Phase-Transition-Induced Protein Redistribution in Lipid Bilayers

Nov 23, 2009 - KcsA proteins were reconstituted in proteoliposomes of POPE/POPG (3:1, mol/mol), and SLBs, including the proteins, were then obtained b...
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Phase-Transition-Induced Protein Redistribution in Lipid Bilayers Heiko M. Seeger,† Carlo A. Bortolotti,†,‡ Andrea Alessandrini,†,§ and Paolo Facci*,† CNR-INFM-S3 National Center on nanoStructures and bioSystems at Surfaces, Via Campi 213/A, 41125 Modena, and Department of Physics, UniVersity of Modena and Reggio Emilia, Via Campi 213/A, 41125 Modena, Italy ReceiVed: August 4, 2009; ReVised Manuscript ReceiVed: October 1, 2009

We report an atomic force microscopy study on the lateral spatial redistribution of an integral membrane protein reconstituted in supported lipid bilayers (SLBs) subjected to a thermally induced phase transition. KcsA proteins were reconstituted in proteoliposomes of POPE/POPG (3:1, mol/mol), and SLBs, including the proteins, were then obtained by the vesicle fusion technique on mica. By decreasing the temperature, the lipid bilayer passed from a liquid disordered (ld) phase in which the proteins are homogeneously distributed to a coexistence of solid ordered (so) and ld domains with the proteins preferentially distributed in the ld domains. The inhomogeneous distribution eventually led to protein clustering. The obtained results are discussed in light of the role that the lipid/protein interaction can have in determining the function of integral membrane proteins. Introduction It is well established that the activity of membrane proteins can be modulated by the lipid bilayer hosting the proteins. At the same time, the behavior of the lipid bilayer can be affected by the presence of protein inclusions.1-3 These mutual modulations can be the result of both specific and nonspecific interactions between the lipids and the proteins taking place at the molecular and macroscopic scales, respectively.4 The physical state of the membrane and its material properties are considered to play the most important role in the case of nonspecific interactions.5 Among the possible descriptors of the lipid-induced regulation mechanisms of membrane proteins, the hydrophobic matching is the central feature which is usually evoked to provide the conceptual framework for understanding the physical nature of such an interaction.6-8 In this scheme it is recognized that membrane protein functional activity is usually obtained by a conformational change involving the protein/lipid interface. The hydrophobic matching mechanism demands matching between the hydrophobic thickness of the proteins and that of the lipid bilayer in the immediate vicinity of the proteins. If the hydrophobic thickness of the protein and that of the lipid bilayer are different, the bilayer thickness in the surroundings of the protein is modified to avoid the formation of a nonpolar/ polar interface. The energetic cost of a change in the lipid bilayer thickness is quantitatively connected to the lipid bilayer physical properties.9 This energetic cost can alter the protein free energy landscape in the membrane, influencing the equilibrium distribution among different conformations. Many of the experimental studies performed to highlight the role of the lipid bilayer on membrane protein function are based on mechanosensitive channels,10,11 sodium ion channels,12 or pore-forming peptides such as gramicidin.13 Most of the obtained results can be * To whom correspondence should be addressed. E-mail: [email protected]. Phone: 0039 059 2055654. Fax: 0039 059 2055651. † CNR-INFM-S3 National Center on nanoStructures and bioSystems at Surfaces. ‡ Present address: Department of Chemistry, University of Modena and Reggio Emilia, Via Campi 183, 41125 Modena, Italy. § University of Modena and Reggio Emilia.

