivation was analyzed by fitting data with a two-exponential equation using a nonlinear least squares minimization method as follows: I(t)/I_max=A₁·exp(-t/₁)+A₂·exp(-t/₂)+A₃. After shifting the time axis so that peak current was set to t=0, the time course of current decay during a voltage step was fitted with single- or double-exponential functions as follows: I(t)=A+[A_i·exp(-t/_i)], where t=0 to 40 milliseconds. This transformation does not alter the fitted values. Results were presented as mean±SEM. Statistical comparisons were made using one-way ANOVA followed by Scheffé's test for multiple comparisons. In some cases, an unpaired Student's t test was used to evaluate the significance of the difference between means. Statistical significance was assumed at P<.05.

	Results

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Functional Differences Between hSkM1 and hH1
Heterologous expression of the recombinant hSkM1 and hH1 Na⁺ channel -subunits in Xenopus oocytes demonstrates isoform-specific differences in both kinetic and steady state inactivation properties. Oocytes expressing hH1 exhibit Na⁺ currents that inactivate rapidly, whereas hSkM1-expressing cells display currents that decay slowly (Fig 1A). In addition, recovery from inactivation is rapid in hH1 but slow in hSkM1 (Fig 1B). Finally, the membrane potential at which 50% of channels are inactivated by a 1-second prepulse (V_1/2) is significantly more hyperpolarized in hH1 compared with hSkM1 (Fig 1C). These distinct electrophysiological properties can be attributed to intrinsic structural differences between the two -subunits. When coexpressed with the human Na⁺ channel ß₁-subunit, hSkM1 exhibits an acceleration of inactivation and recovery from inactivation as well as a negative shift in V_1/2, whereas there is little or no effect on the inactivation properties of hH1 exerted by the ß₁-subunit.¹⁵ These observations indicate that intrinsic differences between -subunit structures are also responsible for the distinct ß₁-subunit effects.

View larger version (18K): [in this window] [in a new window]   Figure 1. Comparison of two-electrode voltage-clamp recordings made in Xenopus oocytes expressing either hSkM1 or hH1 Na+ channel -subunits. A, Representative current tracings recorded during a depolarizing voltage step to -20 mV from a holding potential of -120 mV. Peak current amplitudes were scaled to illustrate differences in the time course of current decay. B, Time course of recovery from inactivation determined using a double-pulse protocol as illustrated. Representative data obtained at a holding potential of -120 mV are shown. The fractional Na+ current amplitude was determined as the ratio of peak currents measured at -20 mV after a given test interval (t) to the maximum current amplitude. C, Steady state inactivation curve using the pulse protocol illustrated in the panel. Values determined for V1/2 were -54.6±1.0 mV for WT-hSkM1 and -74.6±0.9 mV for WT-hH1. Currents were normalized to values obtained with a prepulse of -120 mV. Representative data are shown.

Role of -Subunit Cytoplasmic Regions in Na⁺ Channel Inactivation
In order to identify structural determinants for the observed inactivation differences between hH1 and hSkM1, we studied chimeras consisting of defined regions donated by the two -subunit isoforms. Our initial experiments focused on amino acid sequence differences located within the ID3-4 region^{16 18} because this region is critical for fast inactivation. We also targeted the ID1-2 and ID2-3 cytoplasmic linker regions because of the sequence divergence that exists in these structures between hH1 and hSkM1. These structures were exchanged between the two -subunit isoforms as described in "Materials and Methods."

All chimeric constructs gave functional Na⁺ channels in oocytes with expression levels similar to the WT channels. Fig 2 shows representative current recordings obtained with a test pulse to -20 mV from a holding potential of -120 mV in oocytes expressing either WT or chimeric channels as illustrated. Interdomain chimeras constructed with the hSkM1 background exhibit slow macroscopic inactivation similar to the WT channel. Similarly, all hH1 chimeras exhibit the fast inactivation characteristic of the parent isoform. After making these initial observations, we considered whether combinations of these cytoplasmic regions would contribute to inactivation kinetics. Therefore, we constructed chimeric channels in which all three interdomain regions were exchanged between hH1 and hSkM1. The time course of macroscopic current decay observed for these "triple" interdomain chimeras was also similar to that of the parent channels.

