摘要:

28 I 29 1 303 313 329 339 369 371 Membrane Bending Elasticity and its Role for Shape Fluctuations and Shape Transformations of Cells and Vesicles E. Sackmann, H-P. Duwe and H. Englehardt Adsorption of Phospholipid Vesicles on Solid Surfaces S. Jackson, M. D. Reboiras, I. G. Lyle and M. N. Jones Liposome Electroformation M. I. Angelova and D. S. Dimitrov The Distribution of Substituted Phenols into Lipid Vesicles S. S. Davis, M. J. James and N. H. Anderson Membrane-spanning Symmetric and Asymmetric Diyne Amphiphiles H. Bader and H. Ringsdorf Genera 1 Discussion List of Posters Index of Names28 I 29 1 303 313 329 339 369 371 Membrane Bending Elasticity and its Role for Shape Fluctuations and Shape Transformations of Cells and Vesicles E. Sackmann, H-P. Duwe and H. Englehardt Adsorption of Phospholipid Vesicles on Solid Surfaces S. Jackson, M. D. Reboiras, I. G. Lyle and M. N. Jones Liposome Electroformation M. I. Angelova and D. S. Dimitrov The Distribution of Substituted Phenols into Lipid Vesicles S. S. Davis, M. J. James and N. H. Anderson Membrane-spanning Symmetric and Asymmetric Diyne Amphiphiles H. Bader and H. Ringsdorf Genera 1 Discussion List of Posters Index of Names

ISSN:0301-7249    

DOI:10.1039/DC98681FX001

出版商:RSC    

年代:1986

数据来源: RSC

摘要:

