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The Importance of Hydrophobic Coupling. Effects of Micelle-forming Amphiphiles and Cholesterol

Jens A. Lundbæk^1,2,3, Pia Birn¹, Anker J. Hansen¹, Rikke Søgaard³, Claus Nielsen⁴, Jeffrey Girshman², Michael J. Bruno², Sonya E. Tape², Jan Egebjerg¹, Denise V. Greathouse⁵, Gwendolyn L. Mattice⁵, Roger E. Koeppe, II⁵, and Olaf S. Andersen²

¹ Novo Nordisk A/S, DK-2760, Måløv, Denmark
² Department of Physiology and Biophysics, Weill Medical College of Cornell University, New York, NY 10021
³ Institute of Biological Psychiatry, St. Hans Hospital, DK-4000 Roskilde, Denmark
⁴ Quantum Protein Center, The Technical University of Denmark, DK-2800, Lyngby, Denmark
⁵ Department of Chemistry and Biochemistry, University of Arkansas, Fayetteville, AR 72701

Address correspondence to Jens A. Lundbæk, Institute of Biological Psychiatry, St. Hans Hospital, Boserupvej 2, DK-4000 Roskilde, Denmark. Fax: (45) 46 33 43 67. email: lundbaek{at}dadlnet.dk

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Membrane proteins are regulated by the lipid bilayer composition. Specific lipid–protein interactions rarely are involved, which suggests that the regulation is due to changes in some general bilayer property (or properties). The hydrophobic coupling between a membrane-spanning protein and the surrounding bilayer means that protein conformational changes may be associated with a reversible, local bilayer deformation. Lipid bilayers are elastic bodies, and the energetic cost of the bilayer deformation contributes to the total energetic cost of the protein conformational change. The energetics and kinetics of the protein conformational changes therefore will be regulated by the bilayer elasticity, which is determined by the lipid composition. This hydrophobic coupling mechanism has been studied extensively in gramicidin channels, where the channel–bilayer hydrophobic interactions link a "conformational" change (the monomerdimer transition) to an elastic bilayer deformation. Gramicidin channels thus are regulated by the lipid bilayer elastic properties (thickness, monolayer equilibrium curvature, and compression and bending moduli). To investigate whether this hydrophobic coupling mechanism could be a general mechanism regulating membrane protein function, we examined whether voltage-dependent skeletal-muscle sodium channels, expressed in HEK293 cells, are regulated by bilayer elasticity, as monitored using gramicidin A (gA) channels. Nonphysiological amphiphiles (ß-octyl-glucoside, Genapol X-100, Triton X-100, and reduced Triton X-100) that make lipid bilayers less "stiff", as measured using gA channels, shift the voltage dependence of sodium channel inactivation toward more hyperpolarized potentials. At low amphiphile concentration, the magnitude of the shift is linearly correlated to the change in gA channel lifetime. Cholesterol-depletion, which also reduces bilayer stiffness, causes a similar shift in sodium channel inactivation. These results provide strong support for the notion that bilayer–protein hydrophobic coupling allows the bilayer elastic properties to regulate membrane protein function.

Key Words: gramicidin A • bilayer material properties • bilayer deformation energy • hydrophobic coupling • lipid–protein interactions

Abbreviations used in this paper: ßOG, ß-octyl-glucoside; CMC, critical micellar concentration; DHA, docosahexaenoic acid; DOPC, dioleoylphosphatidylcholine; gA, gramicidin A; GX100, Genapol X-100; HH, Hodgkin-Huxley; MßCD, methylated ß-cyclodextrin; TX100, Triton X-100; rTX100, reduced TX100.

¹ Though G_def⁰ is determined by K_a, K_c, c₀, d₀, and r₀, the precise value will be a function also of the constraints on lipid packing around the protein (Helfrich and Jakobsson, 1990; Nielsen and Andersen, 2000), as well as any spatial variation in the elastic moduli adjacent to the protein (Partenskii and Jordan, 2002). Further, although monolayer compression and bending are independent modes of deformation; the elastic moduli associated with these two modes of deformation are related by K_c=K_a· (Evans and Skalak, 1979), where b 24 (see Rawicz et al., 2000).

² The CMC varies as a function of salt concentration (Walter et al., 2000). The quoted values are for 0.1–0.2 M NaCl solutions.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Regulation of membrane protein function by the host bilayer lipid composition has long been enigmatic. There are numerous examples of such regulation (e.g., Bienvenüe and Marie, 1994; Dowhan, 1997; Killian, 1998; Lee, 2003), but the underlying mechanisms remain unclear, and specific lipid–protein interactions have been identified only in a few cases (e.g., Awasthi et al., 1971; Robinson, 1982; Hilgemann and Ball, 1996; Balla et al., 2000; Hilgemann et al., 2001; Hla et al., 2001). At 100 K, lipids can bind so tightly to membrane proteins that they are identified in crystal structures (Iwata et al., 1995; Luecke et al., 1999b; McAuley et al., 1999; Valiyaveetil et al., 2002). At 300 K, however, the residence time of most lipids in the annulus surrounding a protein is 10^–7 to 10^–6 s, and the specificity of lipid–protein interactions tends to be modest (Marsh and Horváth, 1998). Consistent with this modest specificity, accumulating evidence show that protein function may be regulated by the bilayer-collective properties such as bilayer thickness and monolayer equilibrium curvature (for reviews see Epand, 1997; Brown, 1997; Andersen et al., 1999; Bezrukov, 2000). This "nonspecific" regulation of protein function could result from protein–bilayer hydrophobic interactions that couple protein conformational changes to elastic deformations of the surrounding bilayer. This would provide a mechanistic basis for the regulation of protein function by the bilayer collective properties. In the present study we investigate whether voltage-dependent sodium channels expressed in HEK293 cells are regulated by such a mechanism.