View larger version (46K): [in this window] [in a new window]   Figure 2. Expression of WT and interdomain chimeric Na+ channels in oocytes. Representative current tracings were recorded during a voltage step to -20 mV from a holding potential of -120 mV. Current amplitudes are scaled and plotted on the same time axis to illustrate differences in the time course of current decay. The composition of the Na+ channel chimera is illustrated by the channel drawing located to either the right (hH1 chimeras) or left (hSkM1 chimeras) of the current tracing. In the drawings, heavy lines with filled rectangles depict segments belonging to hH1, whereas fine lines and open rectangles indicate structures from hSkM1. For simplicity, these drawings illustrate only the two domains that neighbor the chimeric segment in the single interdomain chimeras.

Functional properties of each chimera were examined in more detail by determining time constants for current decay and recovery from inactivation, as well as by analyzing steady state inactivation. Inactivation of WT-hSkM1 and the hSkM1-background chimeras was well fit with monoexponential functions. For WT-hSkM1, the average value was 9.2±0.5 milliseconds (n=9) at -20 mV, and all of the hSkM1 chimeras displayed values similar to that of the parent channel (hSkM1-A, =8.7±0.9 milliseconds [n=9]; hSkM1-B, =10.5±0.4 milliseconds [n=8]; hSkM1-C, =13.0±0.7 milliseconds [n=11]; and hSkM1-ABC, =13.0±1.0 milliseconds [n=6]). Inactivation of WT-hH1 and chimeric hH1 channels was best fit with a function containing the sum of two exponential terms. Inactivation of WT-hH1 exhibited a major fast component (_fast=1.5±0.1 milliseconds, 82±2% [n=18]) and a minor slow component (_slow=8.6±0.7 milliseconds, 18±2%). Replacement of ID1-2, ID2-3, or ID3-4 of hH1 with the corresponding region from hSkM1 produced no significant changes in the values (hH1-A, _fast=1.8±0.2 milliseconds [n=7] and _slow=15.9±3.6 milliseconds [n=7]; hH1-B, _fast=1.5±0.2 milliseconds [n=8] and _slow=13.5±4.7 milliseconds [n=8]; hH1-C, _fast=1.2±0.2 milliseconds [n=10] and _slow=18.2±6.7 milliseconds [n=10]; and hH1-ABC, _fast=1.1±0.1 milliseconds [n=4] and _slow=7.9±0.2 milliseconds [n=4]).

Recovery from inactivation was examined using a two-pulse protocol (see inset; Fig 1B) from a holding potential of -120 mV. The time course of recovery in hSkM1 consisted of a fast (_fast=5.9±1.5 milliseconds, 54±4% [n=4]) and a slow (_slow=1320±490 milliseconds, 46±4%) component. Each of the hSkM1-background chimeras except for hSkM1-C exhibited a greater proportion of the slow-recovering component. None of the hSkM1-background interdomain chimeras displayed recovery from inactivation with a time course similar to hH1. values (_fast and _slow) were roughly equivalent to WT-hSkM1 except in the triple interdomain chimera (hSkM1-ABC), which exhibited much slower recovery from inactivation (_fast=13.0±5.5 milliseconds, 27±5%; _slow=3700±500 milliseconds, 73±5% [n=6]). Although WT-hSkM1 fully recovered from inactivation within 25 seconds, hSkM1-ABC required >60 seconds for complete recovery. We confirmed this result with cRNA from three different hSkM1-ABC clones expressed in multiple batches of oocytes, and all reproducibly displayed this very slow recovery from inactivation, indicating that these observations are not due to inadvertent mutations. Because recovery from inactivation was slowed, not accelerated, in this chimera, we cannot attribute the faster recovery of hH1 to structures within any of the cytoplasmic interdomain regions.

