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Journal of Physiology (2001), 535.2, pp. 371-381
© Copyright 2001 The Physiological Society
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ABSTRACT |
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INTRODUCTION |
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Voltage-gated sodium channels are responsible for the initiation and propagation of nerve, skeletal muscle and cardiac action potentials. The orchestrated activation and inactivation gating of sodium channels is vital to normal neuronal signalling, skeletal muscle contraction and normal heart rhythms. Even small syncopations from this normal gating rhythm may alter cellular excitability and whole animal physiology significantly. Several genetic sodium channel diseases have been described, with multiple mutations leading to similar disease phenotypes. Diseases such as paramyotonia congenita (PMC), hyperkalaemic periodic paralysis (HYPP) and long QT syndrome (LQTS), are examples of well studied genetic diseases of skeletal muscle and cardiac voltage-gated sodium channels (see e.g. Cannon et al. 1991; Ptacek et al. 1991, 1992; Cummins et al. 1993; Chahine et al. 1994; Bennett et al. 1995; Dumaine et al. 1996; Fan et al. 1996; Hayward et al. 1996; Ji et al. 1996; Lawrence et al. 1996; Lerche et al. 1996; Wang et al. 1996b; Nagatomo et al. 1998). These mutant channels typically show small 'persistent' sodium currents when studied in vitro. Channel inactivation is altered through mutation, resulting in a small percentage of channels residing in the active state longer than the wild-type channel. The result is a small increase in inward current, causing a slight depolarisation of the membrane. While channel function is only minimally affected, the whole animal phenotype is significantly altered. Patients present with such symptoms as muscle paralysis or cardiac arrhythmias caused by the slight redistribution of channels from one functional state to another and a slight change in resting membrane potential.
Like many other ion channels, voltage-gated sodium channels are molecularly diverse, with different channel isoforms expressed specifically by different cell types or at different times throughout development (for reviews see Kallen et al. 1993; Catterall, 1995; Jan & Jan, 1997; Marban et al. 1998; Goldin, 2001). Often several isoforms are expressed within a single cell. While most efforts have focused on the functional role of conserved structures or the impact of natural mutations on channel function, recent work has studied functional differences between isoforms of the same ion channel species. Comparisons of steady state activation and inactivation voltages, rates of fast and slow inactivation and recovery from inactivation, as well as affinities and actions of toxins and anaesthetics in neuronal, cardiac and skeletal muscle sodium channel isoforms have been described (see e.g. Chen et al. 1992; Nuss et al. 1995a,b; Rehberg et al. 1995; Chahine et al. 1996; Makita et al. 1996; Wang et al. 1996a; Bendahhou et al. 1997; Dib-Hajj et al. 1997; Wright et al. 1997; Deschenes et al. 1998; O'Leary, 1998; Bennett, 1999; O'Reilly et al. 1999; Sheets & Hanck, 1999; Villin et al. 1999; Wei et al. 1999). Generally, these studies were designed to help understand how relatively subtle functional differences between sodium channel isoforms are responsible for the observed physiology in excitable cells.
A recent paper from this laboratory (Bennett, 1999), revealed a novel functional difference between two sodium channel isoforms, adult rat skeletal muscle sodium channel (rSkM1) and human heart sodium channel 1 (hH1). While internal papain treatment removed both rSkM1 and hH1 fast inactivation, the half-activation voltage (Va) for hH1 was shifted dramatically in the hyperpolarised direction with no significant impact on Va for rSkM1. These data imply that cytoplasmic structural differences between cardiac and skeletal muscle sodium channel isoforms exist that affect channel activation differently.
In this study, attempts were made to determine which specific cytoplasmic regions of the cardiac sodium channel are responsible for this papain-induced shift in Va. Studies of the adult human skeletal muscle isoform (hSkM1) indicate that the channel behaves similarly to its rat orthologue. That is, the activation voltage of hSkM1 is unaffected by treatment with papain. Functional comparison of hH1/hSkM1 chimeras indicates that the first two cytoplasmic loops of the cardiac sodium channel joining domain I to II (loop A) and domain II to III (loop B), are both necessary and together sufficient to produce a significant papain-induced, hyperpolarising shift in the voltage at which the cardiac channel activates. In addition, activation voltages of the chimeras differ from Va for wild-type in the absence of papain, with hSkM1 loops A and B imposing a hyperpolarising effect on Va for the channel, while loops A and B of hH1 shift Va for the channel in the depolarising direction.