The "nonspecific" bilayer regulation of protein function has been elucidated in studies on simple systems such as gramicidin and alamethicin channels. In these simple channels, it is well established that channel function is regulated by the elastic properties of the bilayer in which they are imbedded (Sawyer et al., 1989; Andersen et al., 1992; Keller et al., 1993; Lundbæk and Andersen, 1994, 1999; Lundbæk et al., 1996, 1997; Bezrukov et al., 1998). The situation is best understood in gramicidin channels. The structure of the gA channel is known (Arseniev et al., 1986; Ketchem et al., 1997; Townsley et al., 2001; Allen et al., 2003); gA channels are miniproteins formed by transmembrane dimerization of ß^6.3-helical subunits residing in each bilayer leaflet (O'Connell et al., 1990) (see Fig. 1 A), and the channel structure, specifically the helical pitch, does not vary when the bilayer thickness is varied (Katsaras et al., 1992).

View larger version (39K): [in this window] [in a new window]   FIGURE 1. Hydrophobic coupling between channel conformational changes and lipid bilayer deformations/perturbations. (A) Formation of a gA channel by the trans-bilayer association of two monomers causes a bilayer deformation that involves the compression and bending of the bilayer leaflets adjacent to the channel. These deformations incur energetic costs with energy densities of ·2 and ·2, respectively. Ka and Kc denote the bilayer compression and bending moduli, 2u0 the bilayer deformation – the difference between the actual bilayer thickness (l) and the (average) thickness of the unperturbed bilayer (d0) – and c0 the monolayer equilibrium curvature. The bilayer deformation energy (Gdef0) is obtained by integrating the energy density from the channel/bilayer boundary to infinity (Huang, 1986; Helfrich and Jakobsson, 1990; Nielsen and Andersen, 2000). (B) The hydrophobic coupling between a membrane protein and the surrounding bilayer means that a protein conformational change will be associated with a deformation of the surrounding bilayer. The total free energy change associated with the protein conformational change is the sum of the free energy change associated with the intrinsic rearrangement of the protein and the free energy change associated with the bilayer deformation.

Lipid bilayers are liquid crystals, with well-defined elastic properties, such as monolayer equilibrium curvature and compression and bending moduli (Helfrich, 1973; Evans and Hochmuth, 1978; Gruner, 1985; Evans and Needham, 1987; Rand and Parsegian, 1997). The energetic cost of a bilayer deformation therefore can be analyzed using the theory of elastic bilayer deformations (Huang, 1986; Helfrich and Jakobsson, 1990; Nielsen et al., 1998; Dan and Safran, 1998; Nielsen and Andersen, 2000; Partenskii and Jordan, 2002). In simple cases, such as the gA channel where the protein can be approximated as a cylinder, a mismatch between d₀ and l will induce a bilayer deformation that involves compression and bending of the two monolayers (Fig. 1 A). When the equilibrium monolayer curvature (c₀) is zero, the energetic cost of this deformation is equivalent to the deformation of a linear spring, and the deformation-free energy (G_def⁰) is given by, e.g., Nielsen et al. (1998):

(1)

where H_B is a phenomenological spring constant whose magnitude is determined by the bilayer elastic moduli (the compression and bending moduli, K_a and K_c), d₀ and the protein radius, r₀. 2u₀ is the hydrophobic mismatch (2u₀ = d₀ – l). In case c₀ 0, the unperturbed bilayer (where 2u₀ = 0) will possess a curvature frustration free energy that contributes to G_def⁰ (Gruner, 1985). In this case, the expression for G_def⁰ becomes (Nielsen and Andersen, 2000):

(2)

where the coefficients H_X and H_C, like H_B, are determined by d₀, K_a, K_c, and r₀ (Nielsen and Andersen, 2000).¹ Both elastic moduli, as well as c₀, are functions of the profile of intermolecular interactions among the bilayer-forming lipids (Helfrich, 1981; Seddon, 1990; Petrov, 1999). Explicit expressions have proven difficult to obtain (e.g., Helfrich, 1981), but molecular dynamics simulations predict moduli that are in reasonable agreement with experimental values (Lindahl and Edholm, 2000).

Alterations in the intermolecular interactions among the bilayer lipids, and the lipids and imbedded proteins, will express themselves in terms of altered moduli (and curvature)—or altered coefficients in Eqs. 1 and 2, i.e., as changes in bilayer elasticity. The energetic consequences of altered interactions between integral, membrane-spanning proteins and their host bilayer can be described also using other terms, such as: changes in bilayer compression (Mouritsen and Bloom, 1984) or curvature frustration (Gruner, 1985; Epand, 1998) energy, which were combined using the theory of elastic bilayer deformations (Huang, 1986); changes in acyl chain packing (Fattal and Ben-Shaul, 1993), the lateral pressure profile across the bilayer (Cantor, 1997), or lipid-packing stress (Bezrukov, 2000); and changes in bilayer-free volume (Mitchell et al., 1990; Booth et al., 2001). Though couched in different terms, all the above descriptions represent different approaches to parameterize the lateral interactions among the bilayer-forming lipids and between the lipids and imbedded membrane proteins (see Lundbæk et al., 1997; Dan and Safran, 1998; Bezrukov, 2000; Nielsen and Andersen, 2000). The advantage of using the theory of elastic bilayer deformations, as formulated in Eqs. 1 and 2, is that the coefficients H_B, H_X, and H_C can be evaluated knowing K_a, K_c, d₀, and r₀ (Nielsen and Andersen, 2000).

