Can gramicidin ion channel affect the dipole potential of neighboring phospholipid headgroups?

BIOJEC-06867; No of Pages 10 Bioelectrochemistry xxx (2015) xxx–xxx

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Can gramicidin ion channel affect the dipole potential of neighboring phospholipid headgroups? Lucia Becucci ⁎, Rolando Guidelli 1 Department of Chemistry “Ugo Schiff”, Florence University, Via della Lastruccia 3, Sesto Fiorentino, Firenze, Italy

a r t i c l e

i n f o

Article history: Received 11 March 2015 Received in revised form 14 May 2015 Accepted 21 June 2015 Available online xxxx Keywords: Tethered bilayer lipid membrane Surface dipole potential Cyclic voltammetry Dioleoylphosphatidylcholine Dioleoylphosphatidylserine

a b s t r a c t The cyclic voltammetry behavior of a mercury-supported tethered bilayer lipid membrane (tBLM) incorporating gramicidin A was investigated in aqueous 0.1 M KCl at pH 6.8, 5.4 and 3. The distal leaflet of the lipid bilayer consisted of dioleoylphosphatidylcholine (DOPC), dioleoylphosphatidylserine (DOPS), dioleoylphosphatidic acid or a DOPC/cholesterol mixture. In passing from pH 6.8 to pH 3, the midpoint potential between the negative current peak, due to K+ inflow into the spacer, and the positive current peak, due to K+ ejection into the aqueous solution, shifts toward more positive potentials, while the separation between these two peaks decreases. This behavior is interpreted quantitatively on the basis of a model relying on tBLM structural features estimated independently in previous works. The only adjustable parameter is the rate constant for cation translocation across a potential energy barrier located in the hydrocarbon tail region. The behavior is ascribed to a dragging of the lipid headgroups adjacent to the gramicidin channel mouth toward the hydrocarbon tail region, with a resulting decrease in the negative charge of the DOPC phosphate group, or of the DOPS carboxyl group, with decreasing pH. © 2015 Elsevier B.V. All rights reserved.

1. Introduction The intramembrane potential ϕm of a bilayer lipid membrane (BLM), albeit mainly localized in the hydrocarbon tail region, is determined by the charge density and surface dipole potential on the two polar head regions, by the adjacent diffuse layers and by the external applied potential, i.e., the potential difference, Δϕ, between the aqueous phases that bath the two sides of the membrane, often referred to as the transmembrane potential [1]. A symmetrical BLM interposed between two identical aqueous solutions at Δϕ = 0 has a zero intramembrane potential, since charge and surface dipole potential on one side of the hydrocarbon tail region are perfectly counterbalanced by those on the opposite side. The membrane surface dipole potential has been extensively investigated in lipid monolayers by the vibrating plate (Kelvin) or the ionizing electrode method and in lipid bilayers by voltagesensitive fluorescent dyes (for a recent review, see Ref. [2]). More recently, a cryoelectron microscopy (cryo-EM) method using electrons as probes of the surface dipole potential has also been devised. The surface dipole potential affects the structure and function of membraneincorporated proteins [2]. Thus, e.g., some peptides induce a decrease in the surface dipole potential of membranes [3,4]. The pentadecapeptide antibiotic gramicidin A represents an interesting model for studying the interactions of ion channels with the

⁎ Corresponding author. E-mail address: lucia.becucci@unifi.it (L. Becucci). 1 Retired professor from Florence University.

membrane environment and membrane-solution interfaces. The addition of phloretin, known to lower the surface dipole potential, decreases the dissociation constant of the gramicidin dimeric ion channel in a diphytanoylphosphatidylcholine BLM, whereas 6-ketocholestanol, known to raise the surface dipole potential, increases it [5]. By so doing, phloretin addition increases the single-channel conductance of gramicidin A for potassium ions, whereas 6-ketocholestanol decreases it. Conversely, the above two compounds exert on opposite effect on proton conductance across the gramicidin channel. This anomalous proton conductance was explained by means of a Grotthuss mechanism by Phillips et al. [6], and by a transmembrane movement of negative ionic defects along the water wire inside the ion channel by Rokitskaya et al. [7]. Gramicidin itself was found to be as potent as phloretin in lowering the surface dipole potential of dipalmitoylphosphatidylcholine, both in monolayers in the liquid condensed phase and in unilamellar vesicles [8]. This behavior was explained by a direct interaction of gramicidin with the lipid molecules, such as to alter their conformation by inducing reorientation of dipole carrying groups, e.g., phosphocholine groups and/or water molecules forming the hydration shell of lipid headgroups. This work aims at estimating the changes in the surface dipole potential of the headgroups of dioleoylphosphatidylcholine (DOPC), dioleoylphosphatidylserine (DOPS), dioleoylphosphatidic acid (DOPA) and of a DOPC/cholesterol (Chol) mixture of molar composition (70:30) at a mercury-supported biomimetic membrane incorporating gramicidin A, following pH changes in a 0.1 M KCl aqueous solution bathing the membrane. These pH changes are also shown to induce an appreciable change in the applied potential E at which the intramembrane potential ϕm equals zero. The biomimetic membrane

http://dx.doi.org/10.1016/j.bioelechem.2015.06.008 1567-5394/© 2015 Elsevier B.V. All rights reserved.

Please cite this article as: L. Becucci, R. Guidelli, Can gramicidin ion channel affect the dipole potential of neighboring phospholipid headgroups?, Bioelectrochemistry (2015), http://dx.doi.org/10.1016/j.bioelechem.2015.06.008

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L. Becucci, R. Guidelli / Bioelectrochemistry xxx (2015) xxx–xxx