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METHODS |
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Construction of chimeras
The following constructs, hSkM1, hSkM1A, hSkM1B, hSkM1C, hSkM1ABC, hH1, hH1A and hH1B, were kindly provided by Dr A. L. George Jr, as previously described (Makita et al. 1996). These clones, with the exception of the two wild-types, hSkM1 and hH1, were recloned into pCDNA3.1 or pRC-CMV (Invitrogen) containing the CMV promoter. The wild-type constructs were provided in vector pRC-CMV (Invitrogen).
Not I and EcoR I restriction enzymes that cut at unique 5' and 3' sites, respectively, were used to reclone hSkM1A, hSkM1B, hSkM1C and hSkM1ABC into pcDNA3.1. For recloning of hH1B into pcDNA3.1, Hind III and Xba I unique restriction sites on the 5' and 3' end, respectively, were used.
The PinA I/BstE II fragment (3.2 kb) of hH1A-pSP64T (Makita et al. 1996) containing the A loop of hSkM1 was recloned into hH1-CMV to create hH1A-CMV. The 2.2 kb Nhe I fragment of hH1A-pSP64T was recloned into hH1B-pcDNA3.1 to construct hH1AB-pcDNA3.1. Finally, hSkM1AB-pcDNA3.1 was constructed by recloning the 2.2 kb BstE II fragment of hSkM1B-pcDNA3.1 into hSkM1A-pcDNA3.1. Each chimera was characterised using multiple restriction analyses.
CHO cell transfection and tissue culture methods
Chinese hamster ovary (CHO) cells were transfected with each vector using lipofectamine technology as previously described (Bennett, 1999). Briefly, CHO cells were passaged onto 35 mm culture plates at about 25 % confluence. Following incubation for 24 h, cells were exposed to a 1 ml Opti-MEM medium (GIBCO Life Technologies) containing 8 µl lipofectamine (GIBCO) and 1-2 µg DNA, consisting of about 12 % pGreen Lantern-1 (green fluorescent protein, GFP; GIBCO), and about 88 % sodium channel expression vector. Following a 5-24 h incubation at 37 °C with humidified 5 % CO2, the medium was exchanged for normal, non-selective CHO cell growing medium. A post-transfection incubation for 68-76 h preceded the electrophysiological recordings, selecting cells expressing GFP.
Whole cell recording of sodium currents
Transfected cells were studied using the patch clamp whole cell recording technique described previously (Bennett et al. 1997). An Axon Instruments 200B patch clamp amplifier with a CV203BU headstage (Axon Instruments) was used in combination with a Nikon TE300 inverted microscope. Pulse protocols were generated using a 200 MHz Pentium II or 800 MHz Pentium III PC computer (Dell Computers) running Pulse acquisition software (HEKA). The resultant analog signals were filtered at 5 kHz using an eight pole Bessel filter (9200 LPF, Frequency Devices, Haverhill, MA, USA) and then digitised using the ITC-16 analog-digital converter (Instrutech, Great Neck, NY, USA).
A micromanipulator (MP-285 Sutter, Novato, CA, USA) was used to place the electrode onto the cell. Electrode glass (Drummond capillary tubes) was pulled using a two step process on a Sutter (model P-87) electrode puller to a resistance of 1-2 M
measured in the salt solutions used. The external solution used was (mM): sucrose 224, NaCl 22.5, KCl 4, CaCl2 2.0, glucose 5 and Hepes 5; the internal solution used was (mM): sucrose 120, CsF 60, NaCl 32.5 and Hepes 5 (titrated with 1 M NaOH to pH 7.4 at room temperature). Papain (1 mg ml-1, Sigma) was added to the internal solution in all studies except those shown in Fig. 4. All solutions were filtered using 0.2 µm filters (Gelman, Ann Arbor, MI, USA) immediately prior to use. Although series resistance was compensated by 95-98 % for all data, the smaller current produced using the low sodium solutions further minimised any remaining error due to series resistance to < 1 mV. All data shown are recorded at least 5 min after attaining whole cell configuration to assure complete dialysis of the intracellular solution.