As G_def⁰ contributes to the energetic cost of gA channel formation, changes in bilayer elasticity are reflected in channel appearance rate and lifetime, as well as in channel activity (the number of conducting channels in the bilayer, a measure of the channel dimerization constant [Sawyer et al., 1989]). When bilayer elasticity is evaluated from changes in gA channel lifetime as a function of d₀, the results conform to the predictions of Eq. 1 and the experimentally determined H_B is in good agreement with predictions based on the theory of elastic bilayer deformations (Lundbæk and Andersen, 1999). The effects of amphiphiles on gA channel function agree with their effects on bilayer elasticity: micelle-forming amphiphiles, which decrease bilayer "stiffness", increase gA channel appearance rate and lifetime; cholesterol, which increases bilayer stiffness, has the opposite effect (Sawyer et al., 1989; Lundbæk and Andersen, 1994; Lundbæk et al., 1996, 1997). The hydrophobic coupling between a gA channel and the surrounding bilayer thus enables bilayer elasticity to regulate channel function in a well-defined manner. Moreover, it is possible to obtain direct in situ measurements of bilayer elasticity using gA channels as molecular force transducers (Andersen et al., 1999; Lundbæk and Andersen, 1999)

Are integral membrane proteins regulated by bilayer elasticity? Quite apart from the extensive literature on bilayer regulation of protein function, membrane proteins would be expected to be subject to regulation by bilayer elasticity if their function depends on conformational changes that involve the protein/bilayer boundary. There is substantial evidence that integral membrane proteins undergo such changes. Early, low resolution structures of gap junction channels (Unwin and Ennis, 1984) and the nicotinic acetylcholine receptor (Unwin et al., 1988) showed that protein function is associated with changes in quaternary structure (changes in subunit tilt within the bilayer). Similar subunit rearrangements have been proposed based on high-resolution structures of the extra-membranous domains of glutamate-activated channels (Mayer et al., 2001). High-resolution structures of several transmembrane domains also provide evidence for reorganization of the transmembrane domains of H⁺- and Ca²⁺-gated potassium channels (Perozo et al., 1999; Jiang et al., 2002), bacteriorhodopsin (Luecke et al., 1999a; Vonck, 2000), the stretch-activated MscL channel (Chang et al., 1998; Perozo et al., 2002), the sarcoplasmic Ca²⁺-ATPase (Toyoshima and Nomura, 2002), as well as members of the major facilitator superfamily of transport proteins (Abramson et al., 2003; Hirai et al., 2003; Huang et al., 2003). Chemical cross-linking and spectroscopic studies similarly provide evidence for the movement of membrane-spanning -helices relative to each other (Sakmar, 1998; Kaback et al., 2001; Sukharev et al., 2001). Moreover, conformational changes in membrane proteins modulate the bilayer structure. Rhodopsin activation, for example, changes the organization of the adjacent phospholipids (Isele et al., 2000).

As for gA channels, the hydrophobic coupling between a membrane protein and the surrounding bilayer will tend to cause the bilayer's hydrophobic core to adjust in an attempt to match the protein's hydrophobic exterior (Owicki et al., 1978; Mouritsen and Bloom, 1984)—and protein conformational changes that involve the protein/bilayer boundary will perturb the bilayer. In Fig. 1 B the protein conformational change involves a change in hydrophobic length as well as shape—and the bilayer perturbation is represented by a local bilayer thinning (and monolayer bending). As is the case for "simple" length changes, perturbations in local lipid packing due to changes in "shape" also will incur an energetic cost due to altered lipid–lipid and lipid–protein interactions (Dan and Safran, 1998). The free energy change associated with the bilayer deformation (G_def⁰) contributes to the total free energy change associated with protein conformational change (G_tot⁰). To the extent that G_def⁰ is significant—on the order of a few kJ/mol, or larger—its contribution to G_tot⁰ will be important for the distribution between protein conformational states.

It is, in this context, important that the volumetric compressibility modulus of proteins (Gekko and Noguchi, 1979) is one to two orders of magnitude higher than that of lipids (Liu and Kay, 1977), and bilayer-spanning -helices tend to be fairly rigid, compared with lipid bilayers, meaning that they will remain in an -helical conformation irrespective of any hydrophobic mismatch (Zhang et al., 1992). The bilayer-protein hydrophobic coupling therefore will have little impact on any given protein conformation; but if two protein conformations differ in free energy (G_tot⁰) by a few kJ/mole, changes in G_def⁰ will be important for protein function (Sackmann, 1984; Gruner, 1991; Andersen et al., 1992; Lundbæk and Andersen, 1994). The protein–bilayer hydrophobic coupling thus provides a mechanistic link between bilayer lipid elasticity, which varies with bilayer composition (Evans and Needham, 1987; Evans and Rawicz, 1990; Rawicz et al., 2000; Brown et al., 2002) and membrane protein function. In general, however, there is not sufficient information to evaluate a priori how changes in bilayer properties should alter G_def⁰ and protein function (Perozo et al., 2002). It thus becomes necessary to correlate changes in membrane protein function to changes in bilayer properties, using another (reference) system, and thereby determine whether there could be a causal link between changes in bilayer properties and membrane protein function.

