Article pubs.acs.org/JPCB
Lipid Diffusion in Alcoholic Environment Simona Rifici,† Carmelo Corsaro,† Cristina Crupi,† Valeria Conti Nibali,‡ Caterina Branca,† Giovanna D’Angelo,*,† and Ulderico Wanderlingh† †
Dipartimento di Fisica e Scienze della Terra, Università degli Studi di Messina, Messina, Italy Institute for Physical Chemistry II, Ruhr-University Bochum, Bochum, Germany
‡
ABSTRACT: We have studied the effects of a high concentration of butanol and octanol on the phase behavior and on the lateral mobility of 1,2palmitoyl-sn-glycero-3-phosphocholine (DPPC) by means of differential scanning calorimetry and pulsed-gradient stimulated-echo (PGSTE) NMR spectroscopy. A lowering of the lipid transition from the gel to the liquidcrystalline state for the membrane−alcohol systems has been observed. NMR measurements reveal three distinct diffusions in the DPPC−alcohol systems, characterized by a high, intermediate, and slow diffusivity, ascribed to the water, the alcohol, and the lipid, respectively. The lipid diffusion process is promoted in the liquid phase while it is hindered in the interdigitated phase due to the presence of alcohols. Furthermore, in the interdigitated phase, lipid lateral diffusion coefficients show a slight temperature dependence. To the best of our knowledge, this is the first time that lateral diffusion coefficients on alcohol with so a long chain, and at low temperatures, are reported. By the Arrhenius plots of the temperature dependence of the diffusion coefficients, we have evaluated the apparent activation energy in both the liquid and in the interdigitated phase. The presence of alcohol increases this value in both phases. An explanation in terms of a free volume model that takes into account also for energy factors is proposed.
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INTRODUCTION The properties of lipid membranes continue to attract considerable research interest. This relies on the relevance of membrane complexes as models for a range of processes of biochemical and medical interest. In particular, the study of the molecular interaction of short-chain alcohols with biological membranes is getting great value because of its role in metabolism, membrane fusion, drug delivery, alcohol toxicity, alcohol tolerance, and general anesthesia. Despite several mechanisms were proposed to explain this molecular interactions, there is not still a clear picture about it. In fact, it is not still assured if alcohols operate as anesthetics directly acting upon the receptors or indirectly influencing them by perturbing the lipid membranes.1 This problem is even more complicated if we take into account that the structure and phase behavior of a lipid membrane can be affected by the presence of impurities. Alcohols, for instance, decrease the main transition temperature between the gel phase, Lβ, and the fluid phase, Lα, of phospholipid membranes.2 Furthermore, as a consequence of the addiction of these amphiphilic molecules to lipid membranes, an interdigitated bilayer phase, LβI, may appear below the main transition temperature,3 and it is expected at alcohol concentrations above a threshold value equal to 2:1 alcohol/lipid ratio, as it has been observed for the DPPC/nbutanol system.4 The longer the lipid chain, the more energetically favorable the interdigitated state becomes.5,6 Although there are still lots of unresolved questions about the formation of this phase (the distribution of the alcohols in the bilayer, the exact molecular structure of the alcohol-induced © 2014 American Chemical Society
interdigitated phase, and the concentration of alcohol in the lipid bilayer at which interdigitation occurs), the most widespread hypothesis presumes that, because the hydrophobic portion of an alcohol favorably interacts with lipid hydrocarbon chains, the polar hydroxyl group remains free to form hydrogen bonds with polar lipid atoms located near the water/lipid interface. As a general picture of alcohol−membrane interaction, it is assumed that small alcohols (up to three carbon atoms) mainly interact by competing with water for hydration sites on membrane surface, whereas alcohols with longer chain preferentially bind inserting their tail within the membrane. More precisely, the OH− group binds to the phosphate group of the lipid headgroup7 as suggested by NMR and nuclear Overhauser effect spectroscopy experiments,8−11 and the hydrophobic tail sticks into the hydrophobic core of the bilayer. Lateral space is created between the headgroups, and since the nonpolar moieties of these amphiphilic molecules are not as long as the phospholipid hydrocarbon chains, voids between chains in the bilayer interior are created. As a consequence, the lipid chains interdigitate in order to minimize the energy of formation of these voids. This results in a variety of structural changes, like decreases in the hydrophobic thickness, 12,13 increases of lateral pressure,14,15 increases in the polar head area,13,18,19 and increases in lipid lateral mobility.16,17 Received: April 30, 2014 Revised: July 17, 2014 Published: July 18, 2014 9349