All data shown in Figs 1-3 were measured with papain in the internal solution. Data described in Figs 1-3 as '5 min' refer to the data recorded 5 min after attaining whole cell configuration. Thus, papain is present in the intracellular solution for 5 min. Data described in Figs 1-3 as '+ papain' refer to data recorded following saturating effects of papain. This protocol allows one to compare directly the effects of papain on sodium channel function by observing activity of the same population of channels under varying papain-induced conditions (at 5 min and following saturating papain conditions).
Pulse protocols used to determine the conductance-voltage (G-V) relationship
The cell was held at -100 mV, stepping to various depolarised potentials (ranging from -100 to +70 mV in 10 mV increments) for 10 ms, and then returning to the holding potential. Consecutive pulses were stepped every 1.5 s and the data were leak subtracted using the P/4 method, stepping negatively from the -100 mV holding potential. At each test potential, steady-state whole cell conductance was determined by measuring the peak current at that potential and dividing by the driving force (i.e. the difference between the membrane potential and the observed reversal potential). The maximum conductance generated by each cell was used to normalise the data for each cell to its maximum conductance by fitting the data to a single Boltzmann distribution (eqn (1)). The average values of Va (± S.E.M.) throughout are determined from these single Boltzmann distributions. The normalised data were then averaged with those from other cells, and the resultant average conductance-voltage curve was fitted via least squares using the following Boltzmann relation:
Fraction of maximal conductance = [1 + exp(V - Va/Ka)]-1, (1)
where V is the membrane potential, Va is the voltage of half-activation, and Ka is the slope factor.
Data analysis
The data were analysed using a combination of Pulse/PulseFit (HEKA, Lambrecht/Pfalz, Germany) and SigmaPlot 2000 (SSPS Inc., Chicago, IL, USA) software.
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RESULTS |
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Papain removes fast inactivation from human skeletal muscle (hSkM1) and cardiac (hH1) sodium channels - only hH1 activation voltage is altered
Figure 1A and B compares whole cell current traces from a CHO cell expressing hSkM1 and hH1, respectively, at 5 min and following saturating internal papain treatment. Internal treatment with papain removes fast inactivation for both channel types after 15-25 min. After just 5 min of treatment with papain, 93.3 ± 1.9 and 93.5 ± 2.4 % (hSkM1 and hH1, respectively; n = 7 for both isoforms) of the current still inactivates at a -20 mV test pulse. As is often observed with internal protease treatment, there was a general trend towards lower peak current following treatment with papain that was equivalent for the two isoforms. The peak conductance following saturating papain treatment ('+ papain') was 70.0 ± 9.4 and 71.1 ± 8.8 % of the initial peak conductance ('5 min') for hSkM1 and hH1, respectively (n = 7 for each isoform).
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Figure 1. Activation voltage for hH1 shifts in the hyperpolarised direction following treatment with papain - hSkM1 Va is unaffected Whole cell current trace at 5 min (5 minutes) and following saturating papain treatment (+ papain) at a test potential of +30 mV for hSkM1 (A) and hH1 (B). Conductance-voltage (G-V) relationship for hSkM1 (C) and for hH1 (D) at 5 min and following saturating papain treatment. Data points are the mean ± S.E.M. conductance at each membrane potential. Curves are fits of the data to single Boltzmann distributions (eqn (1) in Methods). | ||
Figure 1C and D shows the average conductance-voltage (G-V) relationships for hSkM1 and hH1. All data shown on a single G-V graph in Figs 1-3 were measured from the same set of cells at 5 min and following saturating papain treatment. The G-V relationship for hSkM1is not significantly altered by treatment with papain, while the G-V curve for hH1 following treatment with papain is shifted dramatically in the hyperpolarised direction. The voltage of half-activation (Va) for hH1 is nearly 21 mV more hyperpolarised following treatment with papain.