understood on the basis of a continuum model of the bilayer and have a broad validity not limited to a particular class of membrane proteins. Considering the crowded nature of biological membranes,14 the modification of the lipid bilayer imposed by a membrane protein can significantly influence the behavior of a neighboring protein, giving rise to cooperative effects. In this sense, an interpretation scheme based on the curvature free energy of monolayers and capillary forces can be exploited for the interpretation of protein/protein interactions induced by the interplay between lipids and membrane proteins.15 In this case the interaction between proteins extends beyond the first annular region surrounding each protein.2 The tendency of proteins to form clusters can be modulated both by the lipid:protein ratio and the hydrophobic mismatch between lipids and proteins.16 Another interpretation framework of how lipids can affect membrane protein function is based on the lateral pressure distribution inside the bilayer.17 Moreover, it has been demonstrated that also the coexistence of different physical domains (phases) in the plane of the membrane can influence the behavior of the proteins in it.18,19 This can be accomplished both by positioning the membrane proteins at the interface between two different phases, where packing defects favor conformational changes, and by segregating proteins in one particular phase, thereby increasing the possibility of protein/protein interactions. The latter case has received much attention in recent years due to the raft hypothesis of the biological membrane organization.20 Raft domains, which are cholesterol- and sphingolipid-enriched microdomains, are involved in fundamental trafficking and signaling processes in the cells by concentrating signaling proteins in the same lipid region. Also a lateral heterogeneity caused by the main phase transition between the liquid disordered and solid ordered phase of the lipid bilayer can be relevant for membrane protein function.21,22 It has been found that in model membranes R-helices are excluded from solid ordered and liquid ordered domains.23 The altered conformational change of rhodopsin molecules in membranes below their main phase transition has been attributed to the concentration of membrane proteins in domains of liquid disordered lipids,

10.1021/jp907505m  2009 American Chemical Society Published on Web 11/23/2009

Protein Redistribution in Lipid Bilayers forming dense rhodopsin-rich patches.24 Dealing with the presence of an interface between different phases, it has been reported by Cannon et al.21 that the sarcoplasmic reticulum Ca2+ release channel reconstituted in planar bilayers displays the highest activity at a lipid composition and temperature corresponding to the solid ordered/liquid disordered coexistence region. The presence of an attractive potential toward the domain interface area acting upon the channels has been evoked to explain the observed behavior.21 Thus, the proteins can be positioned in a packing-defect-rich phase where conformational changes are favored. The role of the presence of different domains means that, in general, physiologically relevant parameters, such as pH or ionic strength, which are able to alter the lateral heterogeneity of lipid bilayers, may also have an effect on membrane protein function. In the case of specific interactions between proteins and lipids it has been shown that many membrane proteins require the presence of specific lipids to perform their biological function,3 even if, in most cases, the molecular details of how the partnership between lipids and proteins is realized are not clear.25 The possibility of investigating the structure of a lipid bilayer containing membrane proteins along with their function would greatly enhance the understanding of the protein/lipid mutual interactions. Supported lipid bilayer (SLB) model systems appear to be a suitable experimental tool to reach this goal.26,27 The structure of an SLB and the protein distribution in SLBs at the single-molecule level can be unraveled by atomic force microscopy (AFM) in physiologic-like conditions without the need for fluorescent labels. In a previous work we studied the phase transition behavior of SLBs as a function of physical parameters such as the ionic strength of the solution, pH, and membrane preparation temperature.27 We established that if the SLB is assembled at a temperature higher than that of its main phase transition its behavior resembles that of liposomes, i.e., the bilayer model system usually exploited for studying protein/lipid interactions. This means that, apart from a shift to higher temperatures, the solid ordered to liquid disordered phase transition proceeds via a coupling of the two membrane leaflets instead of an independent melting of the bilayer leaflets as often found for SLBs. On the basis of this observation, it is possible to study some aspects of the protein/lipid interactions by means of SLBs. In the present work we have studied by AFM the redistribution of a K+ ion channel, namely, KcsA from Streptomyces liVidans, in the plane of a POPE/POPG (3:1, mol/mol) SLB upon formation of lipid domains resulting from the temperatureinduced main phase transition of the bilayer. Materials and Methods KcsA Expression, Purification, and Reconstitution. KcsA with an additional N-terminal hexahistidine sequence was expressed in BL21(DE3) cells grown in TB medium. Protein expression was induced by addition of 0.2 mg/L anhydrotetracycline (Acros Organics). The expressed protein was extracted with 20 mM decyl maltoside (DM) and purified by nickel affinity chromatography on a HisTrap FF crude column (Amersham Biosciences). The protein was eluted in 5 mM DM, 25 mM KCl, 50 mM NaH2PO4, 32 mM NaOH, and 400 mM imidazole, pH 7.0. Immediately after nickel affinity purification, the protein was concentrated and reconstituted by dialysis into POPE/POPG, 3:1 (mol/mol) lipid vesicles following the procedure described by Schmitt et al.28 with a protein-to-lipid ratio (w/w) ranging from 0.01 to 0.5.