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Owing to the inherently low sensitivity of the NMR signal in gel-like samples, the limited bore diameter through the center of a high-field superconducting magnet, and the region of the bore that experiences a homogeneous magnetic field, a large amount of sample is required to perform this measurement. Furthermore, a sample for solid state NMR is usually composed of numerous stacked bilayers spread on mica substrates which introduce noise into NMR measurements. In the attempt of minimizing the signal/noise ratio, a particular homemade sample holder, consisting of a mica substrate, rolled inside a small hollow cylinder of Teflon, was realized. More precisely, a ∼30−40 μm thick sheet was obtained by repeatedly flaking and skinning. In this way, a lower quantity of mica was used, and it was possible to obtain a better level of hydration, since more free sample surface was exposed to a humid atmosphere. The mica plate with deposited sample was placed parallel to the magnetic field to test for lateral (in-plane) diffusion. The dynamical properties of pure DPPC and DPPC with alcohol molecules were studied by using a Bruker AVANCE NMR spectrometer operating at 700 MHz 1H resonance frequency. The 1H-PGSTE-NMR pulse sequence provides information on long-range lateral diffusion, up to some millimeter distances. In this sequence, the gradient amplitude, g, or duration, Δ, were varied while keeping all other parameters constant. The obtained data were analyzed by means of the Bruker-supplied XWIN-NMR software and the analysis was based on a modified version of the Stejskal− Tanner equation26
Under macroscopic equilibrium, i.e. without the presence of concentration gradients, the lipid molecules will change their position with time due to the Brownian, thermal motion. This kind of diffusion is called self-diffusion, and it is governed by the Einstein−Smoluchowski relation ⟨r2⟩ = 4Dt, where ⟨r2⟩ is the expected value of the square of the distance a given particle traveled during the time t and D is the self-diffusion coefficient (the area over which a molecule moves in 1 s). This process is very rapid and characterized by a diffusion coefficient of about 1 μm2 s−1. Essentially, studying lateral diffusion in lipids might help to understand the mechanism of membrane permeation and hence provide important information about membrane functionality. Lipids lateral motion, for instance, controls the microviscosity of the headgroup region and of the upper hydrophobic part of the phospholipids.20 Here we present a study about the effects that structural changes have on the phase behavior and on the lateral mobility of a particular artificial membrane, 1,2-palmitoyl-sn-glycero-3phosphocholine (DPPC), especially in the expected interdigitated phase. Studying lateral mobility below the main transition temperature has a particular importance considering the recent interest in characterizing membrane transport in the gel phase. Some proteins have higher affinity to partition into the gel phase with respect to the liquid phase.21 Moreover, the interdigitated phase has recently shown to have an important application in the preparation of potential drug encapsulating liposomes.22 Above all, PGSTE-NMR spectroscopy provides one of the most attractive techniques for studies of molecular transport, and the lipid lateral diffusion coefficients can be directly measured on macroscopically aligned bilayers.23
Ψ(δ , g , Δ, D) =
∑ Ai exp[−γ 2g 2δ 2Di(Δ − δ /3)] i
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(1)
where Ai are the spectral amplitudes without applied gradients, γ is the gyromagnetic ratio, Δ is the time interval between gradient pulses, and δ is the duration of the pulsed field gradients. The number of diffusion components, i, was varied from 1 to 3, and the sums of the squares of the residuals and inspection of the fits were used to determine the actual number of components, appropriate for each measurement. This experimental data analysis based on the multidiffusion processes is commonly used for the interpretation of NMR data for lipid bilayers.27 Measurements were performed at ambient pressure in the temperature range from −10 to 52 °C by means of cooling scans with a rate of 3−5 °C/min. Before starting the cooling process, samples were kept at T = 52 °C for some hours. All measurements were carried out in fully hydration condition.17 Temperature was controlled within ±0.5 °C by a heated air stream passing the sample. At each temperature several diffusion experiments were performed, in which the time interval, Δ, was varied in order to follow both water and lipid diffusions. Samples for DSC measurements, were prepared by dissolving DPPC in a 2:1 = CHCl3:CH3OH (chloroform/methanol) solution. After the evaporation of the solvent in a glovebox under N2 flux to prevent lipid oxidation, the appropriate amount of alcohols and water was added to the dry lipids in order to establish a molar ratio of lipid:alcohol:water of 1:2:10. The samples were incubated at 5 °C for at least 1 week before being studied. Multilamellar vescicles were prepared by subjecting the samples to ultrasonic irradiation in an ultrasonic water bath for about 30 min at a temperature of ∼10 °C above the melting temperature of the lipid. Then, samples were vortexed at room temperature four times and subjected to a