employed for this purpose consists of a lipid bilayer interposed between a hydrophilic chain tethered to mercury, called “spacer”, and the bulk aqueous solution. This mercury-supported tethered bilayer lipid membrane (tBLM) is obtained by anchoring a thiolipid monolayer to a hanging mercury drop electrode (HMDE), and by then self-assembling a phospholipid monolayer on top of it. The thiolipid, called DPTL [9], consists of a tetraethyleneoxy hydrophilic chain terminated at one end with a lipoic acid residue for anchoring to the metal surface, and covalently linked at the other end to two phytanyl chains mimicking the hydrocarbon tails of a lipid. The proximal leaflet of the lipid bilayer moiety of the tBLM consists of phytanyl chains bound by ether linkages to the tetraethyleneoxy spacer. Conversely, the distal leaflet consists of any phospholipid monolayer that can be self-assembled on top of the DPTL thiolipid. In the present work, DOPC, DOPS, DOPA and DOPC/Chol (70:30) distal monolayers were employed. The tetraethylenoxy spacer may accommodate up to two potassium ions per DPTL molecule, corresponding to a charge density of about 45 μC cm−2 [10]. Mercury has several advantages over solid supports, in that it provides a fluid, readily renewable and defect-free surface to the tBLM. Thus, it imparts to the lipid molecules of the whole mixed bilayer fluidity and lateral mobility comparable with those of biomembranes. A mercury-supported tBLM reacts to the presence of proteins, charges and physical forces in a dynamic and responsive manner, by reorganizing upon interaction with external perturbations and mimicking the functionality of living cell membranes. This tBLM has been extensively employed in our laboratory for the investigation of channel-forming peptides and proteins [11,12] and its structural features have been thoroughly investigated [10,13,14]. 2. Material and methods Merck (Darmstadt, Germany) suprapur® KCl was baked at 500 °C before use to remove any organic impurities. Analytical Reagent HCl and K2HPO4 from Merck were used without further purification. Gramicidin A and Chol were purchased from Sigma Aldrich (Seelze, Germany) and used without further purification. DOPC, DOPS and DOPA were purchased in chloroform solution from Avanti Polar Lipids (Birmingham, AL, USA). The 2,3,di-O-phytanyl-sn-glycerol-1tetraethylene-glycol-D,L-α lipoic acid ester thiolipid (DPTL) was provided by Prof. Adrian Schwan (Department of Chemistry, University of Guelph, Canada) [15]; stock solutions of this thiolipid were stored at − 18 °C. Deionized water was distilled once and then redistilled from alkaline permanganate before its use. All measurements were carried out with a home-made HMDE described elsewhere [16]. A home-made glass capillary with a finely tapered tip (about 1 mm in outer diameter) was used. Capillary and mercury reservoir were thermostated at 25 ± 0.1 °C in a water-jacketed box to avoid any changes in drop area due to a change in temperature. The HMDE acted as the working electrode in a three-electrode system, with an Ag/AgCl/0.1 M KCl reference electrode and a platinum coil counter electrode. Mercury-supported tBLMs were obtained by tethering a monolayer of the DPTL thiolipid to the HMDE, upon keeping the mercury drop immersed in a 0.2 mg/mL DPTL solution in ethanol for about 20 min [17]. A phospholipid monolayer was then formed by spreading a 1.3 × 10−3 M solution of the lipid in pentane on the surface of the working aqueous solution, in an amount corresponding to about five lipid monolayers, as estimated by ascribing a cross-sectional area of 65 Å2 to the lipid molecule. This lipid film can be regarded as a monolayer at its equilibrium spreading pressure, i.e., about 50 mN m−1 [18], plus a lipid excess in equilibrium with the monolayer. After allowing the pentane to evaporate, the DPTL-coated mercury drop was immersed into the aqueous solution across the lipid film. This procedure determines the self-assembly of a lipid monolayer on top of the DPTL monolayer, thanks to the hydrophobic interactions between the alkyl chains of the phospholipid and those of the thiolipid. The tBLM was then subjected to repeated potential scans over a potential range from − 0.20 to −1.20 V, while continuously monitoring the curve of the quadrature

component of the current at 75 Hz against the applied potential, E, using AC voltammetry, until a stable curve was attained. Only well-behaved tBLMs, with a capacitance of about −1 μF cm−2 throughout the whole potential range, were employed for further measurements. Gramicidin was incorporated in the tBLM by simply adding its stock solution to the electrolysis cell in an amount corresponding to 0.1 μM. The solution was then stirred for a few minutes, while keeping the electrode at an applied potential of −0.50 V. Impedance spectroscopy, potential-step chronocoulometry and cyclic voltammetry measurements were carried out with an Autolab instrument PGSTAT12 (Echo Chemie, Utrecht, The Netherlands) supplied with FRA2 module for impedance measurements, SCAN-GEN scan generator and GPES 4.9007 software. Potentials were measured vs. a Ag/AgCl electrode immersed in the 0.1 M KCl working solution, and are referred to this reference electrode. 3. Results Fig. 1 shows stabilized cyclic voltammograms (CVs) at a DPTL/DOPC tBLM in an aqueous solution of 0.1 M KCl and 0.1 μM gramicidin, at a scan rate of 50 mV/s and at pH 3, 5.4 and 6.8. The solution at pH 3 was obtained by adding 1 mM HCl, that at pH 5.4 was unbuffered, and that at pH 6.8 was buffered with a HCl–K2HPO4 mixture. These CVs were also obtained at the same tBLM, starting from a 0.1 M KCl unbuffered solution, adding 1 mM HCl and then the appropriate amount of K2HPO4; pH changes at the same tBLM were perfectly reversible, as verified by acidifying a pH 6.8 phosphate buffered solution with HCl to bring it to pH 5.4. The CVs exhibit a negative peak, due to the flow of K+ ions into the hydrophilic spacer, and a corresponding positive peak, due to their ejection from the spacer into the aqueous solution bathing the tBLM. The two peaks are roughly centrosymmetric with respect to the midpoint potential, E1/2, between them, namely the potential at which K+ inflow matches its outflow. Integrating any of the two current peaks yields a charge density of about 45–50 μC cm− 2, which corresponds to saturation of the spacer by K+ ions [10,11]. It is apparent that E1/2 shifts toward less negative potentials with decreasing pH, passing from −0.66 V at pH 6.8 to −0.55 V at pH 5.4 and to −0.44 V at pH 3. The separation between the two peaks decreases with decreasing pH. The negative peak in the pH 5.4 unbuffered solution is broader and less well-formed than the others. This is due to the fact that in this solution the pristine CV shows two partially overlapping negative peaks. Only by several repeated voltage cycles between − 0.20 and − 1.20 V is a steady-state CV attained, in which the two negative peaks merge into one. CVs at a DPTL/DOPS tBLM were recorded under the same conditions as those at the DPTL/DOPC tBLM, yielding identical results within the limits of experimental error; they are shown in Fig. 2. Negative and positive peak potentials, Enp and Epp , and E1/2 values at the DPTL/ DOPC and DPTL/DOPS tBLMs are summarized in Table 1. CVs were also recorded at tBLMs whose distal monolayers consisted of DOPA and of the DOPC/Chol (70:30) binary mixture, which is known to form liquid-order microdomains (lipid rafts) surrounded by a liquid disordered matrix [19–21]. Measurements with DOPA and DOPC/Chol distal monolayers in unbuffered solution of pH 5.4 had no satisfactory level of reproducibility. Hence, only Enp ,Epp and E1/2 values at pH 3 and 6.8 are reported in Table 1. Figs. 3 and 4 show the CVs at DPTL/DOPA and DPTL/(DOPC/Chol) tBLMs at a scan rate of 50 mV/s. Uncertainties in the values of Table 1 are no greater than ±5 mV. Enp, Epp, and E1/2 values at the DPTL/DOPA tBLM are close to those at the DPTL/DOPC and DPTL/ DOPS tBLMs. On the other hand, the E1/2 values at the DPTL/ (DOPC/Chol) tBLM are shifted by about 20 mV toward less negative values at pH 3, and by 30 mV toward more negative values at pH 6.8, with respect to the corresponding values at the DPTL/DOPC and DPTL/ DOPS tBLMs. Fig. 5 shows CVs recorded at a DPTL/DOPC tBLM at a scan rate of 5 mV/s, under conditions otherwise identical with those in Fig. 1. The