We have previously provided qualitative support for such a link, as the effects of amphiphiles on N-type calcium channels correlate with the changes in bilayer elasticity, evaluated using gA channels (Lundbæk et al., 1996). Two amphiphiles that increase gA channel appearance rate and lifetime, ß-octyl-glucoside (ßOG) and Triton X-100 (TX100) (see Fig. 2) cause a hyperpolarizing shift in steady-state N-type calcium channel availability.

View larger version (19K): [in this window] [in a new window]   FIGURE 2. The molecules used in this study. (Top) Micelle-forming amphiphiles promote positive monolayer equilibrium curvature by increasing the average volume of the lipid head groups in a monolayer. The shading of the head group regions GX100, TX100 and rTX100 denotes the approximate region filled by the polyoxyethylene chain (estimate of radius based on Kuga, 1981). (Bottom) Cholesterol promotes negative curvature by decreasing the volume of the head group region. The stippled lines illustrate the hydrophilic-hydrophobic interface.

The present study provides a quantitative evaluation of the effects of bilayer elasticity on membrane protein function. This evaluation was done by comparing the effects of amphiphiles on the function of gA channels and skeletal-muscle voltage-dependent sodium channels, which have a simpler subunit composition and regulation than N-type calcium channels. The experimental design does not assume similarity of the conformational changes in gA and sodium channels; but if the effects of structurally different micelle-forming amphiphiles, on sodium channel function, correlate quantitatively with changes in gA lifetime—and further these effects are the opposite of those caused by cholesterol—it would provide strong support for the importance of bilayer elasticity. Given the complexity of bilayer–membrane protein interactions, we varied both the hydrophobic and the hydrophilic ends of the amphiphiles, but limited the variations to the compounds shown in Fig. 2.

We show: first, that ßOG and TX100, as in N-type calcium channels, cause a hyperpolarizing shift in steady-state channel availability, whereas the voltage-dependent activation appears to be unaltered; second, that these effects are reproduced by reduced Triton X-100 (rTX100) and Genapol X-100 (GX100) (Fig. 2), two other micelle-forming amphiphiles that increase gA channel duration; third, that cholesterol causes a depolarizing shift in channel availability; fourth, that there is a quantitative correlation between the effects on gA channels in planar bilayers and on voltage-dependent sodium channels in living cells; and fifth, that the effects of the amphiphiles on the properties of plasma membranes in living cells correlate with their effects on planar, lipid bilayers. We conclude that sodium channel function most likely is regulated by the bilayer elasticity, and that gramicidin channels provide a tool for quantitative studies of the role of bilayer elasticity (and hydrophobic coupling) for membrane protein function.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Transfection of HEK293 Cells
Full-length cDNA encoding the muscle sodium channel µ1 -subunit (Trimmer et al., 1989) was a generous gift from G. Mandel (State University of New York Stony Brook, NY). The cDNA was inserted into a pBK-CMV vector (Stratagene) and transfected into HEK293 cells using the Lipofectamine method (Life Technologies). HEK293 cells were grown on coverslips in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal calf serum (GIBCO BRL), 100 U/ml penicillin (GIBCO BRL), and 100 µg/ml streptomycin (GIBCO BRL), at 37°C ambient atmosphere with 5% CO₂. Stable clones expressing the sodium channel gene were selected using 0.5 mg/ml G418 (GIBCO BRL). In successfully transfected cells the average peak current was 1,560 ± 210 pA (mean ± SEM, n = 10). The depolarization induced membrane currents were inhibited >99% by 1 µM TTX. In nontransfected cells the average peak current was 20 ± 10 pA (mean ± SEM, n = 22), which is 1% of the peak currents in the transfected cells.

Whole-cell Voltage-clamp Experiments
Sodium channel currents were studied using whole-cell voltage clamp (Hamill et al., 1981) at room temperature (21–24°C). Patch pipettes had a tip resistance of 2–4 M. Voltage-pulse generation and data acquisition were controlled using an Axopatch 200A amplifier and pClamp 6.0 (Axon Instruments, Inc.). Linear leakage corrections were done online using a P/4 pulse protocol (Armstrong and Bezanilla, 1974) or during subsequent analysis using the leakage currents induced by a step in membrane potential from –80 to –90 mV. Unless otherwise noted, the currents were filtered at 10 kHz and sampled at 40 kHz. Pipette and membrane capacitances were electronically compensated for. Only experiments with a series resistance below 4 M (after 80% compensation) and a compensated voltage drop across the series resistance of less than 5 mV were used for analysis. The bathing solution was: 140 mM NaCl, 4 mM KCl, 2 mM CaCl₂, 2 mM MgCl₂, 10 mM HEPES, 10 mM glucose, adjusted to pH 7.4 with NaOH. The electrode solution was: 140 mM CsCl, 20 mM HEPES, 11 mM EGTA, 1 mM CaCl₂, 1.8 mM MgATP, 0.46 mM Na₃GTP, adjusted to pH 7.3 with CsOH. Test solutions of ßOG (>97% purity), GX100 (protein grade), TX100 (protein grade), and rTX100 (protein grade), all from Calbiochem, were prepared by dissolving the compounds directly into the bathing solution. Control or test solutions were applied to the cells using a fast superfusion system (Konnerth et al., 1987). The cell-membrane leak conductance in control cells (0.6 ± 0.2 nS, n = 12) was not significantly different from that in the presence of 30 µM TX100 (1.0 ± 0.4 nS, n = 6), 2.5 mM ßOG (0.3 ± 0.1 nS, n = 6), or in cholesterol-enriched (0.7 ± 0.3 nS, n = 8) or cholesterol-depleted cells (0.9 ± 0.2 nS, n = 7) (P > 0.05). "Timed control experiments", done at a time establishing the whole-cell configuration that matched that in experiments using amphiphiles, were done for all experimental protocols.