This shift in activation voltage for hH1is not due to the uncoupling of inactivation from activation. If the shift in activation voltage were due to uncoupling, then any method of removing (or preventing) fast inactivation would show such a shift in Va. As shown in a previous study from this laboratory (Bennett, 1999), the inactivation deficient mutant, hH1Q3, had the same Va as wild-type hH1. In addition, treatment of hH1Q3 with papain caused a hyperpolarising shift in Va of > 18 mV, nearly identical to that observed for hH1 as shown in Fig. 1D. Therefore papain must have at least a secondary effect on hH1 that is not observed for hSkM1 - altering the voltage at which the cardiac channel activates in a manner that is not directly related to the uncoupling of inactivation from activation. Also, as shown for hH1 in Bennett (1999), there was no time-dependent drift in activation voltage for any constructs tested here (data not shown). Thus, the two sodium channel isoforms are affected differently by internal papain treatment, with hH1 being 'papain sensitive', observed as a hyperpolarising shift in Va of > 20 mV, while hSkM1 is 'papain insensitive', showing no significant shift in Va following treatment with papain.
Activation voltages of channels that contain hH1 cytoplasmic loops A and B are papain sensitive
To locate the hH1 region(s) responsible for this hyperpolarising shift in Va following treatment with papain, chimeras were constructed in an attempt to convert a papain-insensitive hSkM1 into a papain-sensitive channel. Later, we will describe the converse set of mutants, producing a papain-insensitive channel from the papain-sensitive hH1. Because internal papain is used to observe this phenomenon, hH1 cytoplasmic regions are likely to be involved.
Firstly, a relatively large section of hSkM1 was replaced by the analogous hH1 fragment, creating a papain-sensitive channel. Figure 2A shows the G-V relationships for hSkM1 and the chimera, hSkM1ABC, in which all three hH1 cytoplasmic loops joining the four transmembrane domains replaced the analogous hSkM1 loops. A schematic diagram of the corresponding channel is inset above each G-V graph here and throughout this report. The data from each graph compare steady state channel activation at 5 min and following saturation of the papain effect for a single population of cells, and show that the replacement of these three loops is sufficient to produce a papain-induced hyperpolarising shift in activation voltage. The Va for hSkM1ABC shifts by about -22 mV following treatment with papain, nearly identical to the shift of -21 mV observed for wild-type hH1. Steady state activation voltages measured at 5 min and following saturating papain treatment and the resulting shifts in Va with papain treatment for all tested constructs are listed in Table 1.
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Figure 2. The activation voltages of hSkM1 chimeras containing loops A and B of hH1 shift in the hyperpolarised direction following treatment with papain G-V relationships for wild-type hSkM1 and for the tested hSkM1 chimeras at 5 min ( | ||
Figure 2A indicates that loops A, B, and C of hH1 together are sufficient to produce a papain-induced hyperpolarising shift in Va. Next, an attempt to further limit the possible region(s) of hH1 responsible for this shift was made by testing individual loop chimeras in which one loop of hH1 replaced the analogous loop of hSkM1. The conductance-voltage relationships for hSkM1A, hSkM1B and hSkM1C are shown in Fig. 2B. Note that, like wild-type hSkM1, Va for hSkM1C does not shift significantly following treatment with papain, indicating that loop C is not involved in this phenomenon. However, the Va of hSkM1A and hSkM1B each shifts by about 15 mV in the hyperpolarised direction following treatment with papain, consistent with the suggestion that loops A and B both contribute to the papain-induced hyperpolarising shift in activation voltage.
To test this, a double loop chimera was constructed in which loops A and B of hH1 replaced loops A and B of hSkM1 (hSkM1AB; Fig. 2C). Note that the G-V relationship for hSkM1AB, like hH1, shifts sharply in the negative direction following treatment with papain. The data indicate that loops A and B of hH1 are both necessary, and together are sufficient, to produce a large papain-induced hyperpolarising shift in channel activation voltage.