Faraday Discuss. Chem. SOC., 1986,81, 19-28 Equilibrium Studies of Phospholipid Bilayer Assembly Coexistence of Surface Bilayers and Unilamellar Vesicles Norman L. Gershfeld,* William F. Stevens Jr and Ralph J. Nossal National Institutes of Health, Bethesda, Maryland 20892, U.S.A. To understand how single lipid bilayers might form in cells, we have examined the equilibrium phase diagram of dilute dimyristoylphos- phatidylglycerol (DMPG) dispersions in water. Using surface pressure measurements, quasielastic light scattering and phase-contrast microscopy, unilamellar vesicle formation has been observed at a critical temperature, T*, the identical temperature where single bilayers form in films of the lipid at the air-water surface. T* is ca. 8°C higher than the gel-liquid-crystal transition temperature, T,, for DMPG dispersions.At T < T*, DMPG exists as a jelly which appears to consist of a matrix of extended bilayer sheets. At T > T*, DMPG dispersions form rnultilamellar vesicles. The solubility of DMPG in water, X,, was measured over the temperature interval of 8-40 "C. The function dlnX,/dT was found to be discontinuous at T,, but was continuous in the temperature interval that encompasses T*. It is generally believed that the lipids in cell membranes are confined to a single bilayer, and that the lipid bilayer is assemb.led by spontaneous processes similar to those observed in artificial systems. This viewpoint is reasonable given the ubiquity of the bilayer state in phospholipid-water dispersions and the observation that cell membrane bilayer assembly can occur even after protein synthesis is suppressed.'.* However, it is not at all clear why only single lipid bilayers form in cell membranes.Single bilayers, when observed in phospholipid dispersions, generally are accompanied by multilamellar structures: or are formed under non-biological conditions with use of either sonication; organic solvents' or detergents or other lipids not normally present in cells.6 Whether conditions ever exist when only single lipid bilayers form spontaneously is a question of considerable importance for understanding the process of membrane bilayer assembly in cells. We previously sought conditions for spontaneous assembly of single bilayers by examining the properties of the equilibrium air-water surface of phospholipid disper- sions.The results of recent systematic studies indicate that films composed of a single bilayer (the surface bilayer) form spontaneously at air-water surfaces at a temperature T" that depends on the phospholipid in the By thermodynamic arguments it has been deduced that at T", the temperature where surface bilayers appear, single bilayers also form in the equilibrium bulk This prediction has been tested now by examining the phase diagram of aqueous dispersions of a phospholipid that forms a surface bilayer. We have utilized dynamic light scattering, phase-contrast microscopy and calorimetry to demonstrate that single bilayers in the form of vesicles appear in the equilibrium bulk dispersion at the same temperature as that of surface bilayer formation. Finally, from the temperature dependence of the lipid's solubility in water, we establish that the bilayer structure is sustained even in very dilute dispersions. 1920 Pospholipid Bilayer Assembly Dimyristoylphosphatidylglycerol (DMPG) has been selected as the phospholipid for study primarily because its solubility in water is sufficiently high that measurement of the temperature dependence of its chemical potential by standard thermodynamic methods is feasible.Moreover, it exhibits bilayer properties similar to the less soluble phosphatidylcholine analogue, DMPC". As a result of its ionic character, at high dilution DMPG dispersions are expected to form single bilayers.'* Methods Materials DMPG as the sodium salt (Avanti Polar Lipids, Inc., Birmingham, AL) with a purity >99% by thin-layer chromatography, was used without further purification. Hydrolysis of DMPG to the lysophosphatidylglycerol compound was observed in dilute aqueous dispersions; at room temperature the rate of hydrolysis was ca.0.5% in 24 h, increasing slightly with temperature. However, in our experiments hydrolysis was not significant, because all measurements were completed within 48 h. Solubility The procedure for determining the solubility of DMPG in water entailed preparing dispersions by vortexing various amounts of the lipid in water, incubating at various temperatures, and separating the excess lipid from the solution by centrifugation. Generally, 2-5 times the saturation concentration of DMPG was used. A model L5-65 Beckman centrifuge with an SW-40 rotor was employed, with temperature control from 0 to 45 "C maintained to k0.5 "C.Temperature equilibration was attained within 3-4 h. At each temperature six centrifuge tubes were run simultaneously. Aliquots of the supernatant were taken from the uppermost portion of each tube for analysis; they were evaporated and the phosphorus content of each residue was a ~a1ysed.l~ At temperatures below 25 "C, centrifugation at 100 OOOg for 10 h gave so1ut:on concentrations