EXPERIMENTAL DETAILS It is worth observing that all data acquired were consistent and independent of the specific shape of the sample (vesicle or multibilayer) according to the experimental results of Katsaras,24 who has shown that highly aligned DMPC multibilayers and liposomal samples dispersed in an excess of liquid water exhibit the same physical characteristic (e.g., dspacing and transition temperatures) in all phases. Materials and Methods. The phospholipid 1,2-palmitoylsn-glycero-3-phosphocholine (DPPC), was obtained from Avanti Polar Lipids in powder form; butanol and octanol were purchased from Sigma Chemical Co. All chemicals have been used without further purification. Muscovite mica (purity level V-1) was purchased from SPI Chem., in rectangular sheets of about 45 × 25 mm2 and 150 μm thick. Mica was cut to obtain required dimensions. Highly aligned multibilayers samples for PGSTE-NMR experiments were prepared following the procedure suggested by Hallock et al.,25 using as supporting substrate a mica plate covered with approximately 1.5 mg of lipids per cm2. Following the cited procedure, DPPC was dissolved in an excess of 2:1 = CHCl3:CH3OH solution, which provided optimal adhesion and covering of the substrate, and the correct aliquot of alcohol to give a molar concentration of lipid:alcohol = 1:2 was then added. The solvent was removed by evaporation under nitrogen flux, and the dry samples were then stored for at least 2 weeks at 50 °C in 100% humidity atmosphere to be hydrated. The final water content was determined by comparison between wet and dry samples weights. 9350
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respectively. The observed spectra are a superposition of all spectra components present in the sample and provide information about the in-plane diffusion process which takes place in the membrane. Varying the time interval between the gradient pulses, Δ, we obtained a series of attenuated spectra. By measuring the maximum intensity of the 1H NMR spectra from the free-induction decay, the lipid diffusion constants were extracted, according to eq 1. As an example, we show in Figure 2 the diffusion decay of Ψ for (a) pure DPPC, (b) DPPC/butanol, and (c) DPPC/ octanol systems as a function of Q2Δ (where Q = γgδ/2π) at three different temperatures. The temperatures, indicated in the figure, were chosen at about 10 deg above the main transition temperature for each sample to compare all systems in the liquid disordered phase. Furthermore, the curves were normalized to the maximum intensity of Ψ. The transition temperatures were checked by means of DSC experiments. We show in the inset of Figure 2 the DSC heating curves for pure DPPC, DPPC/butanol, and DPPC/octanol systems. In pure DPPC, both pretransition (Tpre) and main transition (Tm) are clearly observed29,30 at 36 and 42 °C, respectively, as indicated by the presence of two peaks observed in the thermogram. The addition of both butanol and octanol results in a broadening and a shifting of the main transition to a lower temperature respect to that of the pure membrane: the longer the alcohol chains, the more the temperature Tm decreases. Furthermore, in all lipid/alcohol systems, the pretransition is suppressed, and this is indicative of a complete interdigitation of lipid chains.2,31 The decay of Ψ for pure DPPC shown in Figure 2a can be described, according to eq 1, as the sum of two components. One component is ascribed to water diffusion between bilayers, and the other is due to lipid diffusion. These two components
temperature cycle for 6 h to form multilamellar liposomes and obtain homogeneity. The comparison between our thermograms with literature data28 confirms that our samples are highly homogeneous multilamellar liposomes. Samples were weighed directly into the aluminum pans of the calorimeter. After samples were sealed, they were equilibrated at the lowest temperature for a period of time ranging from 0.5 to 2 h prior to starting the measurements. DSC scans were carried out from 5 to 45 °C by using a PerkinElmer Calorimeter Pyris 1 at a scan rate of 1 °C/min and repeated three times to ensure that thermal equilibrium was reached and to check for reproducibility.