Please cite this article as: L. Becucci, R. Guidelli, Can gramicidin ion channel affect the dipole potential of neighboring phospholipid headgroups?, Bioelectrochemistry (2015), http://dx.doi.org/10.1016/j.bioelechem.2015.06.008

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Fig. 2. The solid curves are experimental cyclic voltammograms at a DPTL/DOPS tBLM in an aqueous solution of 0.1 M KCl and 0.1 μM gramicidin A at pH 6.8, 5.4 and 3. The experimental cyclic voltammogram in the absence of gramicidin is shown by the dash-dotted curve in the upper frame and is practically identical at all three pH values. Scan rate = 50 mV/s. The corresponding dashed curves were calculated as described in the text. Fig. 1. The solid curves are experimental cyclic voltammograms at a DPTL/DOPC tBLM in an aqueous solution of 0.1 M KCl and 0.1 μM gramicidin A at pH 6.8, 5.4 and 3. The experimental cyclic voltammogram in the absence of gramicidin is shown by the dash-dotted curve in the upper frame and is practically identical at all three pH values. Scan rate = 50 mV/s. The corresponding dashed curves were calculated as described in the text.

separation between the negative and positive peaks is smaller than at a scan rate of 50 mV/s; Enp,Epp and E1/2 values are summarized in Table 1. At a scan rate of 5 mV/s, the negative peaks are higher and larger than the corresponding positive ones, and the charge obtained by their integration over time, inclusive of the small shoulder on their negative side, amounts to about −75 μC cm−2. Integration of the positive peaks yields a charge density close to + 60 μC cm− 2 and, hence, still higher than that from 45 to 50 μC cm− 2, corresponding to spacer saturation by K + ions, as estimated from potential-step chronocoulometric

measurements with all ion channels so far investigated [11,12], including the gramicidin channel [10,17]. Incidentally, the charge following chronocoulometric potential steps carried out in the presence of the gramicidin channel from an initial potential of −0.30 V, where K+ ions are prevented from flowing along the channel by the opposing electric field, to final potentials less negative than −1.20 V, where they enter the hydrophilic spacer, is practically equal in magnitude and opposite in sign to the K+ charge accumulating in the spacer, to maintain the electroneutrality of the whole mercury/water interface [10,17]. In fact, it can be shown that the charge density of the diffuse-layer ions is negligible with respect to that of the K+ ions in the spacer [10]. The anomalously high charge densities obtained by integration of the voltammetric peaks in Fig. 5 can be tentatively explained by postulating that the

Please cite this article as: L. Becucci, R. Guidelli, Can gramicidin ion channel affect the dipole potential of neighboring phospholipid headgroups?, Bioelectrochemistry (2015), http://dx.doi.org/10.1016/j.bioelechem.2015.06.008

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Table 1 Midpoint potential E1/2, negative peak potential Enp and positive peak potential Epp of cyclic voltammograms at mercury-supported tBLMs with distal DOPC, DOPS, DOPA and DOPC/Chol from pH 6.8 to pH 3 and 5.4. (70:30) monolayers in an aqueous solution of 0.1 M KCl and 0.1 μM gramicidin A at a potential scan rate of 50 mV/s, and peak potential shifts ΔEð6:8→pHÞ p For the DPTL/DOPC tBLM, parameters at a scan rate of 5 mV/s are also reported. DPTL/DOPC

pH 3

pH 5.4

pH 6.8

DPTL/DOPS

DPTL/DOPA

DPTL/(DOPS/Chol)

v = 5 mV/s

v = 50 mV/s

v = 50 mV/s

v = 50 mV/s

v = 50 mV/s

Enp

−0.41 V

−0.45 V

−0.45 V

−0.46 V

−0.43 V

Epp

−0.40 V

−0.43 V

−0.43 V

−0.39 V

−0.40 V

E1/2 ΔEnð6:8→3Þ p

−0.41 V +0.28 V

−0.44 V +0.37 V

−0.44 V +0.37 V

−0.43 V +0.35 V

−0.42 V +0.43 V

ΔEpð6:8→3Þ p

+0.18 V

+0.08 V

+0.08 V

+0.05 V

+0.11 V

Enp

−0.61 V

−0.67 V

−0.65 V

Epp

−0.51 V

−0.44 V

−0.45 V

E1/2 ΔEnð6:8→5:4Þ p

−0.56 V +0.08 V

−0.55 V +0.15 V

−0.55 V +0.17 V

ΔEpð6:8→5:4Þ p

+0.07 V

+0.07 V

+0.06 V

Enp

−0.69 V

−0.82 V

−0.82 V

−0.81 V

−0.86 V

Epp

−0.58 V

−0.51 V

−0.51 V

−0.44 V

−0.51 V

E1/2

−0.63 V

0.66 V

−0.66 V

−0.62 V

−0.69 V

very slow accumulation of K+ ions in the spacer at 5 mV/s scan rate may induce a slight elongation of the tetraethyleneoxy chain, with a resulting increase in its spaciousness. To comply with potential-step chronocoulometric measurements, a detailed analysis of the CVs will be confined to those recorded at a 50 mV/s scan rate.

The above experimental behavior and, in particular, the positive shift of E1/2 with decreasing pH, can be explained by a dragging of the lipid molecules immediately adjacent to the mouth of the gramicidin channel

Fig. 3. The solid curves are experimental cyclic voltammograms at a DPTL/DOPA tBLM in an aqueous solution of 0.1 M KCl and 0.1 μM gramicidin A at pH 6.8 and 3. Scan rate = 50 mV/s. The corresponding dashed curves were calculated as described in the text.

Fig. 4. The solid curves are experimental cyclic voltammograms at a DPTL/(70:30 DOPC/Chol) tBLM in an aqueous solution of 0.1 M KCl and 0.1 μM gramicidin A at pH 6.8 and 3. Scan rate = 50 mV/s. The corresponding dashed curves were calculated as described in the text.