Recovery from inactivation of WT-hH1 was resolved into two components with a predominant fast (_fast=9.3±2.4 milliseconds, 94±4% [n=6]) and a minor slow (_slow=265±117 milliseconds, 6±4%) component. All of the hH1 chimeras had time constants and component amplitudes similar to WT-hH1, demonstrating that replacement of hH1 interdomains with those of hSkM1 does not alter the recovery process. The triple hH1 chimera (hH1-ABC) did not exhibit unusual recovery kinetics in contrast to hSkM1-ABC.

Steady state inactivation was assessed by a conventional two-pulse protocol using 1-second prepulses at various membrane potentials (Fig 3). The V_1/2 was significantly shifted toward more positive potentials in hSkM1-A (+4.9 mV, P<.01 [n=4]) and hSkM1-C (+4.6 mV, P<.01 [n=4]), whereas it was shifted to more negative voltages in hSkM1-B (-4.8-mV shift, P<.01 [n=8]) and in hSkM1-ABC (-5.8-mV shift, P<.01 [n=13]). Since the V_1/2 values of hSkM1-B and hSkM1-ABC were intermediate between WT-hSkM1 and WT-hH1, these data indicate that the ID2-3 region accounts in part for the difference in steady state inactivation between the two isoforms. The steady state inactivation properties exhibited by the hH1 background chimeras were not significantly different from those of WT-hH1 (Fig 3, bottom panel). The slope factors determined for all chimeras did not differ significantly from the WT channel values.

View larger version (24K): [in this window] [in a new window]   Figure 3. Steady state inactivation curves for WT and interdomain chimeras. Representative data obtained with the pulse protocol illustrated in each panel are shown. Currents are normalized to the values obtained at a prepulse of -120 mV. Values for V1/2 are as follows (mV): hSkM1-A -49.7±0.8, hSkM1-B -59.4±0.9, hSkM1-C -49.2±1.0, hSkM1-ABC -60.2±2.1, hH1-A -76.4±0.9, hH1-B -71.8±0.8, hH1-C -73.8±0.9, and hH1-ABC -76.2±0.8.

Inactivation Properties of Split Chimeras
To further identify -subunit structures that determine the distinct inactivation characteristics of hH1 and hSkM1, we constructed chimeric channels in which approximately half of each isoform was exchanged. These split chimeras, designated as SH (amino-terminal portion of hSkM1, carboxy-terminal portion of hH1) and HS (amino-terminal portion of hH1, carboxy-terminal portion of hSkM1), were constructed such that the junction between the two sequences occurred immediately before D3. Heterologous expression of both HS and SH in oocytes resulted in robust Na⁺ currents that exhibited inactivation properties intermediate between WT-hH1 and WT-hSkM1 (Fig 4). The time constants of current decay (Fig 4A) and recovery from inactivation (Fig 4B) were intermediate between the two parent isoforms. Similarly, steady state inactivation V_1/2 values in HS and SH were intermediate between the two WT channels (HS, -67.3±0.9 mV [n=6]; SH, -64.2±0.9 mV [n=15]) (Fig 4C). Since neither HS nor SH exhibited an inactivation phenotype that closely resembled one of the parent isoforms, it is likely that inactivation differences are determined by multiple -subunit structures and are not compartmentalized to a single domain.

View larger version (24K): [in this window] [in a new window]   Figure 4. Comparison of two-electrode voltage-clamp recordings from oocytes expressing either WT Na+ channels (hSkM1 and hH1) or split chimeras (HS and SH). A, Representative current tracings recorded during a test pulse to -20 mV from a holding potential of -120 mV. Peak current amplitudes were scaled and superimposed to illustrate differences in the time course of inactivation. B, Time course of recovery from inactivation obtained using the double-pulse protocol illustrated in the panel. C, Steady state inactivation curves obtained using the pulse protocol illustrated in the panel. Current amplitudes were normalized to values obtained at the -120-mV prepulse.