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RESULTS AND DISCUSSION In Figure 1, a typical spin echo spectrum of pure DPPC is shown. The intense peak at 4.7 ppm is due to residual 1H from
Figure 1. NMR spectrum of pure DPPC at T = 40 °C.
intralamellar water while the peaks at 3.1 and 1.1 ppm originate from 1H in the methyl group and acyl chain methylene group,
Figure 2. Diffusion decay in fully hydrated: (a) DPPC bilayer at T = 52 °C, (b) DPPC/butanol at T = 46 °C, and (c) DPPC/octanol at T = 30 °C. Shown are the experimental data (open symbols), total fitting curves (continuous lines), and fitting components corresponding to water (dashdotted line), alcohol (dashed line), and lipid (dotted line) diffusion. In the inset the corresponding DSC heating curves are shown. 9351
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Figure 3. Self-diffusion coefficients as a function of T: (a) DW (circles) and DL (empty circles) in pure DPPC, literature data from ref 34 (left triangle) and ref 14 (right triangle); (b) DW (squares), DA (crossed squares), and DL (empty squares) in DPPC/butanol, literature data from ref 14 (right triangle); and (c) DW (top triangles), DA (crossed top triangles), and DL (empty top triangles) in DPPC/octanol. Transitions between liquid and gel phase are clearly visible and stressed with continuous lines.
are characterized by a diffusion coefficient of DW ∼ 10−10 m2/s and DL ∼ 10−12−10−11 m2/s, respectively, in agreement with previous literature data.27,32,33,35−38 As can be inferred from Figure 2b,c, in both alcohol systems, a further component, ascribed to alcohol molecule diffusion, with DA ranging from 3−11 to 4−10 m2/s has to be considered. In Figure 3 the diffusion coefficients DW, DA, and DL for pure DPPC, DPPC/butanol, and DPPC/octanol systems are shown as a function of T. The DW values of all systems depend slightly on Tm, while both DA and DL show a rapid change on the transition Tm, which is more evident in the sample with octanol. Furthermore, as concerning the lipid diffusion in the liquid phase, we found a good agreement with the literature data for pure DPPC14,34 and DPPC/butanol systems.14 It is worth noting that, to the best of our knowledge, this is the first time that self-diffusion coefficients have been derived for DPPC in the gel phase and for DPPC/alcohol systems in the interdigitated phase. With the aim to compare the lipid diffusion coefficients of the three investigated systems in the gel and in the liquid phase, in Figure 4 we show the DL values as a function of temperature normalized to Tm. In the liquid phase, the DL values decrease slowly with decreasing temperature, and in the systems with alcohols, they are higher respect to the pure membrane, at all temperatures. At temperature near Tm, a sudden slope is observed in all systems, and this is specially evident in the DPPC/octanol sample. In the interdigitated phase all the DL values flatten, showing a slight temperature dependence. Concerning the liquid phase, the presence of both butanol and octanol promotes the lipid diffusion process. A feasible explanation can be derived from the free volume theory, originally developed for describing the transport properties of glass-forming fluids39 and subsequently adapted to free-area
Figure 4. DL values for the three investigated systems as a function of T normalized to the corresponding main transition temperature.