Manipulation of Cellular Cholesterol Content
Cell membrane cholesterol content was increased by exposure to cholesterol complexed with methylated ß-cyclodextrin (MßCD) (Christian et al., 1997). 5 mM MßCD–cholesterol complex (MßCD–cholesterol ratio 10:1) was prepared by adding cholesterol (Sigma-Aldrich) from a stock solution in chloroform–methanol 1:1 (vol:vol) to a glass test tube and evaporating the solvent under nitrogen. MßCD (average MW 1338; Cyclodextrin Technologies Development) dissolved in 5 ml growth medium was added to the glass tube. The solution was vortexed, sonicated for 3 min, and incubated in a rotating water bath at 37°C overnight. Prior to use, the solution was filtered through a 0.45-µm syringe filter to remove excess cholesterol crystals. The cells were exposed to this solution for 21 h. In a second group of cells, the membrane cholesterol content was decreased by exposure to 5 mM cholesterol-free MßCD for 21 h. (This incubation time was chosen for practical reasons, shorter exposures have been used previously [Christian et al., 1997].) That the effects of cholesterol-depletion, in the present study, were reversed by exposure to MßCD–cholesterol shows that they are not due to irreversible changes in cell function.) Experimental conditions in these experiments were as described above, except that MßCD without cholesterol was dissolved in growth medium. In a third group of cells, cholesterol-depletion induced by exposure to cholesterol-free MßCD for 21 h was reversed by a final 2-h exposure to 5 mM MßCD–cholesterol complex (10:1) prepared as described above, but dissolved in the bathing solution used in the patch clamp experiments.

To measure the cell-cholesterol content the growth medium was removed and cell monolayers on coverslips were washed three times with Hanks' balanced salt solution. Cell lipid was extracted with isopropanol for 30 min and cell protein in the lipid-extracted monolayer was recovered with NaOH (McCloskey et al., 1987). The cholesterol content was measured using the "cholesterol CHOD-PAP method" (Boehringer). Cell protein was measured photometrically (Pesce and Strande, 1973).

Data Analysis
All curve fitting was done using the nonlinear least-squares algorithm in Origin 7.0 (Microcal Software). The voltage dependence of channel activation and inactivation was obtained by fitting Hodgkin-Huxley (HH)-type relations (e.g., Hille, 2001) to the peak currents. The voltage dependence of activation was determined by fitting the relation:

to the results. I(V) is the peak current at the test potential V, G_max the maximal conductance, V_act the voltage of half maximal activation, S_act a slope factor, E_rev the reversal potential, and A an offset determined by the peak-to-peak noise. (This offset appears because the baseline current is determined as the mean current, whereas the fitting was to the lower envelope of the current trace, which produced a systematic offset equal to half the peak-to-peak current noise.) The voltage dependence of the steady-state inactivation was obtained by fitting a two-state Boltzmann distribution to the peak currents:

where I(V) is the peak current after a prepulse V, I(–130) the peak current at a prepulse of –130 mV, V_in the voltage of half maximal inactivation, A₁ the maximum value in the fit (A₁ 1), A₂ a constant determined by noninactivating currents and noise, and S_in a slope factor.

gA Channels in Planar Lipid Bilayers
gA channels were studied using the bilayer punch method (Andersen, 1983) at 25 ± 1°C. The bilayers were formed from a dioleoylphosphatidylcholine (DOPC)/n-decane solution (2% wt/vol). The electrolyte solution was unbuffered 1 M NaCl. gA was added to both aqueous solutions from an ethanolic stock; similarly, the micelle-forming amphiphiles were added from ethanolic stock solutions. The amount of ethanol in the chamber never exceeded 0.5% (vol:vol), which has no effect on channel properties (Sawyer et al., 1990). The current signal was recorded using a Dagan 3,900 patch-clamp amplifier (Dagan Instruments). The applied potential was ±200 mV. The current signal was filtered at 2,000 Hz, digitized and sampled at 20,000 Hz, digitally filtered at 500 Hz, and the current transitions were detected on-line (Andersen, 1983); single-channel current transition amplitudes and lifetimes were determined as described previously (Sawyer et al., 1989) with software written using AxoBasic (Axon Instruments, Inc.) or Visual Basic (Microsoft). The lifetime distributions were fitted by single exponential distributions, N(t) = N(0) exp(–t/), where N(t) is the number of channels with duration longer than t, N(0) the total number of channels, and the average channel lifetime.

gA Channels in HEK293 Cells
The effects of the amphiphiles on gA channels in the plasma membrane of native HEK293 cells were studied using whole-cell voltage clamp. The bathing and electrode solutions were the same as in the experiments with voltage-dependent sodium channels, except that 20 mM sucrose had been added to the bathing solution to prevent cell swelling. The membrane leak conductance in the native HEK293 cells (1.0 nS) was comparable to the value in transfected cells. gA from an ethanolic stock solution was added to the bathing solution in the experimental chamber to a nominal gA concentration of 10–100 µM, 10⁶-fold higher than in the planar bilayer experiments. (The ethanol concentration in the experimental chamber never exceeded 0.2%, which has no effect on the membrane conductance.) When the membrane conductance in the presence of gA had increased 10-fold relative to the control value, ßOG or TX100 was added by superfusing the cells with bathing solution, in which these compounds had been dissolved directly.