Determinants of ß₁-Subunit Modulation of Inactivation
We next considered whether the observed differences in the response of hH1 and hSkM1 to a coexpressed ß₁-subunit would segregate with a particular region of the -subunit. To assess this, we coexpressed hß₁¹⁵ with each of the described chimeras. For the eight cytoplasmic interdomain chimeras, coexpression of hß₁ either accelerated (hSkM1 background chimeras) or had no discernible effect on (hH1 background chimeras) the rate of inactivation (data not shown). These results indicate that the hß₁-subunit does not require a cytoplasmic interdomain region of the -subunit for its activity. In contrast, coexpression of hß₁ with the split chimeras resulted in clear changes in inactivation properties (Fig 5). In the HS chimera, coexpression of hß₁ caused a dramatic increase in the rate of inactivation similar to that observed for WT-hSkM1. In contrast, coexpression of hß₁ with chimera SH had no discernible effect on the inactivation time course.

View larger version (29K): [in this window] [in a new window]   Figure 5. Effect of coexpressed hß1-subunit on the rate of inactivation of WT Na+ channels and split chimeras. Currents were recorded and scaled as described in the Fig 1 legend and then scaled and superimposed to illustrate differences in inactivation rate. Representative current tracings are shown for HS (left) and SH (right) in the absence and presence of hß1. Constructs are illustrated by the drawings located above each set of current recordings. Tracings for WT-hSkM1 and WT-hH1 are also shown for comparison. Coexpression of hß1 causes an acceleration of inactivation rate only for the HS chimera.

Quantitative analysis of this effect revealed that hß₁ causes a statistically significant increase in the proportion of fast inactivating current in the HS chimera (HS alone, 28.7±5.9% [n=8]; HS+hß₁, 79.7±5.5% [n=9]; P<.005), whereas its effect on SH was not significant (SH alone, 51.1±6.9% [n=9]; SH+hß₁, 65.4±4.2% [n=8]). Similarly, coexpression of hß₁ causes a significant (P<.05) increase in the proportion of HS channels that exhibit fast recovery from inactivation (HS alone, 75.0±5.7% [n=10]; HS+hß₁, 90.7±1.2% [n=9]; P<.05), but hß₁ had no significant effect on the fast recovering fraction of the SH chimera (SH alone, 73.4±4.5% [n=9]; SH+hß₁, 70.7±3.3% [n=9]). Coexpression of hß₁ did not affect the V_1/2 for steady state inactivation of either HS or SH (HS+hß₁, -64.8±0.8 mV [n=9]; SH+hß₁, -61.1±0.6 mV [n=9] [compared with values obtained in the absence of hß₁ cited above]). These observations strongly suggest that structures located in the carboxy-terminal half of the -subunit molecule are required for the modulatory effect of the ß₁-subunit on inactivation.

We attempted to further localize structures responsible for the ß₁-subunit effect by constructing and expressing additional chimeric Na⁺ channels consisting of D3, D4, or the carboxy terminal of hH1 inserted into the background of hSkM1. Expression of the carboxy-terminal hSkM1 chimera (hSkM1-CT) revealed inactivation properties and a ß₁-subunit response similar to those for WT-hSkM1 (data not shown). The D3 chimera (designated as S124H3) exhibited a time course of inactivation that was slightly slower than that for WT-hSkM1 (=14.6±0.9 milliseconds [n=8] versus 9.2±0.5 milliseconds for WT-hSkM1) (Fig 6, top left panel). In contrast, the D4 chimera (designated S123H4) exhibited a time course of inactivation that was intermediate between WT-hH1 and WT-hSkM1 (=5.2±0.2 milliseconds [n=7]) (Fig 6, top right panel) similar to chimeras HS and SH. Recovery from inactivation for S124H3 resembled WT-hSkM1, whereas S123H4 exhibited a behavior intermediate between WT-hH1 and WT-hSkM1 (Fig 6, bottom panels). Despite our expectations that one of these chimeras would have a blunted or absent ß₁-subunit response, both channels had significant acceleration of both inactivation time course and recovery from inactivation. These data demonstrate that more than one structural domain of the -subunit is involved in the response to the ß₁-subunit.