that were independent of the amount of lipid in the dispersion. Aoove 25 "C, centrifugation for 20 h was required to attain solution concentrations that were independent of the amount of lipid originally added; longer centrifugation times did not change the solution concentrations.Therefore, 20 h centrifugation was used throughout, in order to maintain the same experimental conditions. Longer centrifugation times were avoided to minimize contamination due to hydrolysis. Surface Pressures Surface pressures were measured by a horizontal-float film balance enclosed in a constant temperature chamber as previously described.' Lipid crystals were added directly to the surface of the film balance, and the surface pressure was monitored until values were constant with time for 30 min. To establish that equilibrium had been attained, the surface area was reduced, causing a transient increase in surface pressure followed immediately by a spontaneous decrease to the equilibrium value. Conversely, an increase in film area resulted in a temporary decrease of the surface pressure followed by an increase to the equilibrium value.The pure water surface of the film balance was examined for the presence of lipid by measuring the surface pressure at the end of the experiment when all the lipid had been removed from the film side. The resulting surface pressure usually was near zero within the experimental error of the method (*0.3 dyn cm-I). The temperature was monitored by a thermistor placed in the water surface of the film balance; temperature control was within *0.2 "C.N. L. Gershfeld, W. F. Stevens Jr and R. J. Nossal 21 Microscopy The morphology of the DMPG dispersions was examined with a phase-contrast micro- scope (Zetopan, A/O Reichert, Buffalo, NY) equipped with a constant-temperature stage (Cambion, Cambridge, MA).Temperature control was within *O.l "C; however, because ambient temperatures fluctuated, the precision was *0.5 "C. The temperature of the stage was calibrated with pure crystals having melting points in the temperature range of interest. Morphological states of DMPG were examined by adding a small amount of crystal to a clean slide, then adding ca. 0.1 cm3 of water or lipid dispersion of a known concentration to the slide, and sealing a coverslip to the slide with silicone stopcock grease. Quasielastic Light Scattering The movement of 0.45 pm diameter polystyrene latex beads (Polysciences Inc., Warren- ton, PA), suspended in lipid dispersions, was probed by quasielastic light scattering (A = 633 nm). The spectrometer has been described e1~ewhere.l~ An effective diffusion coefficient, Deff, of the beads moving wihhin the dispersion was obtained as Deff = ( ~ T ' / ~ Q ~ ) - ' , where T ~ / ~ is the time necessary for the temporally varying portion of the photon autocorrelation function to fall to l / e times its maximum value and Q is the magnitude of the Bragg scattering vector.The autocorrelation function was determined as a function of temperature. Constant sample temperature was achieved by using a water-jacketted cuvette (type 50, 10 mm light path, Precision Cells, Hicksville, NY) that was surrounded by an air bath kept at a temperature close to that of the cuvette. Sample temperatures were controlled to kO.1 "C, with a precision of k0.5 "C. The data reported here were taken at a scattering angle, corrected for refraction, of 22.1'.A 5.0 mg cm-3 DMPG dispersion was prepared by mixing dried lipid and water at room temperature on a mechanical rotator for 1 h. Dilutions were made by adding an aliquot of the dispersion to water which already contained suspended beads. The diluted systems then either were vortexed vigorously for ca. 1 min or heated at 35 'C for 12 h. Photon autocorrelation measurements of both preparations yielded similar results. When vortexing or heating was not used, the properties of the diluted samples changed over a 24h period, gradually approaching those of the vortexed or heated samples. We generally used the more rapid methods of sample preparation to avoid possible complica- tions arising from DMPG hydrolysis. The concentration of latex beads was 6 x g ~ m - ~ .Sodium azide (2 x lo-* mol dm-3), which had no effect on results, was added to inhibit growth of micro-organisms. Differential Scanning Calorimetry A Perkin-Elmer DSC model I1 scanning calorimeter was used to evaluate the temperature and latent heat of the gel-liquid-crystal transition. A known amount of anhydrous lipid (ca. 1 mg) and water, in the ratio of 1-2 times the amount of lipid, was added to sample pans and the pans were sealed. The observed values of Tm and AHm were independent of the scan rate in the range 1.25-5 K min-'. Results Surface Pressure-Temperature Phase Diagram for DMPG In previous studies of lecithin bilayer dispersions7-' surface bilayers were observed, by radiotracer measurements, to form at the same temperature where a maximum occurs in the surface pressure us.temperature relation. For each system studied, surface pressure measurements showed an abrupt rise followed by a gradual decline. Thermodynamic22 Pospholipid Bilayer