Statistical Analysis
Statistical analysis was done using SigmaStat 2.0 and either Student's t test or one way analysis of variance (ANOVA) with Dunnetts' method as post hoc test. Results are given as mean ± SEM, (n) where n is the number of cells.

Online Supplemental Material
A detailed kinetic analysis of channel gating at +20 mV is available at http://www.jgp.org/cgi/content/full/jgp.200308996/DC1.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
ßOG and TX100 Reversibly Promote Sodium Channel Inactivation
ßOG and TX100 reversibly inhibit sodium currents, and the inhibition varies as a function of the prepulse potential. Fig. 3 A shows the changes in sodium currents in response to a 25-s application of 5 mM ßOG or 30 µM TX100. After a 300-ms prepulse to either –130 or –60 mV, the cells were depolarized to +20 mV. Both ßOG and TX100 decrease the current, but the effect is larger after the –60-mV prepulse than after the –130-mV prepulse (Fig. 3 A). There is little apparent effect on the time course of activation and inactivation at +20 mV (see online supplemental material). Fig. 3 B shows the effects on the normalized peak currents. Prior to the application of ßOG or TX100, the peak currents after the –60- and –130-mV prepulse on average differed by a factor 2.

View larger version (21K): [in this window] [in a new window]   FIGURE 3. Effects of ßOG and TX100 on sodium channels. (A) Current traces showing the reversible inhibition of sodium currents by ßOG and TX100. (B) The time course of peak current inhibition during and after a 25-s application of 30 µM TX100 or 5 mM ßOG. Pulse protocol: every 5 s the cells were depolarized to +20 mV from a 300 ms prepulse to either –60 mV (•) or –130 mV (). Holding potential –80 mV. The horizontal bar denotes the time period of application. Mean ± SEM (n = 3, 4, TX100, ßOG). (C) Time course of peak current inhibition during 9-min application of 30 µM TX100 followed by a 5-min washout period (mean ± range, n = 2). Horizontal bars denote time periods of application and washout, respectively.

The decrease in the peak currents was qualitatively similar during longer applications. Fig. 3 C shows results from experiments where 30 µM TX100 was applied for 9 min. Every 10 s the cells were depolarized to +20 mV, from a –80-mV holding potential. The peak currents decreased monotonically, with a fast decay to 50% of the initial value over the first 10 s, and a slower decay to an approximately steady-state value over the next 5 min. After a 5 min washout period, the decrease in peak current amplitude could be reversed only partially (Fig. 3 C).

The effects of ßOG and TX100 on sodium channel inactivation were investigated further (Fig. 4) . In both the absence and presence of these amphiphiles, the voltage dependence of the steady-state inactivation after a 300-ms prepulse to varying potentials was well fitted by a two-state Boltzmann distribution (Fig. 4, top and bottom, shows measured and normalized peak currents, respectively.)

View larger version (23K): [in this window] [in a new window]   FIGURE 4. The voltage dependence of steady-state inactivation can be fitted by a two-state model in both the absence and the presence of the micelle-forming compounds. Peak currents (top) and normalized peak currents (bottom). Left: (•) control; () 10 µM TX100; () washout. Center: (•) control; () 2.5 mM ßOG; () washout. Right: timed control experiments. (•) control 1; () control 2; () control 3. Pulse protocol: every 5 s the cells were depolarized to +20 mV from 300-ms prepulses to potentials varying from –130 to +50 mV. Holding potential –80 mV. Results from single experiments.

View larger version (15K): [in this window] [in a new window]   FIGURE 5. Changes in channel steady-state availability induced by ßOG and TX100. 10 µM TX100 () and 2.5 mM ßOG () cause a negative shift in Vin (top) and a positive shift in Sin (bottom). (P < 0.05) For comparison (•) shows results obtained in timed control experiments. Pulse protocol as in Fig. 4. Mean ± SEM based on the changes in individual cells (n = 7, 6, 7, TX100, ßOG, control).

In addition to the current inhibition that is coupled to inactivation, both amphiphiles inhibit the sodium currents by a mechanism that does not appear to be coupled to inactivation. Steady-state inactivation, in both the absence and presence of TX100 or ßOG, is abolished after a 300-ms prepulse to –130 mV (Fig. 4); but both amphiphiles cause some current inhibition after such a prepulse (Figs. 3 and 4). We did not pursue the basis for this contribution to the current inhibition.

The effects of ßOG and TX100 are reproduced by rTX100 and GX100, two other micelle-forming amphiphiles (Fig. 2) of different structure. Both compounds promote sodium channel inactivation in a reversible manner. Fig. 6 shows the concentration dependence of the effects of ßOG, GX100, TX100, and rTX100 on V_in; results from timed control experiments are shown for comparison.