4. Discussion

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identified with that estimated for the region of the lipoic acid residues, which amounts to about 5 μF cm−2 [13,23]. Hence, the experimental dependence of Cil upon the charge density FΓ can be approximately expressed by the equation: C il ¼ ð5 þ FΓ=45  195Þμ F cm−2

ð1Þ

according to which Cil varies linearly with FΓ from 5 to 200 μF cm−2. A DPTL/phospholipid tBLM can be regarded as consisting of: (i) the region of the lipoic acid residues; (ii) the tetraethyleneoxy hydrophilic spacer; (iii) the lipid bilayer moiety; and (iv) the polar head region, which is herein considered together with the diffuse layer region, for simplicity. The potential differences across the above four regions are approximately expressed, in the order, by the equation: Δϕ ¼

Fig. 5. Cyclic voltammograms at a DPTL/DOPC tBLM in an aqueous solution of 0.1 M KCl and 0.1 μM gramicidin A at pH 6.8 (solid curve), 5.4 (dashed curve) and 3 (dash-dotted curve). Scan rate = 5 mV/s.

toward the hydrocarbon tail region of the lipid bilayer. Thus, e.g., the DOPC distal monolayer is characterized by an arrangement of the (H3C)3N+-PO− 3 zwitterion coplanar to the monolayer, due to electrostatic interactions between trimethylammonium and phosphate groups of adjacent lipid molecules, with a resulting stabilization of the unprotonated form of the latter groups [22]. Due to the exposure of the phosphate group to the aqueous solution, its intrinsic pKa is about equal to 0.8, and hence its protonation starts to become appreciable around pH 2. However, if the (H3C)3N+-PO− 3 dipoles of the DOPC molecules adjacent to the mouth of a gramicidin channel are dragged closer to the hydrocarbon tail region, they will tilt with respect to the bilayer plane, with the phosphate group (linked to the glycerol backbone) being in a more embedded position with respect to the trimethylammonium group. The decreased polarizability of the local environment surrounding the phosphate group will tend to favor its protonation more than if it were exposed to the aqueous phase, causing a progressive decrease in its negative charge with decreasing pH from 6.8 to 3. At constant applied potential E, the resulting decrease in the negative potential difference across the polar head region is compensated for by an increasing negative potential difference across the hydrocarbon tail region (i.e., the intramembrane potential ϕm), causing an increase in the K+ flow into the spacer relative to its outflow. Therefore, the electric potential at which the K+ outflow matches its inflow (i.e., the midpoint potential E1/2) is attained at less negative potentials with decreasing pH. Similar conclusions apply to the polar heads of DOPS (see below). The experimental cyclic voltammetry behavior can be interpreted semiquantitatively on the basis of the above considerations by having recourse to the following approximate model. 4.1. Modeling the cyclic voltammetry behavior The K+ charge density, FΓ, where Γ is the K+ surface concentration and F the Faraday, intercalates with the lipoic acid residues and then, progressively, with the tetraethyleneoxy chains of the spacer, as the applied potential becomes progressively more negative [10]. Analysis of the electrochemical impedance spectra of a DPTL/DOPC tBLM incorporating the gramicidin ohmic channel from aqueous 0.1 M KCl reveals that the capacitance, Cil, of the “inner layer” interposed between the K+ charge density FΓ and the opposite charge density on mercury increases almost linearly with FΓ, attaining a value of about 200 μF cm−2 when FΓ attains its maximum saturation value, ranging from + 45 to +50 μC cm−2 [10]. On the other hand, in the absence of ion channels, and hence of ions in the spacer, the capacitance Cil can be approximately

    qM q þ FΓ q þ FΓ γ l þ M þ χsp þ M þ ðqM þ FΓ þ σ i Þ þ ð2Þ C il C sp Cm ε0 εγ ε0 εw

Here, Δϕ is the extra-thermodynamic absolute potential difference across the whole mercury/water interphase, qM is the surface charge density experienced by the diffuse-layer ions, σi is the surface charge density of any ionizable polar-head groups deeply embedded in the polar head region, and χsp is the surface dipole potential of the hydrophilic spacer. Cil, Csp, and Cm are the differential capacitances of the inner-layer region, the spacer and the lipid bilayer moiety, respectively; ε γ and ε w are the dielectric constants of the polar head region and of the aqueous phase, whereas ε0 is the permittivity of free space; ε 0 ε γ/γ is the differential capacitance of the polar head region and ε 0ε w/l is that of the diffuse layer in the linearized version of the Poisson–Boltzmann equation, where l equals √(ε 0 ε wR T/2 F 2c) and c is the concentration of a 1,1-valent electrolyte. The ether linkage between the tetraethyleneoxy spacer and the phytanyl chains and the ester linkage between the glycerol backbone and the fatty acids of the distal lipid monolayer have dipole potentials [24] that may partially compensate each other; in any case, these linkages have a fixed conformation that is not affected by pH changes in the aqueous solution. Hence, their surface dipole potentials are not included in Eq. (2). Upon extracting qM from Eq. (2) as a function of Δϕ and substituting the resulting expression into the equation, ϕm = (qM + F Γ)/Cm, for the intramembrane potential, after straightforward algebraic calculations we obtain: ϕm ≡


qM þ FΓ Δϕ þ FΓ=C il −χ sp −σ i γ=ε0 εγ þ l=ε0 εw   ¼ Cm C m C –1 þ C –1 þ C –1 þ γ=ε εγ þ l=ε εw il

sp

m

0

ð3Þ

0

Setting γ = 10 Å and ε γ = 10 [25], ε 0ε γ/γ is estimated at 9 μ F cm−2. Moreover, setting εw = 78, ε0εw/l amounts to 72 μ F cm−2 at room temperature and for c = 0.1 M. Several pieces of experimental evidence interpreted on the basis of a modelistic approach concur in estimating χsp at about −0.250 V [13]. In the absence of ion channels, Cm and Csp are about equal to 1 and 5 μF cm−2, respectively [23]. In the presence of ions in the spacer, induced by a pore or ion channel, both capacitances increase to some extent, but Cm remains smaller than Csp. Thus, C−1 m is higher than the reciprocal of all other capacitances, and, to a first approximation, Eq. (3) is simplified as follows:
ϕm ¼ Δϕ þ FΓ=C il –χ sp −σ i γ=ε0 εγ þ l=ε0 εw :

ð4Þ

The current I is obtained from the rate theory of ion transport across membranes [26–29], as applied to ion translocation across the potential energy barrier located in the lipid bilayer moiety of the tBLM:       ð1−α ÞFϕm αFϕm FΓ I∝ Fkt csp exp : −c0 exp − 1− 45 RT RT

ð5Þ

Please cite this article as: L. Becucci, R. Guidelli, Can gramicidin ion channel affect the dipole potential of neighboring phospholipid headgroups?, Bioelectrochemistry (2015), http://dx.doi.org/10.1016/j.bioelechem.2015.06.008