Converse chimeras confirm that loops A and B of hH1 are both necessary and together sufficient to produce a papain-induced hyperpolarising shift in activation voltage
Figure 3A compares the G-V relationships of the converse chimera, hH1AB (in which loops A and B of hSkM1 replace the analogous hH1 loops) with the G-V relationships of the wild-type hH1, each at 5 min and following saturating treatment with papain. Note that hH1AB shows no shift in Va following treatment with papain, much like wild-type hSkM1. These data further verify that loops A and B of hH1 together are sufficient to confer papain sensitivity on a channel. When both hH1 loops are removed, the channel is papain insensitive.
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Figure 3. Both loops A and B of hSkM1 are necessary and together sufficient to prevent a papain-induced hyperpolarising shift in activation voltage G-V relationships for wild-type hH1 and for the tested hH1 chimeras at 5 min ( | ||
In order to determine whether both hSkM1 loops are necessary to remove papain sensitivity, the converse single loop chimeras, hH1A and hH1B, were studied. Figure 3B compares the G-V relationships of hH1A and hH1B at 5 min and following saturating treatment with papain, showing that the Va shifts in the negative direction for both constructs following treatment with papain; however, while Va for hH1 shifted by -21 mV, the Va of each chimera shifted by a significantly smaller -12 mV for hH1A, and -15 mV for hH1B. Together with the data shown in Fig. 3A, these data indicate that both loops A and B of hH1 must be removed (and replaced by analogous hSkM1 loops) in order to prevent a papain-induced shift in the voltage dependence of channel activation.
The first two large cytoplasmic loops of each isoform impose a direct and isoform-specific effect on channel activation voltage
All data presented thus far were collected in the presence of internal papain (at 5 min and following saturating papain treatment). The data indicate that disruption of the first two large cytoplasmic loops of the cardiac sodium channel causes a dramatic hyperpolarising shift in the activation voltage of the channel. As will be discussed later, there may be a physiological/pathological role for cytoplasmic proteases in the regulation of cardiac sodium channel function through this protease-induced hyperpolarisation of the cardiac channel activation voltage.
We next sought to determine whether these two loops have some direct effect on the voltage at which a sodium channel activates. Thus, the activation voltages of the wild-type and double loop chimeras in the absence of papain were compared. Figure 4 shows the G-V relationships for hSkM1 and hSkM1AB (Fig. 4A), and for hH1 and hH1AB (Fig. 4B) in the absence of papain. These data clearly indicate that loops A and B together directly alter the voltage dependence of channel activation. hH1 loops added to hSkM1 (hSkM1AB) cause a nearly 20 mV depolarising shift in activation voltage, and consistently but conversely, replacement of hH1 loops with hSkM1 loops (hH1AB) produces nearly a 9 mV hyperpolarising shift in activation voltage (see Tables 2 and 3). In addition, the activation voltage for each single loop chimera (hSkM1A, hSkM1B, hH1A and hH1B) measured in the absence of papain was shifted from the Va for the wild-type channel by a smaller, but significant 5-7 mV (data not shown; n = 7-9 for each chimera). These data show that inclusion of either loop A or B of hSkM1 produces a significant hyperpolarising shift in Va while the inclusion of either loops A or B of hH1 produces a significant depolarising shift in Va.
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Figure 4. Channel activation voltage is altered directly and in an isoform-dependent manner by loops A and B G-V relationships for the two wild-type and two double loop chimeras measured in the absence of papain. Data points are the mean ± S.E.M. conductance at each membrane potential. Curves are fits of the data to single Boltzmann distributions. A, hSkM1 ( | ||
These data indicate that loops A and B of each isoform impose a direct effect on the voltage at which a channel activates. The effect of the two cytoplasmic loops on Va is isoform specific, with loops A and B of hH1 causing a depolarising shift in Va, while loops A and B of hSkM1 cause a smaller, but significant, hyperpolarising shift in Va.