View larger version (20K): [in this window] [in a new window]   FIGURE 6. Concentration dependence of the effects of the amphiphiles on Vin. Top left: Effects of 10, 30, and 100 µM TX100. Top right: Effects of 0.3, 1, and 2.5 mM ßOG. Middle left: Effects of 10 and 30 µM rTX100. Middle right: Effects of 4.5 and 15 µM GX100. Bottom: Timed control experiments. Pulse protocol: every 5 s the cells were depolarized to +20 mV from a 20 ms prepulse to potentials varying from –130 to +50 mV. Holding potential –80 mV. Filter 10 kHz, sampling rate 20 kHz. Mean ± SEM. n = 4, 6, 4 (3, 10, 30 µM TX100); 3, 6, 7 (0.3, 1, 2.5 mM ßOG); 4, 1 (10, 30 µM rTX100); 3, 5 (4.5, 15 µM GX100); 6 (timed control).

Lack of Effects on Sodium Channel Voltage Activation
To test whether the changes in sodium channel inactivation reflect a general change in all aspects of channel function, we investigated the effects of ßOG or TX100 on the channels' voltage activation (Fig. 7) .

View larger version (17K): [in this window] [in a new window]   FIGURE 7. Peak sodium currents versus test potential in the absence and the presence of the micelle-forming amphiphiles. In all cases the peak current voltage relations were fitted with a two-state model (see MATERIALS AND METHODS). Top: (•) control; () 10 µM TX100; () washout. Middle: (•) control; () 2.5 mM ßOG; () Washout. Bottom: Timed control experiments. (•) Control 1; () control 2; () control 3. Pulse protocol: every 5 s the cells were depolarized from a holding potential of –80 mV to test potentials ranging from –90 to +50 mV. Results from single experiments.

Effects on Inactivation Are Not Due to Unspecific Membrane Damage
ßOG, GX100, TX100, and rTX100 are detergents that, at concentrations close to (and above) their CMC, may cause irreversible cell-membrane damage and membrane-protein denaturation (Weltzien, 1979). The promotion of sodium channel inactivation is not due to such extraneous, irreversible effects. First, the nominal concentrations of the amphiphiles always were well below their CMC. Second, the modulation of inactivation is reversible, at least for short applications (Figs. 3 and 6). (When the amphiphiles were applied for 5–15 min the changes in peak currents and midpoint potentials were not always completely reversed by a 5-min washout period. We did not investigate whether the apparently irreversible effects of longer applications could be reversed by even longer washouts.) Third, the membrane leak conductance is not increased by 30 µM TX100 or 2.5 mM ßOG (see MATERIALS AND METHODS). Fourth, the effects of the amphiphiles are specific in the sense that they depend on the physiological state of the channel (Fig. 3).

We also can exclude that promotion of inactivation is due to other irreversible, time-dependent processes caused by the whole-cell configuration (changes in bilayer–cytoskeleton or channel–cytoskeleton interactions or the loss of channel subunits) because: first, the changes in channel inactivation are reversible; and second, all experiments were compared with timed control experiments (which in their own right show only modest changes in V_in; Figs. 5 and 6).

Membrane Cholesterol Inhibits Sodium Channel Inactivation
TX100 and other micelle-forming amphiphiles increase bilayer elasticity, as evaluated from changes in gA channel lifetimes (Sawyer et al., 1989); cholesterol decreases bilayer elasticity as evaluated from changes in bilayer elastic moduli (Evans and Rawicz, 1990; Needham and Nunn, 1990) and gA channel lifetimes (Lundbæk et al., 1996). Thus, if the amphiphile-induced changes in sodium channel gating arise simply from changes in the bilayer elasticity, cholesterol-depletion should to a first approximation have effects similar to those observed after adding the micelle-forming amphiphiles—and cholesterol enrichment should have the opposite effects.

The cholesterol content of HEK293 cells was modified using MßCD (see MATERIALS AND METHODS). In control cells, the total cholesterol content was 25.0 ± 6.0 µg cholesterol/mg protein (n = 3), which is comparable to the value found in other cells types (Christian et al., 1997; Levitan et al., 2000). After incubation with 5 mM MßCD–cholesterol (10:1) for 21 h, the cholesterol content was increased to 53.0 ± 6.0 µg cholesterol/mg protein (n = 3). After incubation with cholesterol-free MßCD for 21 h, the cholesterol content was decreased to 7.2 ± 1.3 µg (n = 3) cholesterol/mg protein. When the cholesterol-depleted cells were incubated for 2 h with MßCD–cholesterol, the cholesterol content was increased to 41.6 ± 4.1 µg cholesterol/mg protein. (Cholesterol-depletion causes dissolution of sphingolipid- and cholesterol-enriched membrane rafts [e.g., Brown and London, 2000].) It is not known whether sodium channels reside in rafts; but, whether the channels reside in rafts or not, cholesterol-depletion will decrease the cholesterol content of the bilayer surrounding the channel.)

Fig. 8 shows the effects of changes in cell-cholesterol content on the voltage dependence of steady-state inactivation. In all cases, the voltage dependence of inactivation is well described by a two-state Boltzmann distribution (Fig. 8 A). Fig. 8 B shows the effects of modulation of cell cholesterol content on V_in and S_in. These experiments were done as a separate series, after the experiments with ßOG and TX100, and the effects of cholesterol were compared with a separate control group investigated during the same time period. Further, as each cell cannot be used as its own control, the results for the cholesterol experiments are reported as absolute values rather than relative changes.