J. Phys. Chem. B, Vol. 113, No. 52, 2009 16655 Sample Preparation and Atomic Force Microscopy Imaging. The supported lipid bilayers were prepared by the vesicle fusion technique. The protein/lipid suspension was sonicated for 30 s in an ultrasonic bath to obtain small unilamellar proteoliposomes (SUVs). Then we added 70 µL of our proteoliposome suspension on the mica. The lipid suspension was incubated for 15 min at 32 °C and then rinsed abundantly with 450 mM KCl, 25 mM Hepes, pH 7 buffer solution. As shown in an earlier work from our group, the incubation temperature chosen assured that the two membrane leaflets were coupled during the temperature-induced phase transition.27 The solution was then exchanged for the imaging solution (150 mM KCl, 10 mM potassium dihydrogen citrate at pH 7) by extensive rinsing. Then the mica support with the formed lipid bilayer was mounted on the temperature-controlled stage of the AFM instrument. AFM experiments were performed with a Bioscope equipped with a Nanoscope IIIA controller (Veeco Metrology). We built a temperature-controlled stage based on a circulating water bath on which we could mount the Bioscope head. Imaging was performed in tapping mode at a scan rate of 1-2 lines/s using triangular silicon nitride cantilevers (Olympus OMCL-TR400PB1, Japan) with a nominal spring constant of 0.09 N/m and a resonance frequency in liquid between 8 and 9 kHz. The force applied to the membrane was adjusted to the lowest possible value allowing reproducible imaging by varying the amplitude set point and/or the drive amplitude. The sample temperature was continuously monitored by a digital thermometer, Fluke 16 (Fluke, Italy), equipped with a small K-thermocouple probe (Thermocoax GmbH, Germany) in direct contact with the imaging buffer. ATR-FTIR Spectroscopy. Attenuated total reflection Fourier transformed infrared (ATR-FTIR) spectroscopy was exploited to correlate the number of bumps protruding from the lipid bilayers observed in AFM images and the amount of protein in the reconstituted liposomes. ATR-FTIR spectra were obtained by a Jasco 470 Plus spectrometer equipped with a DTGS detector. Spectra were acquired by averaging 256 acquisitions at 4 cm-1 resolution. The measurement chamber was constantly purged with a N2 flux at a flow rate of 1-2 L/min, and an equilibration time of 5 min was allowed after each chamber opening. A silicon trapezoidal crystal (80 × 10 × 5 mm3) with an incident angle of 45° allowing a total of 10 reflections on the sample side was used for the attenuated total reflection mode. Before each measurement the silicon crystal was exposed to a piranha solution (1:3 H2O2/H2SO4), washed extensively in bidistilled water (18 MΩ cm), and exposed to an oxygen plasma discharge in a plasma cleaner (Diener Electronic GmbH, Germany). This cleaning procedure assured a hydrophilic surface for the ATR crystal. To obtain oriented multibilayers, 300 µL of a solution containing KcsA reconstituted in proteoliposomes (lipid concentration 8 mg/mL) was added onto one side of the silicon crystal and dried under a N2 stream. The solution was spread uniformly all over the surface to obtain a homogeneously distributed film. It is well established that this preparation procedure results in the formation of stacked multibilayer structures in which the membranes are oriented with their plane parallel to the crystal surface.29 Results and Discussion A single solid supported proteolipid bilayer was assembled by the vesicle fusion technique on a mica surface and imaged by temperature-controlled AFM in tapping mode in liquid. An

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Figure 1. (a) AFM image of a POPE/POPG (3:1) supported lipid bilayer with reconstituted KcsA molecules. The bilayer was assembled by the vesicle fusion technique from proteoliposomes with a protein-to-lipid weight ratio of 300 µg/mg. (b) Statistics of the particle height above the lipid bilayer.

Figure 2. (a) Schematic diagram of the vectorial incorporation of KcsA molecules in vesicles. The cytoplasmic side of the molecules is located in the inner region of the vesicle. (b) After the vesicle fusion on the mica support the KcsA molecules expose their cytoplasmic side to the bulk water and the AFM tip.