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DISCUSSION |
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The activation mechanisms of the human skeletal muscle and cardiac sodium channels differ
Internal papain treatment of hH1 and hSkM1 (Fig. 1) removed fast inactivation from each channel type, but caused a 21 mV hyperpolarising shift in Va of hH1 only. Previous work showed that this shift in Va cannot be caused by the uncoupling of inactivation from activation (Bennett, 1999). Papain must produce at least a secondary effect on hH1 that is not observed for hSkM1. Minimally one can conclude that hH1 and hSkM1 activation mechanisms are not identical.
Disruption of two cytoplasmic loops of the cardiac sodium channel are necessary and together sufficient to produce a hyperpolarising shift in the voltage of channel activation
An inclusive and converse set of chimeras was studied to determine which region(s) of the cardiac sodium channel is involved in this papain-induced hyperpolarising shift in activation voltage. The data shown in Fig. 2 indicate that loops A and B of hH1 each contributes to this phenomenon. The Va of individual loop chimeras, hSkM1A and hSkM1B, each shifts by about -15 mV following treatment with papain. Further, the double loop chimera, hSkM1AB, produced a hyperpolarising shift in Va of > 25 mV following treatment with papain.
These data are verified further through converse mutagenesis as shown in Fig. 3, in which hSkM1 loops replaced hH1 loops. The data are completely consistent with the data of Fig. 2. The Va of the double loop chimera, hH1AB, is unaffected by papain, while the Va of individual loop chimeras, hH1A and hH1B, each shifts significantly in the negative direction following treatment with papain, but by less than wild-type hH1. The papain sensitivity of hH1A and hH1B presumably is caused by the remaining papain-sensitive hH1 loop in each of these constructs.
Together, the data show that the first two cytoplasmic loops of the cardiac sodium channel that join domain I to II, and domain II to III, are both necessary, and together sufficient to produce a papain-induced hyperpolarising shift in the voltage at which channels activate.
As noted in Tables 2 and 3, the activation voltages of the two papain-insensitive constructs (hSkM1 and hH1AB) measured at 5 min with papain present (
, Fig. 2A and Fig. 3A) are not significantly different from the values ofVa measured in the absence of papain (Fig. 4). This verifies that papain has no significant effect on the activation voltage of these channels. However, the values ofVa of the papain-sensitive constructs, hSkM1AB and hH1, measured at 5 min with papain present (, Fig. 2C and Fig. 3A) are significantly more hyperpolarised than the values ofVa measured when papain is absent (Fig. 4). These data indicate that after just 5 min, papain may already disrupt some loops, resulting in a partial shift in Va of the channel. If so, then one might infer that papain induces a significantly larger hyperpolarising shift in Va than those reported in Table 1 and shown in Figs 1-3.
The data described above indicate that papain may disrupt some A and/or B loops within the first 5 min of application. Yet very few if any C loops are disrupted at the 5 min measurement since nearly all fast inactivation is still intact (> 93 % intact). This indicates that under identical conditions of treatment with papain, disruption of loops A and B of hH1 leading to a hyperpolarising shift in Va may occur more rapidly than does disruption of loop C resulting in the removal of fast inactivation.
The first two large cytoplasmic loops directly alter the voltage dependence of channel activation in an isoform-specific manner
As shown in Fig. 4 and in Tables 2 and 3, the Va of hSkM1AB measured in the absence of papain was more depolarised than the Va of wild-type hSkM1, while the Va of hH1AB was more hyperpolarised than the Va of wild-type hH1. In the absence of papain, the Va of each single loop was also consistently and significantly shifted from wild-type Va, albeit to a lesser extent. Together, the data indicate that if a channel contains intact loops A or B of hH1, the channel activates at more depolarised potentials and conversely, if a channel contains intact loops A or B of hSkM1, the channel activates at more hyperpolarised potentials. If both loops A and B are intact, then activation voltage shifts more than and in the same direction as that observed for the single loop chimeras. These data provide the first evidence of isoform-specific cytoplasmic regions of voltage-gated sodium channels that directly and differently alter the voltage of channel activation. The physiological impact of these isoform-specific differences on channel function is obvious.
A second physiological/pathological effect - does endogenous protease activity affect cardiac sodium channel function?