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Here, kt is the translocation rate constant, α is the transfer coefficient, csp is the volume concentration of the ions in the spacer, and c0 is that just outside the tBLM surface. The (1 − FΓ/45) factor is introduced to account approximately for the probability of finding free sites available for ion accommodation at the inner mouth of the ion channel; it vanishes when the K+ charge density in the spacer attains its saturation value of 45 μ C cm− 2. The cyclic voltammetry current I is exclusively controlled by ion translocation, and no ion depletion is assumed in the aqueous solution adjacent to the outer mouth of ion channels. Hence, c0 is set equal to the K+ bulk concentration. The csp value, in moles per unit volume, is obtained by dividing the moles of K+ ions in the spacer by its volume; thus, we have csp = Γ/d, where d is the length of the hydrophilic spacer, estimated at 2.25 nm [10]. For simplicity, the potential energy barrier in the lipid bilayer is assumed to be symmetrical by setting α = 0.5. Cyclic voltammetry curves are determined numerically by subdividing the potential range of interest into 1 × 106 δE steps. The current I is calculated at each step by Eq. (5) and is integrated by adding the corresponding charge contribution, δ(FΓ) = I δE/v, to the sum of all preceding contributions, where v is the potential scan rate. The resulting FΓ values are used to continuously update csp = Γ/d, Cil via Eq. (1) and ϕm via Eq. (4), and all these values are feedbacked into Eq. (5) for the current. Fig. 6 shows a number of CVs calculated for different values of the rate constant kt. The separation, Epp–Enp, between the positive and negative peaks decreases linearly with log kt up to kt = 104 cm/s, and then tends to zero, as shown by the inset in Fig. 6. The above model relies on parameters estimated independently in previous works on mercury-supported DPTL/phospholipid tBLMs and requires the translocation rate constant kt as the only adjustable parameter for the fitting to experimental CVs. This inherent simplicity is attained at the expense of a number of simplifying assumptions whose validity must be checked. One main assumption consists in considering that the average electric potential within the whole lipid bilayer also applies to the interior of the ion channel. In a previous work [29], we estimated the local electric

Fig. 6. Calculated curves of the current I against the absolute potential difference Δϕ across the mercury/(aqueous solution) interface at a mercury-supported DPTL/phospholipid tBLM incorporating a cation-selective ion channel and immersed in a 0.1 M aqueous solution of a 1,1-valent electrolyte. The rate constant kt for cation translocation equals: 1 × 102 (dashed curve), 1 × 103 (dash-dotted curve), 1 × 104 (short-dashed curve), 1 × 105 (dotted curve), and 1 × 106 cm/s (solid curve). Scan rate = 50 mV/s. The inset is a calculated plot of the separation, Epp–Enp, between the positive and negative peaks as a function of logkt. The solid circles mark the experimental values of (Epp–Enp) for the three cyclic voltammograms in Fig.1.

potential (called micropotential) created by a layer of dipoles in a cylindrical ion channel incorporated in a Hg-supported lipid monolayer enclosed between the metal and a perfectly conducting solution phase. The estimate, based on perfect conducting–conducting imaging conditions, depends on the length of the dipoles, assumed to point toward the metal, and on their location along the monolayer. The model predicts a symmetric micropotential profile with a smooth maximum or minimum at the midpoint of the monolayer. For a layer of short dipoles located on the solution side of the monolayer (i.e., in the polar head region), the micropotential profile at the midpoint of the monolayer practically coincides with the corresponding average potential, which equals σi l/(2ε 0 ε) (Fig. 7 in Ref. [29]); in this expression, l is the dipole length, σi is the charge of the positive pole, regarded as turned toward the hydrocarbon tail region, and ε is the dielectric constant of the dipole layer. This estimated dipolar contribution to the micropotential in the middle of the cylindrical ion channel is, therefore, in fairly good agreement with that, −σiγ/(ε 0 ε γ), present in the intramembrane potential expression of Eq. (4), in which the dipole is considered to be directed away from the hydrocarbon tail region, as in the case of the (H3C)3N+-PO− 3 dipole of a DOPC polar head dragged toward the hydrocarbon tail region. This provides some justification for the application of the average potential to the gramicidin channel. Another approximation consists in ascribing to the K+ volume concentration, csp, at the inner mouth of the ion channel its average value, Γ/d, in the spacer. In a preceding work, we calculated the K+ equilibrium concentration profile in the spacer moiety of a mercury-supported DPTL/DOPC tBLM incorporating the gramicidin channel as a function of the transmembrane potential, by solving the appropriate Poisson– Boltzmann equation (Fig. 7 in Ref. [10]). As expected, the K+ concentration profile increases in an approximately exponential manner in the direction of the metal at all potentials, up to the attainment of its maximum limiting value. However, the experimental value of the K+ charge saturating the spacer amounts to + 45 μ F cm−2, which corresponds to two potassium ions per tetraethyleneoxy chain of the hydrophilic spacer [10], whose length is estimated at 2.25 nm. Considering that the diameter of a desolvated K+ ion equals 0.30 nm and that of a water molecule equals 0.29 nm, the spacer length is almost completely covered by aligning the pentameric ring of the lipoic acid residue, which anchors the spacer to the metal and is about 0.30 nm in diameter, with two K+ ions solvated on both sides: (0.30 + 0.30 × 2 + 0.29 × 4) nm = 2.06 nm. In this respect, the K+ concentration profile calculated in Ref.

Fig. 7. Experimental (solid curve) and calculated cyclic voltammogram (dashed curve) at a DPTL/DOPC tBLM in an aqueous solution of 0.1 M KCl and 0.1 μM gramicidin A at pH 6.8. Scan rate = 5 mV/s.