AFM image of such a sample obtained from proteoliposomes with a protein-to-lipid ratio of 300 µg/mg acquired at 28 °C is shown in Figure 1a. At this temperature the SLB is completely in the liquid disordered phase.27 Many bumps emerge from the bilayer plane. These protrusions are ascribable to KcsA molecules exposing the intracellular portion to the AFM probe. KcsA is a homotetramer in which each subunit contains 160 residues with two transmembrane R-helices. Its crystal structure has been resolved by Doyle et al.30 for the protein lacking the C-terminus (from residue 126 to residue 158). The N-terminal residues together with the C-terminal ones form the cytoplasmic domain of the channel. The height distribution of the observed protrusions is peaked at 2 nm (Figure 1b). It is to be stressed that this protruding protein portion can be partially deformed by the scanning tip, accounting for the obtained dispersion of the height distribution. The obtained sample most likely results from a vectorial incorporation of the KcsA molecules in the POPE/POPG liposomes. This type of reconstitution has already been established in the case of KcsA in asolectin vesicles, where the channels are incorporated almost exclusively in the outsideout configuration as confirmed by a proteolytic assay.31 It has been demonstrated that the most probable pathway for vesicle rupture on hydrophilic surfaces results in a configuration in which the outer leaflet of the vesicles, once on the support, is the one which faces the support.32 Thus, vectoriality of the incorporation leads to a planar lipid bilayer in which the cytoplasmic side of KcsA faces the probe tip (Figure 2).

Moreover, the presence of a layer of trapped water between the lipid bilayer and the support avoids a direct contact between the membrane and the solid surface. This allows the lipid to maintain a lateral mobility.33 In addition, this configuration prevents proteins from a strong interaction with the mica support which could result in their partial denaturation and, likely, in a severe loss of lateral mobility. It has been reported that membrane proteins in supported lipid bilayers can be essentially immobile.34 This is especially the case when extramembraneous portions of the proteins are in direct contact with the substrate. Nonetheless, in AFM measurements of the lateral mobility of small membrane proteins, such as mobile influenza A M2, a protein diffusion coefficient on the order of 10-14 cm2/s has been reported.35 This value is about 4 orders of magnitude lower than that obtained on the upper surface of a multibilayer at the same temperature. The strongly reduced mobility can be attributed both to protein substrate interaction and to the fact that in AFM measurements only proteins which move not too fast with respect to the time resolution of the AFM are tracked, and as a consequence, inhomogeneities of the protein behavior are more evident. In our case, some of the molecules showed relative movements on the order of a few tens of nanometers between two consecutive images (time lag of 8 min) obtained at 30 °C (data not shown), highlighting a slow diffusion on the order of 10-14 cm2/s. However, we observed a strong dependence of the phenomenon upon the imaging temperature. Moreover, bumps attributed to the proteins are much more

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Figure 3. ATR/FT-IR spectra of POPE/POPG (3:1) plus KcsA multibilayers obtained for different protein-to-lipid weight ratios. The carbonyl and amide I and II regions are shown. The data have been normalized to the carbonyl peak intensity.

defined when the sample is imaged at low temperature, consistent with a reduced protein diffusion. It is to be stressed that a reduced mobility of KcsA channels in supported lipid bilayers does not necessarily imply a loss of channel functionality. A recent fluorescence study on the cooperativity of KcsA subunits performed on a solid SLB demonstrated that, in spite of an apparent absence of mobility for all the molecules, only a small fraction showed no activity in response to a change in pH.36 To further confirm that the observed bumps protruding from the bilayer can be associated with KcsA molecules, we analyzed the signal from ATR/FT-IR spectra on multibilayers obtained from the same proteoliposomes (KcsA in POPE/POPG, 3:1, at different weight ratios) used to obtain SLBs (Figure 3). The density of bumps observed by AFM on the SLBs assembled from proteoliposomes with different protein-to-lipid ratios was proportional to the corresponding amide signal. This evidence supports strongly the identification of the bumps with KcsA molecules. Moreover, the amide I signal is peaked at 1654 cm-1 and can be attributed mainly to the R-helix contribution accordingly with the known structure of KcsA, suggesting the preservation of the secondary structure of the proteins. In a previous investigation27 we studied the melting behavior of POPE/POPG SLBs in different conditions and found that, under preparation and imaging conditions similar to those of the present case, the transition from a solid ordered to a liquid disordered phase occurred with a coupled behavior of the two membrane leaflets. We then imaged the SLB with reconstituted KcsA molecules at a temperature of 28 °C, where all the bilayer was in the liquid disordered state (Figure 4a). We decreased the temperature to 26.5 °C, inducing the presence of a solid ordered lipid domain (Figure 4b). Remarkably, the growing solid domain tended to exclude the proteins, which remained mainly confined in the liquid disordered phase. Moreover, most of the proteins were found at the interface between the solid ordered domain and the liquid disordered area. By further decreasing the temperature, the solid ordered domain area increased and the proteins were mainly present in the decreasing liquid disordered fraction of the bilayer (Figure 4c-f). The proteins were eventually induced to form clusters. Parts b and c of Figure 4 show two consecutive images obtained after the cell temperature was set to 26.5 °C, highlighting the slow kinetics of domain formation in this system as observed in the absence of proteins.27 The KcsA channel has been thoroughly characterized as being the first K+ channel whose structure has been determined by X-ray crystallography. In particular, the interac-