Assembly LO 30 2 0 10 0 1 I 20 30 LO 50 TI "C Fig. 1. Equilibrium surface pressure Il,, as a function of temperature T for DMPG-H20 disper- sions. Surface bilayer forms at T* = 31.5 k0.5 "C, the temperature where II, is a maximum. arguments indicate' that two experimental conditions are sufficient to establish the temperature, T*, of surface bilayer formation: ( a ) a maximum in the surface pressure- temperature phase diagram and ( b ) the presence of bilayers in the dispersed phase. In fig. 1 we demonstrate that DMPG has characteristics of surface pressure vs.temperature similar to those observed for DMPC and other lipids:',' DMPG shows a surface pressure maximum at 31.5 f 0.5 "C. Also, in agreement with other reports," we find from diff eren- tial scanning calorimetry of DMPG dispersions that the gel-liquid-crystal transition occurs at T, = 24 "C, and AH = 7 kcal mol-'.* Thus, at the temperature of the surface pressure maximum, 31.5 "C, the DMPG dispersion consists of the lamellar Iiquid- crystalline phase. Since the necessary conditions for surface bilayer formation have been met, the surface pressure maximum is the temperature of surface bilayer formation, T*. Morphological States of DMPG Dispersions Using phase-contrast microscopy, at temperatures near the gel-liquid-crystal transition (24 "C) large crystals of DMPG disappeared when covered with water.However, upon heating the slide to temperatures above 30"C, amorphous structures were seen. The structures became clearer with further heating until at ca. 32 "C a multitude of vesicles seemed to explode in the microscope field. The vesicles were 5-50 pm in diameter, with smaller sizes predominating. The vesicles also exhibited a distribution of wall thick- nesses, from very thin undulating walls to fairly thick walls. The phenomenon is * 1 cal=4.184 J.N. L. Gershfeld, W. F. Stevens Jr and R. J. Nossal 23 reversible, for upon cooling below 30 "C all structures disappeared, but reappeared upon reheating to 32°C. The transformations occur within 5-10 min after changing tem- peratures.Since the interbilayer spacing increases upon dilution of ionized lipids like DMPG," we also examined the effect of dispersion concentration on this striking phenomenon of vesicle formation. With concentrations of 5.0, 1 .O, and 0.25 mg cm-', pronounced Tyndall scattering occurred when a beam of light was passed through the dispersion. However, under phase-contrast microscopy no structures were observed; the particles in the dispersion were too small to be seen in the light microscope. We thought that by focussing on a crystal large enough to be seen in the microscope, but too small to change significantly the concentration of the dispersion, we would be able to study the influence of lipid concentration on the morphology of the vesicles formed by the crystal.Small crystals of DMPG were deposited by evaporating 5.0 mm3 of the 0.25 mg cm-3 dispersion on a microscope slide; they were then covered with 0.1 cm-' of the dispersion. In addition, a small number of glass beads (5 p m diameter) was added to each of the dispersions. The glass beads served two functions: to facilitate focussing on the crystal and to speed formation of the vesicles. The resulting observations were independent of the number of beads present. Very small crystals in each of the three dispersions yielded essentially the same results as large crystals in water: below 30 "C no structures were visible in the microscope, even at a magnification of 640x; at 32 "C extensive vesicle formation was observed. In addition, vesicles were seen occasionally at 30°C in perhaps one of every five slides examined, and with an increase in temperature the number of vesicles grew exponentially.The temperature interval (30-32°C) where vesicles first appear was the same for all three lipid dispersions (0.25, 1.0 and 5.0 mg cm-'). However, with increasing concentra- tion the proportion of thick-walled vesicles to thin-walled vesicles increased. At the lowest dispersion concentration (0.25 mg cm-') the majority of the vesicles were very thin-walled and difficult to see. The vesicle wall thickness also increased when the slide was heated above 32°C. The thickening of the walls above 32 "C was gradual, and became more obvious when the temperature was raised by ca. 5-10 "C. Thus thin-walled vesicles appear to form optimally at 32 "C, which, within experimental error, is the temperature where surface bilayers form.Quasielastic Light Scattering The appearance of vesicles in DMPG dispersions at 32 "C, and the absence of any visible structure below 30"C, suggested that a phase transition occurs in this temperature interval. To examine the nature of the lipid phase at the lower temperatures, dynamic light scattering studies were performed on dispersions of DMPG. Initial results with 1 .O and 0.25 mg cm dispersions yielded very low levels of photon scattering. Polystyrene latex beads thus were added to probe for structural transformations of the dispersed lipid. Although scattering from the beads is only an indirect measure of lipid structure, it enabled us to observe a