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[10] provides a local probability of finding a K+ ion, rather than a proper K+ local volume concentration. In practice, we can feature the movement of K+ ions within the spacer along its normal as the surmounting of two low potential energy barriers to diffusion. This justifies the approximate use of the average K+ concentration in the spacer. Moreover, the outer valley of such a potential energy surface for K+ diffusion is expected to start being statistically filled only after the inner valley is almost completely filled. Hence, the probability for an inflowing K+ ion to find a free adsorption site can be regarded as roughly proportional to the fraction of the spacer “surface area” not yet occupied, i.e. (1 − FΓ/45), as assumed in Eq. (5). On the other hand, no hindrance to K+ flow out of the spacer is expected during the positive-going potential scan, at all K+ average concentrations csp. A minor approximation consists in ignoring Coulomb ion–ion interactions within the gramicidin channel. This is justified by the gramicidin channel being so narrow that ions can only move in a single file. Several experimental observations indicate that an ion moving across the gramicidin channel drags with it a long column of six to nine water molecules in single file [30], causing ion–ion interactions within the channel to be negligible. In addition, the electrostatic repulsion between the K+ ions stored in the spacer is notably screened by the electron charge on the mercury surface, which is almost equal in magnitude and opposite in sign to that of the K+ ions, to ensure the electroneutrality of the whole interfacial region. Potassium ions are also shielded by the water molecules interposed between them, and those in close contact with the metal surface by electron spillover from the metal. 4.2. A comparison with experimental cyclic voltammograms Comparing Figs. 1 and 2 with Fig. 6 might induce one to explain the pH dependence of the experimental CVs by an appreciable increase in kt with decreasing pH. The experimental (Epp–Enp) values for the DPTL/DOPC tBLM at pH 6.8, 5.4 and 3 and at a scan rate of 50 mV/s (see Table 1) are marked by solid circles in the inset of Fig. 6. They point to an increase in the translocation rate constant kt from 5 × 103 cm/s at pH 6.8 to 6 × 105 cm/s at pH 3. However, this approach is unrealistic, because it is highly improbable for a pH change in the bulk solution to have such a drastic effect on the potential energy barrier located inside the lipid bilayer moiety of the tBLM. Moreover, a mere increase in kt with decreasing pH would maintain E1/2 constant, as shown in Fig. 6, whereas its experimental value shifts in the positive direction. It is more reasonable to ascribe the pH dependence of the experimental CVs to a change in the surface dipole potential located in the polar heads of the distal lipid monolayer, as expressed by the σi(γ/ε0εγ) term in Eq. (4). A change in the charge density σi of ionizable polar-head groups is to be expected, especially if the ion channel is shorter than the thickness of the whole lipid bilayer, thus dragging the polar heads adjacent to its mouth closer to the hydrocarbon tail region. This is the case with the gramicidin channel, which has a length of 2.5–3.0 nm [31], as compared with an estimated lipid bilayer thickness of 3.5–4.0 nm. Unfortunately, there is no accurate way of estimating the absolute value of the σi(γ/ε 0 εγ) surface dipole term at a particular pH, and one must be satisfied with estimating the changes in this term with varying pH with respect to a pH value arbitrarily chosen as reference. For all the distal lipid monolayers investigated, we will choose as reference the experimental CV at pH 6.8. This choice is appropriate under the hypothesis that the positive shift of E1/2 with decreasing pH is determined by polar head dipoles with a negatively charged protonizable end turned toward the hydrocarbon tails. In this case the absolute value of the negative surface dipole potential decreases with decreasing pH. For the DPTL/DOPC and DPTL/DOPS tBLMs, both characterized by a (Epp–Enp) value of 0.31 V at pH 6.8, the corresponding calculated I vs. Δϕ curve was obtained from Eqs. (1), (4) and (5), with χsp = − 0.250 V [13] and σi = 0. The translocation rate constant providing the best fit was found to be 5 × 103 cm/s. To compare the calculated CV with the

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experimental CVs at pH 6.8 in Figs. 1 and 2, we must consider that the absolute potential difference Δϕ across the mercury/water interface is approximately related to the potential E measured against the Ag/ AgCl/0.1 M KCl reference electrode by the equation Δϕ = E + 0.190 V [13]. When the calculated CV is plotted against the E axis, it turns out to be shifted by 0.140 V toward more positive potentials with respect to the experimental ones at pH 6.8. To attain the best overlapping between the experimental CVs at DPTL/DOPC and DPTL/DOPS tBLMs and the calculated one, the latter was shifted by −0.140 V along the horizontal E axis. Moreover, its height was normalized to that of the experimental CVs, i.e., it was forced to be equal to that of the experimental CVs (see the dashed curves in Figs. 1 and 2). In view of Eq. (4) and of the fact that the calculated curve was obtained by setting σi = 0, one might be tempted to regard the −0.140 V shift as a measure of the surface dipole potential, σ i ðγ=ε 0 εγ þ l=ε 0 εw Þ at pH 6.8. However, no such significance should be attached to this value, since it depends on the arbitrary choice of the experimental CV at pH 6.8 for the initial fitting. Moreover, the shift was made with the only aim of obtaining the best overlapping of the calculated CV with the experimental ones at this reference pH value. A more reliable approach consists in identifying the changes, ΔEpnð6:8→pHÞ, in the negative , in the positive peak potentials, peak potentials and those, ΔEpð6:8→pHÞ p when passing from pH 6.8 to pH 5.4 and to pH 3, with the corresponding changes, Δσ i ðγ=ε0 εγ þ l=ε 0 εw Þ, in the surface dipole potential. Peak potentials and their changes are summarized in Table 1. Curves of I against Δϕ at pH 5.4 and 3 were calculated from Eqs. (1), (4) and (5) by setting again χsp = −0.250 V and kt = 5 × 103 cm/s, and equating σ i ðγ=ε0 εγ þ l=ε0 ε w Þ in Eq. (4) toΔEnð6:8→pHÞ along the negative voltage scan p along the positive one. In other words, they were genand to ΔEpð6:8→pHÞ p erated from the calculated curve at pH 6.8, taken as reference, by solely identifying the surface dipole potential changes with the corresponding peak potential changes. These calculated CVs were plotted against the E = (Δ ϕ − 0.190 V) axis, then shifted by − 0.140 V along the E axis, as done for the calculated curve at pH 6.8, and finally normalized to the height of the experimental CVs. Shifting the CVs calculated at pH 5.4 and 3 by − 0.140 V along the E axis amounts to starting from the calculated CV at pH 6.8, after its superimposition on the experimental one via a − 0.140 V shift along the E axis, and generating the other two calculated CVs by solely substituting σ i ðγ=ε0 εγ þ l=ε0 εw Þ in Eq. (4) with surface dipole potential changes equal to the corresponding peak potential changes. Differently stated, shifting all three calculated curves by the same quantity −0.140 V along the E axis serves to maintain their relative positions along the potential axis equal to those of the corresponding experimental CVs. The resulting calculated curves of the current density j against the applied potential E at pH 5.4 and 3 are shown in Figs. 1 and 2, together with the corresponding experimental CVs. This approach explains the progressive decrease in the separation between the negative and positive peak potentials with decreasing pH by a change in the surface dipole potential, rather than by a highly improbable notable increase in kt from 5 × 103 cm/s at pH 6.8 to 6 × 105 cm/s at pH 3. It also justifies the positive shift of the midpoint potential E1/2 with decreasing pH by a concomitant shift in the surface dipole potential of the polar heads of the distal lipid monolayer. The ΔEpnð6:8→pHÞ values for DOPC and DOPS in Table 1 being more values demand an explanapositive than the corresponding ΔEpð6:8→pHÞ p tion. As already stated, the DOPC distal monolayer is characterized by an arrangement of the (H3C)3N+-PO− 3 zwitterion coplanar to the monolayer [22]. However, if the (H3C)3N+-PO− 3 dipoles of the DOPC molecules adjacent to the mouth of a gramicidin channel are dragged closer to the hydrocarbon tail region, the phosphate group assumes a more embedded position with respect to the trimethylammonium group, and the decreased polarizability of its local environment favors its protonation more than if it were exposed to the aqueous phase. At pH 6.8

Please cite this article as: L. Becucci, R. Guidelli, Can gramicidin ion channel affect the dipole potential of neighboring phospholipid headgroups?, Bioelectrochemistry (2015), http://dx.doi.org/10.1016/j.bioelechem.2015.06.008