Figure 4. AFM images (5000 × 5000 nm2) of a POPE/POPG (3:1) supported lipid bilayer with reconstituted KcsA molecules (300 µg/ mg) at different temperatures. In (a) all the bilayer at 28 °C is in the liquid disordered phase. Many KcsA molecules are found in the bilayer. In (b) we decreased the temperature to 26.5 °C while imaging the same area. A solid ordered domain developed, and the proteins were mainly excluded from the solid phase. The solid ordered domain area shape has been overlaid on the image in (a) to show the protein exclusion phenomenon. (c-f) Images at decreasing temperature (c, 26.5 °C; d, 26.0 °C; e, 25.0 °C; f, 23.0 °C) showing the growth of the solid ordered domain and the induced clustering of the proteins confined to the remaining liquid disordered areas. In (b) and (c) two consecutive images (time lag ∼8 min) at 26.5 °C are shown highlighting the slow kinetics of the domain formation process in the system.

tion of the KcsA with the lipid bilayer has been the subject of many investigations both theoretically and experimentally.37-39 The crystal structure of KcsA predicts a hydrophobic thickness for the protein of 3.7 nm, and binding constants for phosphatidylcholines established an optimum chain length of about C22, corresponding to a hydrophobic thickness of 3.13 nm.39 For our particular mixture of POPE/POPG (3:1) one can assume a hydrophobic thickness of 2.8-2.9 nm40 in the liquid disordered phase. These values correspond well to the hydrophobic thickness of a typical biological membrane in the liquid disordered phase. This means that our lipid bilayer in the liquid disordered phase has a hydrophobic thickness significantly lower than that of the best hydrophobic matching,41 indicating the presence of a large hydrophobic mismatch which has to be compensated by lipid deformation or, to a lower extent, by a tilt of R-helices. The difference in thickness between the solid ordered phase and the liquid disordered phase of the POPE/ POPG (3:1) bilayer, as measured from AFM topography, is 1.2-1.4 nm. Even if we consider a small contribution from the different mechanical properties of the two lipid phases in AFM images (no appreciable variation is found for this difference by changing the force applied by the tip), the hydrophobic thickness of solid ordered domains is about 4 nm. These data place the hydrophobic span of KcsA between those of the solid ordered phase and liquid disordered phase of our lipid system. The presence of the protein in a liquid disordered domain or in a solid ordered domain would imply a stretching or a compression