tramformation in the dispersion.Results are shown in fig. 2, where DeE us. temperature is plotted for a 0.25 mg cm-' DMPG dispersion containing 0.45 p m beads. The measured diffusion coefficients for beads moving in lipid-free water follow closely the dotted line representing values calculated by the Stokes-Einstein relation, D = kT/6.srr)a7 for beads of this size. ( k is Boltzmann's constant, T is the absolute temperature, a is the bead radius and r) is the viscosity of the solution). In the lipid dispersions, for T > 32 "C (T"), values of Defi are equal to those for beads which move freely in water. However, for T < T" the beads move more slowly than if they were in water alone, and Defi appears to be independent of temperature. At temperatures above T" the autocorrelation function approximates24 6 - 4.Pospholipid Bilayer Assembly I I 1 I I I , 1 , - I I , I , I , , I 6 a single exponential. Below T* a long, slowly decaying 'tail' is clearly evident in the autocorrelation function. The data shown in fig. 2 were obtained by alternating between temperatures above and below T*, and we thus conclude that the transformation is reversible with respect to temperature. The data are consistent with the presence of an extended lipid lattice within the dispersion when T < T". Additional evidence of an extended spatial structure was obtained by inserting a concentrated droplet of 0.45 p m beads into a sample of the dispersion. The high local concentration of beads was easily viewed by eye. When left overnight at room temperature, the beads remained localized near the point of insertion.A control which contained water and beads, but no lipid, initially showed the same pattern as did the DMPG dispersion. However, after overnight incubation the beads in water had diffused throughout the entire volume of the sample. In samples maintained at 35 "C beads were uniformly distributed after 6 h whether or not DMPG was present. Changes in DMPG concentration also have a marked effect on bead movement. Generally, the higher the concentration, the greater the inhibition of motion for T < T*. In samples of 5 mg cmP3 DMPG, which were noticeably slurry-like when viewed in a test tube, bead movement was almost totally inhibited and the autocorrelation functions were almost flat out to times of 200ms. Solubility Fig.3 gives solubility of DMPG, X,, as a function of temperature, where X , is the mole fraction of DMPG in solution. Since In X , is proportional to the chemical potential, it will be discontinuous at the gel-liquid-crystal transition temperature, which for DMPG is 24 "C. Line AB was drawn by a linear regression analysis of the data obtained below 25 "C; the slope of AB is 0.123 K-', with the correlation coefficient r = 0.998. For temperatures above 25 "C solubility is not linear with temperature; line BC was therefore drawn as an asymptote to be consistent with the solubility data. The lines AB and BC are discontinuous at 24 "C and yield the calorimetrically determined latent heat forN. L. Gershfeld, W. E Stevens Jr and R. J. Nossal 25 - 1 1 -12 -1 3 -14 - -15 -16 - 17 s u r f ace bil a yer - A I I I I I I I I 10 20 30 LO T / "C Fig.3. Solubility of DMPG in HzO, as a function of temperature T. X, is the mole fraction of DMPG in solution. Line BC has been drawn as the asymptote to the point B of the curve BD (see text). gel-liquid-crystal transition (see Discussion).* We have already noted that longer centrifugation times are required to separate the lipid above 25 "C (see Methods); this is to be expected since phospholipid densities generally decrease upon heating above the gel-liquid-crystal transition. At temperatures above 36 "C the solubility rises steeply with temperature; the solutions, free of excess lipid dispersion, exhibit Tyndall scattering, suggesting that micelles are present in the solutions.However, for solutions prepared at temperatures near T, (the gel-liquid-crystal transition) Tyndall scattering is not evident. Discussion Spontaneous Formation of Unilamellar Vesicles in DMPG Dispersions The principal objective of this study was to establish the equilibrium conditions where single bilayers form in DMPG dispersions. Thermodynamic properties of the equilibrium air-water surface provide an experimental framework for identifying the conditions required to form single bilayers in the bulk dispersion. Two aspects of the surface properties of phospholipid dispersions are of particular importance. The first is the occurrence of a surface pressure maximum at the temperature where surface bilayers f o r ~ n . ~ - ~ From thermodynamic arguments we deduced','' that unilamellar vesicles form in the dispersion at the temperature of surface bilayer forma- tion.The analysis may be summarized briefly as follows. From the Gibbs adsorption relation and the Gibbs-Duhem equation, the surface and bulk lipid states have identical composition when the surface pressure is a maximum; in addition, the partial molar * The discontinuity actually appears at ca. 24.5 "C. A slightly higher T,,, for the centrifuged material is consistent with the elevated pressures generated in the centrifuge tube.26 Pospholipid Bilayer Assembly entropy of each component in the surface film is equal to the corresponding partial molar entropy in the bulk liquid-crystal phase.8 Therefore, the surface and bulk lipid states are identical when the temperature is that of the surface pressure maximum.If the dispersed lipid contains bilayers, as DMPG dispersions do, then the surface also must be in a bilayer state. However, the measured film density is that of a single bila~er;'.