theory to model two-dimensional diffusion in lipid bilayers in the liquid-crystalline phase.40−42 According to this theory, lateral diffusion of a particle with the size of the molecules that constitute the fluid takes place only when a free volume greater than a certain critical size exists next to the particle. Free volumes smaller than this critical size do not contribute to diffusion, and the problem is reduced to determining the distribution of free volume in the system. More specifically, in our case, the lateral diffusion coefficient DL should depend on the free area and on the packing properties as follows:40
D L ∼ e −a 0 / a f 9352
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Figure 5. Arrhenius plots of the temperature dependence of the obtained lateral diffusion coefficients (DL) for DPPC (a), DPPC with butanol (b), and DPPC with octanol (c). The solid lines are Arrhenius fits to the data. The apparent activation energies, Ea, and the corresponding error values, obtained both in the liquid and in the interdigitated phase areas of the investigated systems, are also indicated. Dashed regions refer to the liquid phase.
Figure 6. Arrhenius plots of the temperature dependence of the lateral diffusion coefficients (DA) for butanol (a) and octanol (b). The solid lines are Arrhenius fits to the data. The apparent activation energies, Ea and the corresponding error values, obtained both in the liquid and in the interdigitated phase areas, are also indicated. Dashed regions refer to the liquid phase.
e−Ea/kBT, where Ea is the activation energy associated with diffusion. In the case of a lipid bilayer, the activation energy takes into account (a) the interactions of a lipid molecule with its neighbors and with the bounding fluid in the bilayer and (b) the energy required to create a hole next to the diffusing molecule, whenever this event is locally associated with an energy change.40 Macedo and Litovitz43 proposed an hybrid equation
where a0 corresponds to the average cross-sectional molecular area for a DPPC molecule and af is a measure for the average amount of free area per molecule in the bilayer. The increasing of the polar head area of lipids caused by the addiction of alcohol in the bilayers13,18 gives rise to a smaller molecular packing; as a consequence, higher DL values are expected respect to the pure membrane. Furthermore, to jump to an adjacent empty site, a diffusing molecule needs energy to overcome an activation barrier. In the free area theory this is accounted for by supposing the lateral diffusion coefficient be proportional to a Boltzmann factor
D ∼ e(−a0 / af − Ea / kBT ) 9353
(3)
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Notes
which includes both the area and the energy factors and contains eq 2 as a limiting case.39 In Figure 5 the Arrhenius plots of the temperature dependence of the DL coefficients for DPPC, DPPC/butanol, and DPPC/octanol are shown. The apparent activation energies, Ea, and the corresponding error values, obtained both in the liquid and in the interdigitated phase, are also indicated. The presence of alcohol increases the value of Ea both in the liquid and in the interdigitated phases. The longer alcohol chains, the higher the apparent activation energies are. We want to underline that our data are in good agreement with previous literature data.44 Interestingly, we observe that, despite the increase of Ea, DL decreases. According to eq 3, we can argue that the area factor is more efficient than the energy factor in controlling the lateral diffusion of lipid. Concerning the interdigitated phase, lipid diffusion is hindered by butanol, and this effect is enhanced with increasing alcohol chain length. This can be explained taking into account the higher Ea with respect to the pure sample and considering that in the interdigitated phase the energy of formation of voids inside the bilayer is minimized. This results in a more packed structure and in a less free area per molecule af, available for diffusion. This is more evident for DPPC/octanol system respect to DPPC/butanol system, since interdigitation is more energetically favorable for alcohols with longer chains.5,6 Finally, in Figure 6 the Arrhenius plots of the temperature dependence of the lateral diffusion coefficients (DA) for butanol and octanol are shown. In the liquid phase, Ea is higher for butanol respect to octanol, while an opposite behavior is observed in the interdigitated phase. As concerning the interdigitated phase, tentatively, we suggest that the free volume available for butanol diffusion is larger respect to that of octanol and this results in a energetically favored configuration. But the exact explanation for these findings needs a dedicated study.