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of the lipids in the surroundings of the protein, respectively. In this situation the partition coefficient of KcsA in the two different phases, which represents the preference of the protein for the liquid disordered phase relative to the solid ordered one, cannot be calculated on the basis of the established theory which considers the free energy of lipid/protein interaction at the chainmelting transition.1 Moreover, a detailed calculation of the energetic cost of the lipid deformation to accommodate the hydrophobic mismatch could be performed on the basis of the theory of elastic bilayer deformations42 once the intrinsic monolayer curvature is known. In our case, the lipid composition of the solid ordered and liquid disordered domains may differ, due to a nonideal mixing of POPE and POPG lipids, leading to two different values for the intrinsic monolayer curvature. Our experimental evidence shows that KcsA, in the POPE/POPG (3:1) lipid bilayer, has a preference for the liquid disordered phase. The behavior in different lipid systems is currently under investigation. In those cases in which the hydrophobic thickness of the membrane protein is intermediate between the hydrophobic thicknesses of two different lipid domains, it has been shown, both experimentally and theoretically, that it is possible for the proteins to accumulate at the domain boundaries, where the hydrophobic thickness of the lipid bilayer is the most likely to satisfy the matching requirements.43,44 A possible consequence of the reduced area availability due to KcsA confinement in the bilayer liquid disordered phases is that protein clustering possibility increases. This situation can have a strong effect on the protein activity. It has been reported that the transport function of membrane proteins in lipid bilayers presents changes in the slope of the Arrhenius plot which are related to lipid phase transitions.45 In another report, the activity of a calcium channel of the sarcoplasmic reticulum has been related to the phase state of the hosting lipid bilayer.21 In the latter case an attractive potential acting on the proteins toward the defect-rich domain interface has been invoked as the driving force for the localization of the proteins in regions where their conformational changes are favored. In all these reports the localization of the proteins in the lipid bilayer is not established directly by imaging techniques, but only the correspondence of the phase transition region with the change in protein function is shown. In another study a modulation of KcsA ion channel activity has been attributed to protein clustering.46 In our study we establish by imaging evidence at the single-molecule level that membrane protein localization can be strongly influenced by a phase transition, which provides further support to the experimental findings of the cited works. In the specific case of KcsA, it has been demonstrated that the presence of specific lipids can modify the functional behavior of the channel.47 The presence of anionic lipids (POPG) was required for the functionality of KcsA, with a relationship between the open channel probability and the anionic lipid content. It is important to consider that a distribution of KcsA in specific lipid domains can alter the average lipid environment probed by the channel. In fact, the liquid disordered domains in a POPE/POPG (3:1) mixture are rich in POPG, leading to a probable effect on the channel functionality. It has to be stressed that a phase transition in lipid bilayers can be induced also by other parameters besides temperature. At constant temperature, a phase transition can occur due to pH, ionic strength, pressure, or lipid compositional changes. Even local changes of one of these parameters can be relevant to induce a localized phase transition of a lipid bilayer, modifying the function of the proteins embedded in that region.