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the phosphate group can be regarded as still largely deprotonated, and hence the surface dipole potential term σ i ðγ=ε0 εγ þ l=ε0 ε w Þ is expected to be negative. However, at pH 5.4, and even more so at pH 3, the embedded phosphate group is expected to become appreciably protonated. The resulting loss of negative charge of the phosphate group determines a gradual positive shift in the Δσ i ðγ=ε0 ε γ þ l=ε0 εw Þ term as pH decreases from 6.8 to 3, in accordance with the experimental Δ values in Table 1. The ΔEnð6:8→pHÞ values being Epnð6:8→pHÞ and ΔEpð6:8→pHÞ p p values can possibly be exhigher than the corresponding ΔEpð6:8→pHÞ p plained by the dynamics of K+ flow in and out of the spacer and by its different directionality. The inflow of K+ ions across the channel mouth is favored by the adjacent (H3C)3N+-PO− 3 dipoles being directed with their negative pole toward the hydrophilic spacer, and may also tend to enhance their dipole moment. Conversely, K+ outflow is disfavored by such a dipole orientation, and may tend to decrease the tilt angle of the dipoles with respect to the bilayer plane. The proposed effect of K+ in- and outflow rates is supported by the ΔEnð6:8→pHÞ values p for DOPC being higher at a scan rate of 50 mV/s than at 5 mV/s, as shown in Table 1. It should be noted that the modelistic calculation of CVs through Eqs. (1), (4) and (5) also accounts satisfactorily for the experimental decrease in the separation between the negative and positive peaks with a decrease in the voltage scan rate, as appears by comparing the CVs for DOPC in Figs. 1 and 5 at the same pH. As an example, Fig. 7 shows the I vs. Δϕ curve calculated using the same parameters adopted for the calculated curve at pH 6.8 and 50 mV/s scan rate in Fig. 1, except for the voltage scan rate, which is set equal to 5 mV/s. This curve is directly compared with the corresponding experimental CV in Fig. 5 (solid curve in Fig. 7), after shifting it by −100 mV along the E axis and normalizing its height to the experimental one, in order to point out the fairly good agreement. The − 100 mV shift being smaller than the −140 mV shift required to superimpose the calculated CV at 50 mV/s on the corresponding experimental one in Fig. 1 is in keeping with the expected decrease in the polar-head surface dipole potential with a decrease in the K+ ion flow. The conclusions drawn about the pH effect at DPTL/DOPC tBLMs can be extended to DPTL/DOPS tBLMs. In the absence of exogenous species, the DOPS monolayer was shown to be positively charged at pH 3, almost neutral around pH 6 and negatively charged with a further increase in pH [22]. This behavior is explained by assuming an arrangement with the H3N+-COO− dipole coplanar to the monolayer, due to electrostatic interactions between ammonium and carboxyl groups of adjacent lipid molecules. The phosphate group, whose pK is about 8, is buried inside the polar head region; it is almost completely protonated at pH 6 and only partially deprotonated at pH 7. However, the dipoles of the DOPS molecules adjacent to the mouth of a gramicidin channel are expected to be dragged closer to the hydrocarbon tail region. Under these conditions, the phosphate group linked to the glycerol backbone will be in a more embedded position and, consequently, completely protonated even at pH 6.8. If, as a consequence of their inward dragging at the channel mouth, the H3N+ − COO− dipoles are somewhat tilted with respect to the bilayer plane, with the negative pole directed toward the hydrocarbon tail region, then the carboxyl group may play the same role as the phosphate group in the DOPC molecule. Thus, in spite of having a pKa of about 3 far from the channel mouth [22], it will be already partially protonated at pH 5.4 and completely protonated at pH 3, with a resulting gradual loss of its negative charge with decreasing pH. The values of the surface dipole potential differences reported in Table 1 should not be regarded as local values around the mouth of a gramicidin channel. Rather, they are average values throughout the whole polar head region of the distal monolayer. As such, they depend on the number density of the gramicidin channels in the lipid bilayer. Gramicidin has a low solubility in water (≤ 1 × 10−4 μM according to Hladky and Haydon [32]) and has been found to partition strongly

into the hydrophobic region of lipid membranes [33]. Hence, at the 0.1 μM gramicidin concentration adopted herein, much higher than its solubility value in pure water, it is quite reasonable for its concentration in the lipid bilayer to assume a value close to the 1:10 peptide/lipid molar ratio adopted in a theoretical analysis of hydrophobic matching [34], in a densitometry and sound velocimetry study on the effect of gramicidin [35], and in a STM imaging of gramicidin in a dimyristoylphosphatidylcholine (DMPC) monolayer on Au [36]. In particular, the counting of the lipid molecules surrounding each gramicidin channel yields an average number of 7 ± 1 [36], which is consistent with molecular dynamics calculations that assume eight nearestneighbor lipids [37]. This implies that the majority of lipid molecules in a gramicidin/lipid (1:10) mixture are recruited by the gramicidin ion channels. The cross-sectional area of a gramicidin channel is ~ 250 Å2 and that of a phosphatidylcholine molecule is ~62 Å2 [34,37]. Hence, assuming that the mouth of a single gramicidin channel is surrounded by eight nearest neighbor DOPC molecules, in the limiting case in which the phosphate group is entirely protonated the positive charge density for a 1:10 gramicidin/DOPC molar ratio amounts to 8e/[(250 + 10 × 62) Å2] = 14.7 μC cm−2, where e is the proton charge. The surface dipole potential expressed by σiγ/(ε0ε) is, then, given by 1.66 V, upon setting γ = 10 Å and ε = 10 [25]. The experimental CV for the DOPA monolayer at pH 6.8 (solid curve in Fig. 3) was fitted by an I vs. Δϕ curve calculated by the same procedure adopted for DOPC and DOPS. The calculated curve (dashed curve in Fig. 3) was obtained from Eqs. (1), (4) and (5) by setting χsp = −0.250 V, σi = 0 and kt = 2.5 × 103 s−1. It was, then, converted to the E = (Δϕ − 0.190 V) scale and overlapped with the experimental CV by shifting it by 100 mV toward more negative potentials and normalizing its height to that of the experimental CV. The curve of I against Δϕ for DOPA at pH 3 was then calculated from Eqs. (1), (4) and (5) by setting again χsp. = −0.250 V and kt = 2.5 × 103 s−1, and equating σ i ðγ=ε0 εγ þ l=ε0 ε w Þ in Eq. (4) to ΔEnð6:8→3Þ along the negative voltage scan p along the positive one; the experimental peak potential and to ΔEpð6:8→3Þ p shifts in passing from pH 6.8 to pH 3 are reported in Table 1. Overlapping of the calculated curve (dashed curve in Fig. 3) with the experimental CV was carried out as before, with the same negative shift by 100 mV along the E axis. The two protonation constants of the phosphate group of DOPA in biomimetic membranes assume values close to 1 × 108 M−1 and 1 × 104 M−1 [38,39]. Hence, the phosphate group of DOPA is uncharged at pH 3 and monoanionic at pH 6.8 far from the mouth of a gramicidin channel. At pH 6.8, the DOPA molecules around the gramicidin channel mouth, even if dragged closer to the hydrocarbon tail region, are not expected to become protonated, because they remain in close contact with water molecules. The fact that the pH dependence of the CVs for DOPA is close to that for DOPC and DOPS (see Table 1) lends support to the statement that, near a gramicidin channel, the phosphate group of DOPC and the carboxyl group of DOPS are protonated and neutral at pH 3, while they are negatively charged at pH 6.8. The experimental CVs for the DOPC/Chol distal monolayer were fitted by I vs. Δϕ curves calculated from Eqs. (1), (4) and (5) by an identical procedure, using a translocation rate constant kt of 2.7 × 103 s−1 and a negative shift of 160 mV along the E axis in order to overlap the calculated curves (dotted curves in Fig. 4) with the experimental CVs. Agreement between experimental and calculated curves is less satisfactory than in the case of the DOPC and DOPS monolayers, probably because the DOPC/Chol monolayer contains lipid rafts [19–21], and gramicidin channels are likely to be unevenly distributed between them and the liquid disordered matrix. The most significant difference between the cyclic voltammetry behavior of DOPC/Chol and that of the three pure phospholipids investigated is represented by an increase of about 60 mV in ΔEnð6:8→3Þ and of 30 mV inΔEpð6:8→3Þ , i.e., an increase p p in the quantities that we have identified with corresponding changes in surface dipole potential. This result is apparently surprising,