~ consequently the equilibrium state in the dispersion must be unilamellar. The second aspect concerns the nature of the transformations leading to the surface bilayer state. The surface bilayer forms only at a unique temperature, and the transforma- tions leading to the surface bilayer resemble higher-order transition^.^"^ The analogous process for forming single bilayers in the dispersion would, therefore, be expected to occur over a range of temperatures. For DMPG dispersions the temperature of surface bilayer formation is T*= 31.5 f 0.5 "C, where the surface pressure-temperature phase diagram shows a maximum (fig.1). Within experimental error, this is the temperature where thin-walled vesicles are found in large numbers by phase-contrast microscopy (see Results). Moreover, the transformation to vesicles appears to be higher order, commencing with the appearance of just a few vesicles at 30 "C, and with the thin-walled vesicle population increasing rapidly to a maximum at T". The motion of polystyrene latex beads within the dispersion also reflects the formation of vesicles. At temperatures below 30°C the dispersion is a loose jelly-like substance which retards the movement of the beads (fig. 2). At 30 "C the movement of the beads increases until at T* it becomes identical to the movement of beads in pure water; T" is the temperature where vesicles are first seen in large numbers.The transformation from the jelly-like state to vesicles occurs over a 2 "C range, characteristic of higher-order transitions. Because the movement of the beads is impeded at T < T* the few vesicles that are observed at those temperatures are likely to be attached to the jelly-like matrix and, from the perspective of thermodynamics, they cannot be considered as isolated unilamellar vesicles. The beads move freely at T* when the jelly-like matrix vanishes; it is only at this temperature that isolated thin-walled vesicles form. But are these vesicles unilamellar? The microscopy indicates that, with dilution of the dispersion, the vesicles appear to become predominantly thin-walled at T"; indeed, the vesicle walls at this temperature are the thinnest we have observed, and are extremely difficult to bring into focus.Since X-ray diffraction studies indicate increasing separation of ionized bilayers with dilution,'* it is reasonable to expect that, for the very dilute dispersions examined here, unilamellar structures form at T*. The formation of uni- lamellar vesicles at 32 "C therefore is directly analogous to the formation of surface bilayers; the surface and bulk states occur at the same temperature. Thus our present experimental results support the thermodynamic proof of the correspondence between surface bilayer formation and unilamellar vesicle formation at T*.*>'O An important feature of the transformation from the jelly-like material to unilamellar vesicles is that it occurs at a unique temperature, T".Indeed, all lipid states which exist above and below T" jelly, multilamellar vesicles (liposomes) and surface films at T" into the single bilayer state. This property is characteristic of a critical point. The presence of multilamellar bilayers along with unilamellar vesicles in phospholipid dispersions may now be attributed to lack of adequate temperature control. When dispersions are prepared, they generally are heated to some ill-defined temperature above the gel-liquid-crystal transition temperature. If the critical temperature T" is reached unilamellar vesicles will form, but multilamellar structures also will form if the temperature is not maintained at the critical point.Phase Diagram of DMPG Bilayer Assembly A schematic phase diagram summarizing the various bilayer states formed in DMPG dispersions is given in fig. 4. There are three types of organization: the solution phaseN. L. Gershfeld, W. F. Stevens Jr and R. J. Nossal D I B I L A Y E R GEL L l Q U l D CRYSTAL - J E L L Y u LV + T m c e l l e _ _ _ _ _ - - - - M m o n o m e r 27 Fig. 4. Schematic phase diagram for dilute DMPG-H20 dispersions. Above line ABCD, the lipid in the dispersion consists of bilayers. At T,, gel and liquid-crystal bilayers coexist in a jelly-like matrix. At T*, the matrix transforms to unilamellar vesicles (ULV); at T > T* the unilamellar vesicles become multilamellar (MLV). Below line ABCD, DMPG exists in solution. Micelles are expected to form above the line KM, where K is the Krafft point.containing monomer or micelle, the two bilayer phases (gel and liquid crystal) and the three morphologically distinct bilayer states belly, unilamellar vesicle (ULV) and multi- lamellar vesicle (MLV)]. In