View larger version (19K): [in this window] [in a new window]   Figure 6. Expression of D3 (S124H3) and D4 (S123H4) chimeras. Top, Representative current tracings recorded from oocytes expressing either S124H3 (left) or S123H4 (right) in the presence (+) or absence (-) of hß1-subunit. Currents were recorded as described in the Fig 1 legend. Bottom, Effect of hß1 on the time course of recovery from inactivation for S124H3 (left) and S123H4 (right) chimeras. Experiments were performed as described in "Materials and Methods" using a holding potential of -120 mV. t indicates test interval.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
In the present study, we have used chimeric human Na⁺ channels to exploit the natural functional differences between heart and skeletal muscle isoforms in order to delineate structural components of the -subunit that govern its inactivation properties. Although previous work has helped identify generic structures essential for Na⁺ channel inactivation, little is known about isoform-specific differences in the kinetic and steady state attributes of this important gating phenomenon. Such differences may be clues to understanding subtle principles of structure-function relationships in voltage-gated Na⁺ channels.

Inactivation Phenotype Is Determined by Multiple -Subunit Domains
There is substantial experimental evidence indicating that the ID3-4 region of the -subunit plays a major role in Na⁺ channel inactivation. Deletions and mutations in this region of mammalian Na⁺ channel -subunit isoforms lead to removal or slowing of inactivation.^{5 7 8 22} Specific antisera directed against this substructure have also been shown to interfere with this gating process.⁶ However, the participation by other -subunit domains in inactivation is poorly defined. Evidence for a role of structures other than ID3-4 in the inactivation of human skeletal muscle Na⁺ channels has been obtained by biophysical studies of naturally occurring mutant channels found in hyperkalemic periodic paralysis and paramyotonia congenita, which are rare autosomal-dominant muscle diseases.^{19 23 24 25 26} Missense mutations in several locations of the hSkM1 -subunit, especially concentrated within D4, cause subtle yet physiologically relevant disturbances in either the kinetics or steady state properties of inactivation. Whether the affected structures have a physiological role in inactivation or the mutations perturb gating by more indirect mechanisms is not yet clear. Nevertheless, there is clearly a need for more experimentation to discern the role of structures other than ID3-4 in Na⁺ channel inactivation.

We have carried out a systematic study of the interdomain regions in the recombinant human Na⁺ channel -subunit isoforms hSkM1 and hH1 to assess the role of these structures in determining inactivation properties. Our results clearly show that amino acid differences within the ID3-4 region are not sufficient to explain the observed differences in inactivation rate, recovery rate, and steady state inactivation properties between these two channels. This observation confirms results obtained by Hartmann et al,²² who replaced the nonconserved ID3-4 residues of hH1a with the corresponding amino acids of rat brain IIA, another slowly inactivating channel isoform. We have also examined the importance of the two other cytoplasmic interdomain structures (ID1-2 and ID2-3), which differ greatly in primary sequence between hSkM1 and hH1. Our data indicate that none of these cytoplasmic domains alone or in combination is sufficient to explain the gating differences observed between WT-hH1 and WT-hSkM1.

The intermediate rates of inactivation observed for the split and D4 chimeras strongly suggest that differences in the inactivation properties between hSkM1 and hH1 are determined by multiple -subunit domains. We hypothesize that portions of the inactivation gate or its receptor are formed by structures contributed by two or more structural domains and that primary sequence differences within these structures determine the isoform-specific inactivation properties. Although these domains may not be adjacent in the primary structure, they may be contiguous in the folded channel protein in situ. Candidates for such structures include the S4-S5 loop, which has been shown to contribute to inactivation in Shaker K⁺ channels,²⁷ and the S6 segments, by analogy to work done in the L-type Ca²⁺ channel ₁-subunit.²⁸ Structures within D1 in the rat skeletal muscle Na⁺ channel -subunit have been shown to partially explain the inactivation differences between muscle and cardiac Na⁺ channels,^{29 30} but similar to our work with the split chimeras, the degree of change in the inactivation properties was not complete, suggesting a requirement for other structures.