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notwithstanding the fact that Chol determines a significant increase in the dipole potential, when it is included into a biomimetic membrane [40,41]. In fact, such an increase is not expected to be directly affected by a change in pH. Moreover, the DOPC concentration is clearly less in the DOPC/Chol monolayer than in the DOPC one. However, it was more recently shown that indirect effects of Chol on the dipole potential of its mixtures with phosphatidylcholine (PC) are much more important than the direct ones. Starke-Peterkovic et al. [42] reported an increase in dipole potential from about + 0.35 V to + 0.45 V, positive toward the hydrocarbon tail, upon the addition of 40 mol% Chol to DOPC vesicles, and a six times higher increase with DMPC in place of DOPC. Different explanations for these indirect effects, which might also act synergically, were proposed. Thus, based on molecular dynamics simulations, Tu et al. [43] suggested that the PC polar heads rotate toward the bilayer to fill spaces left by cholesterol molecules, which sit more deeply in the membrane. Another indirect cause of dipole potential increase by Chol is its condensing effect on PC membranes [42]. Thus, the presence of 40 mol% Chol decreases the molecular area of egg PC from 61 to 49 Å2 [44] and that of DMPC from 58.5 to 53.8 Å2 [45]. The most significant indirect effect of Chol proposed by Starke-Peterkovic et al. [42] is represented by a significant decrease in water penetration into the bilayer, with a resulting consistent decrease in the dielectric constant in the region of the ester linkages of the glycerol backbone with the carboxyl groups of the fatty acids.

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Hg-supported tBLMs [11,12] is a powerful and versatile electrochemical technique, since it may also monitor the effect of a change in the rate of ion flow upon the current by varying the potential scan rate. It should be noted that applications of cyclic voltammetry to biomimetic membranes have been so far confined to the electron transfer across the lipid bilayer mediated by some redox couple [50,51], and not extended to the translocation of inorganic ions of biological interest, with only some exceptions at BLMs [46,47,52]. To the best of our knowledge, the present work provides the first quantitative treatment of CVs at tBLMs. This treatment relies on reasonable assumptions and on the knowledge of tBLM structural parameters deduced by independent means; in practice, the translocation rate constant kt is the only truly adjustable parameter. The conclusions drawn about the positive shift of the midpoint potential E1/2 and the decrease in the separation between the negative and positive peaks with decreasing pH can also be extended to other ion channels. Thus, we have recently verified that a similar behavior with varying pH is exhibited by the lipodepsipeptide syringopeptin (unpublished results). Moreover, the CV of the voltage gated ion channel of the lipodepsipeptide syringomycin shows a shift in the zero current potential with varying pH that is almost identical with the shift in the midpoint potential E1/2 of gramicidin A [53]. Acknowledgments

The different extent of the peak potential shifts ΔEnð6:8→3Þ and Δ p from distal monolayers of pure phospholipids to the DOPC/Chol Epð6:8→3Þ p mixture confirms that the change in the conformation of the lipid polar heads around the mouth of gramicidin channels with varying pH is sensitive to the lipid composition. The cyclic voltammetry behavior is also expected to be influenced by the dipole moment of the ion channel itself, when its mole fraction in the lipid bilayer becomes significant. Thus, the notable dependence of the current vs. potential curves of alamethicin upon its concentration in conventional bilayer lipid membranes [46,47] is likely to be affected by the high dipole moment of the alamethicin ion channel. In this respect, the situation of the gramicidin channel is particularly favorable. In fact, it is generally accepted that the major conformation of the gramicidin channel consists in a Nterminus-to-N-terminus helical dimer, with the C-terminus Trp residues in the two polar head regions of the membrane, due to their Hbonding capability and favorable electrostatic interactions [31,48,49]. Hence, the permanent dipole moment of the gramicidin channel is vanishingly small. 5. Conclusions In the absence of an external applied potential Δϕ, the source of the intramembrane potential of bilayer lipid membranes (BLMs) resides in its two polar heads. A symmetrical BLM interposed between two identical aqueous solutions has a zero intramembrane potential at Δϕ = 0, since charge and surface dipole potential on one side of the hydrocarbon tail region are perfectly counterbalanced by those on the opposite side. However, biological membranes are asymmetrical. The polar heads on the cytoplasmic side of plasma membranes are often more negatively charged than those on the extracellular side. Moreover, biological membranes incorporating proton pumps create a proton electrochemical potential gradient across them. It is, therefore, of interest to estimate the effect that pH changes in the solution bathing one side of the membrane exert on the contribution to the intramembrane potential from the polar heads on that side. A Hg-supported tBLM is particularly suitable in this respect, thanks to the very small and rigidly blocked polar-head region of its proximal lipid leaflet and to its high fluidity, mechanical stability and reproducibility. Its high resistance to electric fields allows the recording of CVs over a broad electric potential range under different environmental conditions, thus monitoring the ion flow both in and out of the hydrophilic spacer of tBLMs. Cyclic voltammetry applied to

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Please cite this article as: L. Becucci, R. Guidelli, Can gramicidin ion channel affect the dipole potential of neighboring phospholipid headgroups?, Bioelectrochemistry (2015), http://dx.doi.org/10.1016/j.bioelechem.2015.06.008