The authors declare no competing financial interest.
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CONCLUSIONS The effects of structural changes, caused by the addiction of a high butanol or octanol concentration, on the phase behavior and on the lateral mobility of DPPC have been studied by performing DSC and PGSTE-NMR spectroscopy measurements. Both liquid, Lα, and interdigitated, LβI, phases have been investigated, with a special attention given to the LβI phase. We found that the addition of alcohols causes a decrease of the main transition temperature of lipid membrane. The longer the alcohol chains, the more the temperature Tm decreases. Furthermore, in all lipid/alcohol systems, the pretransition appears to be suppressed, and this is indicative of a complete interdigitation of lipid chains. In the liquid phase the lipid diffusion process is promoted by the presence of both butanol and octanol, and the DL values decrease deeply as the temperature is decreased. An opposite trend is found in the interdigitated phase with DL coefficients that slightly depend on temperature. Lipid diffusion is hindered by the presence of butanol, and this effect is enhanced in systems with longer alcohol chain. These findings have been explained with a two dimension diffusion model that takes into account for both the free area available for diffusion and the activation energy of the diffusion process.
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REFERENCES
(1) Uda, N. R.; Rajashekar, K.; Wei, S. Calorimetric Investigation of Membrane Anesthetic Interactions. Thesis, 2008. (2) Löbbecke, L.; Cevc, G. Effects of Short-Chain Alcohols on the Phase Behavior and Interdigitation of Phosphatidylcholine Bilayer Membranes. Biochim. Biophys. Acta 1995, 1237, 59−69. (3) Slater, J. L.; Huang, C. H. Interdigitated Bilayer Membranes. Prog. Lipid Res. 1988, 27, 325−359. (4) Zhang, F.; Rowe, E. S. Titration Calorimetric and Differential Scanning Calorimetric Studies of the Interactions of n-Butanol with Several Phases of Dipalmitoylphosphatidylcholine. Biochemistry 1992, 31, 2005−2011. (5) Adachi, T. A New Method for Determining the Phase in the XRay Diffraction Structure Analysis of Phosphatidylcholine/Alcohol. Chem. Phys. Lipids 2000, 107, 93−97. (6) Adachi, T.; Takahashi, H.; Ohki, K.; Hatta, I. Interdigitated Structure of Phospholipid-Alcohol Systems Studied by X-Ray Diffraction. Biophys. J. 1995, 68, 1850−1855. (7) Chiou, J. S.; Krishna, P. R.; Kamaya, H.; Ueda, I. Alcohols Dehydrate Lipid Membranes: an Infrared Study on Hydrogen Bonding. Biochim. Biophys. Acta 1992, 1110, 225−233. (8) Barry, J. A.; Gawrisch, K. Direct NMR Evidence for Ethanol Binding to the Lipid-Water Interface of Phospholipid Bilayers. Biochemistry 1994, 33, 8082−8088. (9) Holte, L. L.; Gawrisch, K. Determining Ethanol Distribution in Phospholipid Multilayers with MAS-NOESY Spectra. Biochemistry 1997, 36, 4669−4674. (10) Feller, S. E.; Brown, C. A.; Nizza, D. T.; Gawrisch, K. Nuclear Overhauser Enhancement Spectroscopy Cross-Relaxation Rates and Ethanol Distribution Across Membranes. Biophys. J. 2002, 82, 1396− 1404. (11) Patra, M.; Salonen, E.; Terama, E.; Vattulainen, I.; Faller, R.; Lee, B. W.; Holopainen, J.; Karttunen, M. Under