As discussed above, the cytoplasmic loops of two voltage-gated sodium channel isoforms directly alter channel activation voltage. In addition to this, Va of hH1 is altered through disruption of two cytoplasmic loops by papain treatment. Papain, a cysteine protease, was used as a means to uncover and quantify this phenomenon. Substantial work has shown that such papain-like proteases, in particular calpains and caspases, are active in the cytoplasm under physiological and pathological conditions. Normal cellular function appears to rely on the balance between these proteases and their inhibitors (for review, see Croall & Demartino, 1991; Saido et al. 1994; Chapman et al. 1997; James, 1998; Kidd, 1998; Nunez et al. 1998). Also, protease activity has been implicated in pathological conditions, and is specifically involved in the apoptotic pathway of many cells, including cardiac myocytes (James, 1998). This suggests that the balance between protease and inhibitor activity shifts towards increased protease activity as the cell enters the apoptotic pathway, leading to cell death. In addition, cysteine proteases were shown to cleave transmembrane proteins such as calcium ATPases and integrins, and protease activity is implicated in long term potentiation (Saido et al. 1994; Du et al. 1995).
If cardiac myocytes rely on a balance between papain family proteases and their inhibitors expressed in the cytoplasm and near the inner plasma membrane regions of the cell, then a physiological role for low basal protease activity on cardiac sodium channel function can be described. It is intriguing to suggest that physiological levels of the protease would cause a small percentage of channels to be non-inactivating, and would also cause a shift in the activation voltage of this small percentage of channels. Both phenomena would lead acutely to greater sodium current at small depolarisations, leading to larger calcium fluxes, and potentially stronger contractility. However, as the equilibrium between protease and inhibitor activity shifts toward greater protease activity, depolarisation of the resting membrane potential of the myocyte will develop due to an increased persistent sodium current, potentially leading to pathological conditions, including arrhythmias. Consistent with this is the observation that during the early stages of cardiac myocyte apoptosis, there is enhanced proteolytic activity and increased excitability, perhaps due to an increased sodium conductance (James, 1998).
Thus, endogenous protease activity in the heart may be a means by which sodium channel activity is modulated acutely. Under physiological conditions, small changes in basal protease activity may slightly alter the levels of persistent sodium current as well as readjust channel activation voltage. Under pathological conditions, protease activity apparently increases dramatically, perhaps removing all inactivation as well as shifting severely the activation voltage of the channels, increasing ionic conductances to toxic levels.
Summary
Here we have shown that the activation mechanisms of two human voltage-gated sodium channel isoforms, hH1 and hSkM1, are different. The large cytoplasmic loops of hH1 and of hSkM1 that join domain I to II (loop A) and domain II to III (loop B) apparently alter the activation voltage of the sodium channel directly and in opposing directions. These data are the first to indicate that differences in cytoplasmic structures between voltage-gated sodium channel isoforms directly and differently alter the channel activation mechanism. Experiments are continuing in an attempt to describe the mechanism by which these loops affect the voltage dependence of gating. In addition, disruption of these cardiac sodium channel loops through protease treatment causes a large hyperpolarising shift in Va. The role of these loops and proteases in cardiac myocyte function is under investigation in order to find out whether cardiac myocyte excitability is affected by the balance (or change in this equilibrium) between disrupted and intact cytoplasmic loops of the cardiac sodium channel.
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Acknowledgements
I am indebted and grateful to Dr A. L. George Jr for his generosity in providing many of the sodium channel constructs used throughout this study. His contribution proved invaluable to the successful completion of this work. Also, many thanks to Drs Jahanshah Amin and E. Truitt Sutton for their suggestions, and to Jeanie Harper for her technical help. This work was supported in part by NSF IBN-9816685, NIH AR45169-01A1, and a Grant-In-Aid from the American Heart Association, Florida Affiliate.
Correspondence
E. Bennett: Department of Physiology and Biophysics, MDC 8, University of South Florida, College of Medicine, 12901 Bruce B. Downs Blvd, Tampa, FL 33612, USA.
Email: esbennet{at}hsc.usf.edu
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, data from the same set of cells recorded following saturating papain treatment (+ papain). n = 7 for both groups.