Seeger et al. In the case of the present study, KcsA is a channel whose activity is gated by a change in pH.31 In general mixtures of PE and PG lipids have a strong dependence on pH.48 Performing differential scanning calorimetry experiments on the POPE/ POPG (3:1) mixture, we found a linear relationship between the transition midpoint temperature and pH.27 By decreasing the pH, the solid ordered phase is stabilized with an increase of the main phase transition temperature. A pH change can thus induce a modification of the lipid phase and the lateral lipid distribution, which could indirectly have an influence also on the membrane protein function. In the present work we have shown the correlation between lipid phase changes and protein localization/clustering in the bilayer. This evidence could impact further studies aimed at elucidating the mechanisms of activation of membrane proteins in interaction with lipid bilayers. Acknowledgment. This work has been performed with partial financial support of the Italian Ministry FIRB2007 Project “Italnanonet”. KcsA clones were a kind gift of C. Miller and A. Accardi. References and Notes (1) Marsh, D. Biochim. Biophys. Acta 2008, 1778, 1545. (2) Lee, A. G. Biochim. Biophys. Acta 2003, 1612, 1. (3) Lee, A. G. Biochim. Biophys. Acta 2004, 1666, 62. (4) Phillips, R.; Ursell, T.; Wiggins, P.; Sens, P. Nature 2009, 459, 379. (5) McIntosh, T. J.; Simon, S. A. Annu. ReV. Biophys. Biomol. Struct. 2006, 35, 177. (6) Mouritsen, O. G.; Bloom, M. Biophys. J. 1984, 46, 141. (7) Jensen, M. O.; Mouritsen, O. G. Biochim. Biophys. Acta 2004, 1666, 205. (8) Marsh, D Biophys. J. 2008, 94, 3996. (9) Andersen, O. S.; Koeppe, R. E. Annu. ReV. Biophys. Biomol. Struct. 2007, 36, 107. (10) Perozo, E.; Kloda, A.; Cortes, D. M.; Martinac, B. Nat. Struct. Biol. 2002, 9, 696. (11) Sukharev, S. I.; Sigurdson, W. J.; Kung, C.; Sachs, F. J. Gen. Physiol. 1999, 113, 525. (12) Lundbaek, J. A.; Birn, P.; Hansen, A. J.; Sogaard, R.; Nielsen, C.; Girshman, J.; Bruno, M. J.; Tape, S. E.; Egebjerg, J.; Greathouse, D. V.; Mattice, G. L.; Koeppe, R. E.; Andersen, O. S. J. Gen. Physiol. 2004, 123, 599. (13) Andersen, O. S.; Lundbaek, J. A.; Girshman, J. In Modelling the Dynamics of Biological SystemssNonlinear Phenomena and Pattern Formation; Mosekilde, E., Mouritsen, O. E., Eds.; Springer: New York, 1995; p 131. (14) Takamori, S.; Holt, M.; Stenius, K.; Lemke, E. A.; Gronborg, M.; Riedel, D.; Urlaub, H.; Schenck, S.; Brugger, B.; Ringler, P.; Muller, S. A.; Rammner, B.; Grater, F.; Hub, J. S.; De Groot, B. L.; Mieskes, G.; Moriyama, Y.; Klingauf, J.; Grubmuller, H.; Heuser, J.; Wieland, F.; Jahn, R. Cell 2006, 127, 831. (15) Botelho, A. V.; Huber, T.; Sakmar, T. P.; Brown, M. F. Biophys. J. 2006, 91, 4464. (16) Gil, T.; Sabra, M. C.; Ipsen, J. H.; Mouritsen, O. G. Biophys. J. 1997, 73, 1728. (17) Cantor, R. S. Chem. Phys. Lipids 1999, 101, 45. (18) Zhang, W.; Kaback, H. R. Biochemistry 2000, 39, 14538. (19) Micol, V.; Sanchez-Pinera, P.; Villalain, J.; de Godos, A.; GomezFernandez, J. C. Biophys. J. 1999, 76, 916. (20) Almeida, P. F. F.; Pokorny, A.; Hinderliter, A. Biochim. Biophys. Acta 2005, 1720, 1. (21) Cannon, B.; Hermansson, M.; Gyorke, S.; Somerharju, P.; Virtanen, J. A.; Cheng, K. H. Biophys. J. 2003, 85, 933. (22) Op Den Kamp, J. A. F.; Kauerz, M. T.; Van Deenen, L. L. M. Biochim. Biophys. Acta 1975, 406, 169. (23) Mall, S.; Broadbridge, R.; Sharma, R. P.; East, J. M.; Lee, A. G. Biochemistry 2001, 40, 12379. (24) Baldwin, P. A.; Hubbell, W. L. Biochemistry 1985, 24, 2633. (25) Hakizimana, P.; Masureel, M.; Gbaguidi, B.; Ruysschaert, J. M.; Govaerts, C. J. Biol. Chem. 2008, 283, 9369. (26) Sackmann, E. Science 1996, 271, 43. (27) Seeger, H. M.; Marino, G.; Alessandrini, A.; Facci, P. Biophys. J. 2009, 97, 1067. (28) Schmidt, D.; Jiang, Q. X.; MacKinnon, R. Nature 2006, 444, 775.

Protein Redistribution in Lipid Bilayers (29) Fringeli, U. P.; Gu¨nthard, H. H. Mol. Biol. Biochem. Biophys. 1981, 31, 270. (30) Doyle, D. A.; Cabral, J. M.; Pfuetzner, R. A.; Kuo, A. L.; Gulbis, J. M.; Cohen, S. L.; Chait, B. T.; MacKinnon, R. Science 1998, 280, 69. (31) Cuello, L. G.; Romero, J. G.; Cortes, D. M.; Perozo, E. Biochemistry 1998, 37, 3229. (32) Reimhult, E.; Kasemo, B.; Hook, F. Int. J. Mol. Sci. 2009, 10, 1683. (33) Kim, J.; Kim, G.; Cremer, P. S. Langmuir 2001, 17, 7255. (34) Salafsky, J.; Groves, J. T.; Boxer, S. G. Biochemistry 1996, 35, 14773. (35) Hughes, T.; Strongin, B.; Gao, F. P.; Vijayvergiya, V.; Busath, D. D.; Davis, R. C. Biophys. J. 2004, 87, 311. (36) Blunck, R.; McGuire, H.; Hyde, H. C.; Bezanilla, F. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 20263. (37) Marius, P.; Alvis, S. J.; East, J. M.; Lee, A. G. Biophys. J. 2005, 89, 4081. (38) Deol, S. S.; Domene, C.; Bond, P. J.; Sansom, M. S. P. Biophys. J. 2006, 90, 822. (39) Williamson, I. M.; Alvis, S. J.; East, J. M.; Lee, A. G. Cell. Mol. Life Sci. 2003, 60, 1581.

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