fig. 4, lines AB and BC show the temperature dependence of the solubilities of the gel and liquid-crystal bilayers. To demonstrate the self-consistency of the solubility data we calculate the latent heat of the gel-liquid-crystal transition from the enthalpies of bilayer dissolution ( i e . the negative of bilayer assembly) near T,. The standard free energy of bilayer assembly may be written as AGi = RT In X , +f( T, +) ( 1 ) where the first term on the right accounts for the assembly of uncharged bilayer from monomer, and where f( T, +) is a term which accounts for the work of charging the bilayer to a potential +.In principle, the Gibbs-Helmholtz relation applied to. an eqn ( 1 ) yields the change in enthalpy for assembly of bilayer from monomer in solution. While the expression f( T, +) cannot be determined from our data, we assume that the temperature dependence off( T, +) is the same below and above T,. Consequently, we do not need the exact form of f( T, +) in order to determine the latent heat of the gel-liquid-crystal transition. From the data in fig. 3 we find that AHm = 7 kcal, in agreement with the calorimetric measurements. Implicit in eqn ( 1 ) is the assumption that at Tm the solution contains monomer and not micelles. This is based on the observation that solutions of soaps and detergents, in general, form micelles only when the Krafft point is exceeded.16 The Krafft point is the temperature where the solubility rises sharply, and is usually higher than the melting point of the lipid; in the case of DMPG the melting point is T, = 24 "C, and the solubility28 Pospholipid Bilayer Assembly first increases noticeably at ca.36 "C (fig. 3). Moreover, our preliminary observations that DMPG solutions above 36 "C visibly scatter light, while solutions near T, do not, also support the assumption that the solutions contain monomers. We have indicated the Krafft point as K in fig. 4, with the dotted line KM as the temperature dependence of the critical micelle concentration. While the precise value for the Krafft point has not been obtained in this study, we have indicated its probable location at T > T".The jelly-like material which transforms at T* to unilamellar vesicles, ULV, represents a three-dimensional matrix which we believe consists of sheets of single bilayers that interact weakly to form an extended three-dimensional structure. Our inability to see any structures under phase-contrast microscopy is consistent with this model. The transformation from the jelly to the vesicle is higher-order, and therefore will not display a latent heat. Thus the solubility changes continuously with temperature at T", the critical point for single bilayer formation. With heating, the unilamellar vesicles form multilamellar vesicles, MLV; this transformation is also higher order, and without a latent heat.Is the formation of unilamellar vesicles at T" a general property of phospholipids? While details of the phase diagram may differ for specific lipids, surface bilayer formation has been demonstrated with a wide range of phospholipids, including various phos- phatidylcholines' and lipid extracts of erythrocytes and bacterial membranes.1° The surface-thermodynamic argument we have used for DMPG dispersions is general, and therefore each of the foregoing lipid systems is expected to form unilamellar vesicles at the temperature of surface bilayer formation. Of particular significance is the observa- tion that T" for erythrocyte and bacterial membrane lipids is the same as the growth temperature of the cell. lo The unilamellar vesicle, an equilibrium state governed only by the conditions of T and composition, will form whenever conditions are appropriate, whether in artificial dispersions or in cell membranes. Thus the spontaneous assembly process manifested for DMPG unilamellar vesicles is likely to be important in the in vivo assembly of membrane bilayers. References 1 I. Kahane and S. Razin, Biochim. Biophys. Acta, 1969, 183, 79. 2 L. Mindich, J. Mol. Biol., 1970, 49, 433. 3 R. M. Servuss, W. Harbich and W. Helfrich, Biochim. Biophys. Acta, 1976, 436, 900. 4 C. Huang, Biochemistry, 1969, 8, 344. 5 S. Batzri and E. D. Korn, Biochim. Biophys. Acta, 1973, 298, 1015. 6 N. E. Gabriel and M. F. Roberts, Biochemistry, 1984, 23, 4011. 7 N. L. Gershfeld and K. Tajima, Nature (London), 1979, 279, 708. 8 K. Tajima and N. L. Gershfeld, Biophys. J., 1985, 47, 203. 9 L. Ginsberg and N. L. Gershfeld, Biophys. J., 1985, 47, 211. 10 N. L. Gershfeld, Biophys. J., 1985, 47, 426a. 1 1 E. J. Findlay and P. G. Barton, Biochemistry, 1978, 17, 2400. 12 M. E. Loosley-Millman, R. P. Rand and V. A. Parsegian, Biophys. J., 1982, 40, 221. 13 G. Rouser, S. Fleischer and A. Yamamoto, Lipids, 1970, 5, 494. 14 R. Nossal and S. H. Chen, J. Phys. (Paris) Suppl., 1972 33~1, 171. 15 N. L. Gershfeld, in Cell Surface Dynamics, ed. A. S. Perelson, C. DeLisi and F. W. Wiegel (Marcel 16 K. Shinoda, T. Nakagawa, B. Tamamushi and T. Isemura, Colloidal Surfactants (Academic Press, New Dekker, New York, 1984), chap. 4, pp. 111-114. York, 1963), pp. 7 and 8. Received 19th December, 1985