View larger version (16K): [in this window] [in a new window]   FIGURE 8. Effect of changes in membrane cholesterol content on sodium channel steady-state availability. (A) Steady-state availability curves obtained in cells that are cholesterol-depleted or -enriched. At all cholesterol contents the voltage dependence of steady-state inactivation is fitted with a two-state model. (•) Control; () cholesterol depleted; () cholesterol enriched. Results from single experiments. (B) Effects of altering cell membrane cholesterol content on Vin (Top) and Sin (Bottom). Experimental conditions as in Fig. 4. Cholesterol-depletion changed both Vin and Sin compared with the control group (P < 0.05). Reversal of cholesterol-depletion again changed both Vin and Sin compared with the cholesterol-depleted group (P < 0.05). Mean ± SEM (n = 9, 8, 14, 6, cholesterol enriched, cholesterol-depleted, cholesterol-depletion reversed, timed control experiments).

Decreasing the cell-cholesterol content has similar effects as adding micelle-forming amphiphiles: a hyperpolarizing shift in V_in and a positive shift in S_in. Increasing the cholesterol content of cholesterol-depleted cells has the opposite effects; but the effects of cholesterol on inactivation appear to saturate when the cholesterol content is increased above the level in control cells. This latter result is not due to sequestration of the added cholesterol into a pool that does not affect the sodium channels, as cholesterol enrichment causes a shift in the voltage dependence of channel activation (see below).

Effects of Cholesterol on Sodium Channel Voltage Activation
In all experiments where the cell cholesterol content was manipulated, the peak current-voltage relations were well described by an HH description. Table II summarizes the effects of the cholesterol manipulations on V_act, S_act, and E_rev.

Kinetics of Inactivation
The effects of ßOG, TX100, and cholesterol on the time course of inactivation were investigated at –60 mV, which is close to V_in for the control situation (Figs. 4 and 8). The cells were depolarized from a holding potential of –110 to –60 mV, for 0.4 to 810 ms, and channel availability was determined using test pulses to +20 mV (Fig. 9) .

View larger version (30K): [in this window] [in a new window]   FIGURE 9. Effects of amphiphiles on the time course of inactivation at –60 mV. Every 10 s the cells were depolarized from a holding potential of –110 mV to a prepulse of –60 mV for a time interval that varied from 0.4 to 811.2 ms before the channel availability was determined at a potential of +20 mV. (A) (•) Control; () 30 µM TX100; () washout. (B) (•) Control; () 2.5 mM ßOG; () washout. (C) Timed control experiments. (•) Control 1; () control 2; () control 3. (D) (•) Control; () cholesterol depleted; () cholesterol enriched. Results are given as mean ± SEM of peak current values in individual cells. (The time to 75% channel availability determined from these mean values may differ from the mean of the time to 75% channel availability in the individual cells, which is given in the text.) (n = 6, 6, 5, 6, 7, and 7: TX100, ßOG, timed controls, cholesterol enriched, cholesterol-depleted, and timed controls for cholesterol experiments).

The effects of ßOG, TX100, and cholesterol-depletion on the return from inactivation were investigated at –80 mV. After a 300-ms depolarization to +10 mV, the cells were repolarized to –80 mV for 1–128 ms, and channel availability was determined at +10 mV (Fig. 10) .

View larger version (33K): [in this window] [in a new window]   FIGURE 10. Time course of recovery from inactivation. (A) (•) Control; () 10 µM TX100; () washout. (B) (•) Control; () 2.5 mM ßOG; () washout. (C) Timed control experiments. (•) Control 1; () control 2; () control 3. (D) (•) Control; () cholesterol depleted. Pulse protocol: every 5 s the cells were depolarized from –80 to +10 mV for 300 ms, followed by a repolarization to –80 mV for time intervals varying from 1 to 128 ms; finally the channel availability was determined at +10 mV. Mean ± SEM (n = 3, 3, 6, 6, and 5: TX100, ßOG, timed controls, cholesterol-depleted, timed controls for cholesterol experiments).

Effects on Gramicidin Channels
To determine whether the changes in sodium channel inactivation could be related to changes in bilayer elastic properties, we examined the effects of the micelle-forming amphiphiles and cholesterol on the function of gA channels in planar lipid bilayers (Fig. 11) and HEK293 cell membranes (Fig. 12) . We first studied the changes in gA channel function in dioleoylphosphatidylcholine (DOPC)/n-decane bilayers.

View larger version (28K): [in this window] [in a new window]   FIGURE 11. Effects of ßOG, GX100, TX100, rTX100, and cholesterol on gA channel activity and lifetimes. (A) Single-channel current traces recorded from the same bilayer before (top) and after (bottom) the addition of 0.3 mM ßOG. (B) Lifetime distributions (plotted as normalized survivor plots) for gA channels in DOPC/n-decane bilayers at GX100 concentrations of 0, 4.5, and 15 µM. (C) Lifetime distributions in the absence and presence of cholesterol (1:2) in the bilayer-forming solution. (D) Concentration dependence of the variation in observed with ßOG, GX100, TX100, and rTX100. The results are plotted as /cntrl, where the subscript "cntrl" denotes the control situation versus the amphiphile concentration divided by the relevant CMC: () ßOG; () GX100; (•) TX100; () rTX100. Each data point denotes results (mean ± SEM) obtained in three or more separate experiments.

View larger version (12K): [in this window] [in a new window]   FIGURE 12. Effects of 2.5 mM ßOG on the conductance of a HEK293 cell plasma membrane in the presence of gA. After establishing the whole-cell configuration, 10 µM gA was added to the extracellular solution (first arrow). When the membrane conductance in the presence of gA had reached a steady level, ßOG was added using a superfusion system as in the experiments with sodium channels (second arrow). Results from a single experiment.

All four amphiphiles increase the overall