Structures Involved in Na⁺ Channel Subunit Association
Na⁺ channels expressed in brain and skeletal muscle are heteromeric complexes of - and ß-subunits.¹ In muscle, one ß₁-subunit is noncovalently associated in a 1:1 stoichiometry with the -subunit.³¹ The presence of ß-subunits in cardiac tissue has been demonstrated by biochemical and immunochemical techniques,³² but these proteins may not be associated specifically with the cardiac -subunit isoform.³³ Additional data obtained from the functional reconstitution of recombinant ß₁-subunit pairs have demonstrated dramatic effects on brain³⁴ and skeletal muscle¹⁴ Na⁺ channels but little effect on hH1.¹⁵ We have exploited this difference in response to the ß₁-subunit to delineate structures required for this phenomenon. In our split chimeras, HS and SH, acceleration of inactivation occurred only when the carboxy-terminal half of the channel was donated by hSkM1. The effect of the ß₁-subunit on recovery kinetics also depends on this structural region of the -subunit. These observations strongly support the notion that the ß₁-subunit effect on hSkM1 is dependent on this carboxy-terminal region of the channel. We attempted to localize this effect to D3, D4, or the carboxy terminal, and our data indicate that the region responsible for the ß₁ effect is not confined to a single one of these structures.

Structural Implications
Our data demonstrate that certain functional differences between heart and skeletal muscle Na⁺ channel isoforms cannot be localized to a single cytoplasmic or intramembranous domain structure. These results have possible implications for the tertiary structure of the Na⁺ channel -subunit molecule. Either the structural differences (which are widely distributed in the -subunit) determine isoform-specific inactivation properties, or protein domains (which are separated in the primary sequence of the channel but are in close proximity in the final folded channel molecule) are responsible for the observed differences in inactivation and subunit modulation. Other functional properties of Na⁺ channels have also been shown to depend on more than one structural domain. The interaction of -scorpion toxin with rat brain Na⁺ channels can be disrupted by antibodies directed against the extracellular portion of the S5-S6 loop in both D1 and D4.³⁵ The requirements for multiple domain interactions have also been recognized to be important in determining functional properties of other proteins, such as subunit specificity in G proteins,³⁶ and in determining ligand binding selectivity in ß-adrenergic receptors.³⁷ When the complex biophysical nature of inactivation and the large size of mammalian Na⁺ channels are taken into account, it is easy to consider that differences in inactivation properties result from multiple structural differences.

	Selected Abbreviations and Acronyms

 

= time constant D (associated with number) = domain hß1 = human ß1-subunit hH1 = human heart -subunit isoform hH1-A, -B, and -C = hH1 chimeras with cytoplasmic interdomains from hSkM1 HS and SH = split chimeras with the relative position of hH1 and hSkM1 sequences noted by H and S, respectively hSkM1 = human skeletal muscle -subunit isoform hSkM1-A, -B, and -C = hSkM1 chimeras with cytoplasmic interdomains from hH1 ID (associated with number range) = interdomain PCR = polymerase chain reaction S (associated with number) = segment V1/2 = half-maximal inactivation WT = wild-type

	Acknowledgments

 
This study was supported by grants from the National Institutes of Health (HL-51197, HL-46681, and NS-32387) and the American Heart Association, Tennessee Affiliate, Inc. Dr Bennett is an Established Investigator of the American Heart Association, and Dr George is a Lucille P. Markey Scholar. The authors thank Dan Roden for his advice and critical review of the manuscript.

Received April 14, 1995; accepted October 13, 1995.

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