the Influence of Alcohol: the Effect of Ethanol and Methanol on Lipid Bilayers. Biophys. J. 2006, 90, 1121−1135. (12) Rifici, S.; Crupi, C.; D’Angelo, G.; Di Marco, G.; Sabatino, G.; Conti Nibali, V.; Trimarchi, A.; Wanderlingh, U. Effects of a Short Length Alcohol on the Dimyristoylphosphatidylcholine System. Philos. Mag. 2011, 91 (13), 2014−2020. (13) Wanderlingh, U.; D’Angelo, G.; Conti Nibali, V.; Crupi, C.; Rifici, S.; Corsaro, C.; Sabatino, G. Interaction of Alcohol with Phospholipid Membrane: NMR and XRD Investigations on DPPC/ Hexanol System. Spectroscopy 2010, 24, 375−380. (14) Dickey, A.; Faller, R. How Alcohol Chain-Length and Concentration Modulate Hydrogen Bond Formation in a Lipid Bilayer. Biophys. J. 2007, 92, 2366−2376. (15) Tu, K.; Tarek, M.; Klein, M. L.; ScharfTu, D. Effects of Anesthetics on the Structure of a Phospholipid Bilayer: Molecular Dynamics Investigation of Halothane in the Hydrated Liquid Crystal Phase of Dipalmitoylphosphatidylcholine. Biophys. J. 1998, 75, 2123− 2134. (16) Chen, S. Y.; Yang, B.; Jacobson, K.; Sulik, K. K. The Membrane Disordering Effect of Ethanol on Neural Crest Cells in Vitro and the Protective Role of Gm1 Ganglioside. Alcohol 1996, 13, 589−595. (17) Klemm, W. R. Biological Water and its Role in the Effects of Alcohol. Alcohol 1998, 15, 249−267. (18) Kranenburg, M.; Smit, B. Simulating the Effect of Alcohol on the Structure of a Membrane. FEBS Lett. 2004, 568, 15−18. (19) Ly, H. V.; Longo, M. L. The influence of Short-Chain Alcohols on Interfacial Tension, Mechanical Properties, Area/Molecule, and Permeability of Fluid Lipid Bilayers. Biophys. J. 2004, 87, 1013−1033. (20) Venable, R. M.; Zhang, Y.; Hardy, B. J.; Pastor, R. W. Molecular Dynamics Simulations of a Lipid Bilayer and of Hexadecane: an Investigation of Membrane Fluidity. Science 1993, 262, 223−226.
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(42) Tian-xiang, X. Translational Diffusion in Lipid Bilayers: Dynamic Free-Volume Theory and Molecular Dynamics Simulation. J. Phys. Chem. B 1999, 103, 385−394. (43) Macedo, P. B.; Litovitz, T. A. On the Relative Roles of Free Volume and Activation Energy in the Viscosity of Liquids. J. Chem. Phys. 1965, 42, 245−256. (44) Oradd, G.; Lindblom, G.; Westerman, P. W. Lateral Diffusion of Cholesterol and Dimyristoylphosphatidylcholine in a Lipid Bilayer Measured by Pulsed Field Gradient NMR Spectroscopy. Biophys. J. 2002, 83, 2702−2704.
(21) Clay, A. T.; Sharom, F. J. Lipid Bilayer Properties Control Membrane Partitioning, Binding, and Transport of P-Glycoprotein Substrates. Biochemistry 2013, 52, 343−354. (22) Smith, E. A.; Wang, W.; Phoebe, K. Dea. Effects of Cholesterol on Phospholipid Membranes: Inhibition of the Interdigitated Gel Phase of F-DPPC and F-DPPC/DPPC. Chem. Phys. Lipids 2012, 165, 151−159. (23) Oradd, G.; Lindblom, G. Lateral Diffusion Studied by Pulsed Field Gradient NMR Oriented Lipid Membranes. Magn. Reson. Chem. 2004, 42, 123−131. (24) Katsaras, J. Adsorbed to a Rigid Substrate, Dimyristoylphosphatidylcholine Multibilayers Attain Full Hydration in all Mesophases. Biophys. J. 1998, 75, 2157−2162. (25) Hallock, K. J.; Henzler Wildman, K.; Lee, D. K.; Ramamoorthy, A. An Innovative Procedure Using a Sublimable Solid to Align Lipid Bilayers for Solid-State NMR Studies. Biophys. J. 2002, 82, 2499−2503. (26) Stejskal, E. O.; Tanner, J. E. Spin Diffusion Measurement: Spin Echoes in the Presence of a Time-Dependent Field Gradient. J. Chem. Phys. 1965, 42, 288−292. (27) Oradd, G.; Westerman, P. W.; Lindblom, G. Lateral Diffusion Coefficients of Separate Lipid Species in a Ternary Raft-Forming Bilayer: A Pfg-NMR Multinuclear Study. Biophys. J. 2005, 89, 315− 320. (28) Suurkuusk, M.; Singh, S. K. Microcalorimetric Study of the Interaction of 1-Hexanol with Dimyristoylphosphatidylcholine Vesicles. Chem. Phys. Lipids 1998, 94, 119−138. (29) Tristam-Nagle, S.; Wiener, M. C.; Yang, C. P.; Nagle, J. F. Kinetics of the Subtransition in Dipalmitoylphosphatidylcholine Dispersions. Biochemistry 1987, 26, 4288−4294. (30) Mellier, A.; Diaf, A. Infrared Study of Phospholipid Hydration. Main Phase Transition of Saturated Phosphatidylcholine/Water Multilamellar samples. Chem. Phys. Lipids 1988, 46, 51−56. (31) Kurniawan, Y.; Venkataramanan, K. P.; Scholz, C.; Bothun, V. N-Butanol Partitioning and Phase Behavior in DPPC/DOPC Membranes. J. Phys. Chem. B 2012, 116, 5919−5924. (32) Wassal, S. R. Pulsed Field Gradient-Spin Echo NMR Studies of Water Diffusion in a Phospholipid Model Membrane. Biophys. J. 1996, 71, 2724−2732. (33) Lindblom, G.; Wennerstrom, H.; Arvidson, G. Translational Diffusion in Model Membranes Studied by Nuclear Magnetic Resonance. J. Quantum Chem. 1977, 12, 153−158. (34) Lindblom, G.; Oradd, G.; Filippov, A. Lipid Lateral Diffusion in Bilayers with Phosphatidylcholine, Sphingomyelin and Cholesterol An NMR Study of Dynamics and Lateral Phase Separation. Chem. Phys. Lipids 2006, 141, 179−184. (35) Hansen, F. Y.; Peters, G. H.; Taub, H.; Miskowiec, A. Diffusion of Water and Selected Atoms in DMPCLipid Bilayer Membranes. J. Chem. Phys. 2012, 137, 204910(15 pages). (36) Lindblom, G.; Oradd, G. NMR Studies of Translational Diffusion in Lyotropic Liquid Crystals and Lipid Membranes. Prog. Nucl. Magn. Reson. Spectrosc. 1994, 26, 483−515. (37) Sanders, M.; Mueller, R.; Menjoge, A.; Vasenkov, S. Pulsed Field Gradient Nuclear Magnetic Resonance Study of Time-Dependent Diffusion Behavior and Exchange of Lipids in Planar-Supported Lipid Bilayers. J. Phys. Chem. B 2009, 113, 14355−14364. (38) Wang, L.; Schö n hoff, M.; Mö h wald, H. Swelling of Polyelectrolyte Multilayer-Supported Lipid Layers. 1. Layer Stability and Lateral Diffusion. J. Phys. Chem. B 2004, 108, 4767−4774. (39) Cohen, M. H.; Turnbull, D. Molecular Transport in Liquids and Glasses. J. Chem. Phys. 1959, 31, 1164−1169. (40) Almeida, P. F. F.; Vaz, W. L. C.; Thompson, T. E. Lateral Diffusion in the Liquid-Phases of Dimyristoylphosphatidylcholine Cholesterol Lipid Bilayers-a Free-Volume Analysis. Biochemistry 1992, 31, 6739−6747. (41) Vaz, W. L. C.; Hallmann, D.; Clegg, R. M.; Gambacorta, V.; De Rosa, M. A Comparison of The Translational Diffusion of a Normal and a Membrane-Spanning Lipid in Lα Phase 1-Palmitoyl-2oleoylphosphatidylcholine Bilayers. Eur. Biophys. J. 1985, 12, 19−24. 9355
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