ISSN:0301-7249    

DOI:10.1039/DC9868100019

出版商:RSC    

年代:1986

数据来源: RSC

摘要:

Faraday Discuss. Chem. SOC. 1986, 81, 29-37 Directly Measured Deformation Energy of Phospholipid HI, Hexagonal Phases Sol M. Gruner Physics Department, Princeton University, Princeton, New Jersey 08544, U.S.A. V. Adrian Parsegian" National Institutes of Health, Bethesda, Maryland 20892, U.S. A. R. Peter Rand Biological Sciences, Brock University, St. Catharines, Ontario L2S 3AI, Canada Osmotic stress combined with X-ray diffraction has been used to measure the energies of inverted hexagonal (HI*) phases containing dioleylphos- phatidyl-choline (DOPC) and -ethanolamine (DOPE). To distinguish chain packing energies from the action of other factors governing stability, energies with and without added tetradecane have been compared. Little hydrocarbon strain energy is stored in the HII lattice, but alkane promotes the transition from a lamellar to an HI1 structure.In the presence of excess water, various HII lattices seek spontaneous radii of curvature, r,, of the aqueous-polar interface. Under low osmotic stress, the work of deforming from this spon- taneous radius appears to be quadratic in curvature, can be well described by a bending modulus that is independent of the initial r,, and is related to bending moduli measured on planar bilyaers. At high stress, the work of water removal and lattice deformation seems to be that of removing water from a polar 'mash' composed of lipid headgroups and water, dependent only weakly on the DOPE/DOPC ratio. The inverted hexagonal phase ( HII) '-9 formed by many phospholipids in water has been relatively neglected compared to lamellar phases.The lack of attention is puzzling given the many instances where the formation of this structure of water pores is invoked to explain membrane fusion This paper reports the first direct measurements of the work of deforming a non-lamellar phase, accomplished by the simultaneous application of X-ray diffraction and osmotic stress measurements to map the phase diagram and to find both the underlying phase structures and the free energies of their formation. It has long been known that phosphatidylcholines (PC) and phosphatidyl- ethanolamines (PE) form lamellar ( L a ) and inverted hexagonal ( HII) liquid crystals, which exhibit a limited affinity for water.' Beyond some level of hydration the lattice dimensions remain fixed. The water affinity of the La phase is known to arise from a competition between strongly repulsive, short-range, exponentially decaying hydration force^.'^ modified by mechanical fluctuation^,'^"^ and weaker, algebraically decaying, attractive van der Waals forces.At the point of aqueous excess, these forces are in balance and the system is in stress-free equilibrium with liquid water. If the chemical activity of the water is now decreased by osmotic tress,'^,'^-^^ the lipid-water system must accordingly adjust its lattice parameters to bring the system into equilibrium with the new chemical potential of the water. Osmotic stress may be used to measure the work of removing water from between the bilayers. A plot of osmotic pressure vs. lattice water volume is actually a P-V diagram whose integral is the work done to extract 2930 HII Phase Deformation Energy water from the lattice.The osmotic stress applied to a lamellar lattice is then the force/area between facing bilayers. A very different set of forces has been suggested to explain the stability of HII phase^.^-^ The lipid aggregates which form the cylindrical water-containing tubes are thought to have a spontaneous radius of curvature, r, (also known as an equilibrium or intrinsic radius of curvature), characteristic of the mixture of component lipids. The work of bending the layers to a new radius, r, might be written as The finite length of the lipid chains appears also to set limits on the diameter of the HII tubes and may be expected to exert forces within the lattice. In the present study we have measured the work needed to remove water from HII structures of pure dioleylphosphatidylethanolamine (DOPE) and of a 3 : 1 DOPE-DOPC mixture with and without tetradecane added to relax strains in the hydrocarbon region.While alkane added to the 3 : 1 DOPE-DOPC mixture is needed to form the Hll structure at room temperature, it appears to have no effect on the stresses encountered in pure DOPE HII lattices. The measured work of deformation near ro can indeed be charac- terised by a bending modulus, whose value is close to that measured in planar bilayers.22-26 Surprisingly, at high osmotic stress the work of dehydration seems to depend only weakly on the PE/PC ratio. Methods The lipids, DOPE, DOPC and tetradecane, were combined in the desired ratios and then hydrated either gravimetrically at known weight ratios or by equilibration with polyethyleneglycol solutions of measured osmotic pressure or with the vapours of saturated salt solutions of known vapour pressure.We investigated the structural responses of the HII phase to applied osmotic or vapour pressure stress in exactly the same way as we did for many multilamellar phases . 5720,2 First, the structures formed by the lipid-water mixtures of measured composition were characterized by X-ray diffraction. All samples discussed here formed a single two-dimensional hexagonal phase characterized by a series of X-ray spacings bearing ratios to the fundamental repeat, d, of l , l / f i , 1/2, l/d7, 1/3, l/v"ii,. . .. From the known weight percent lipid, c, the unit cell was divided into two compartments as shown in fig. 1, one containing all the water in the form of aqueous cylinders of radius r situated on the six-fold axes of symmetry distance d (axis) apart, and another containing all the lipid and filling the rest of the unit cell. (Note that in all our stated dimensions we use the Luzzati2 volume-average definition for lipid-water boundaries, where the lipid-water interface is drawn to divide all water from all lipid.) One should recognize, though, that in fact there is a polar region containing both lipid and water [see, e.g. ref. (19)] and that at the lower radii of curvature there may not be any purely aqueous regions. We define s = d(axis) = 2 d / d taking phospholipid and water densities as equal.(In the actual reduction of data we have corrected dimensions for the weight density of components using 1 .O for the specific volume of water and phospholipid molecules and 1.2 for the specific volume of added alkane.) Then the intercylinder distance d, = s - 2r. The area on the surface of the water cylinder available to each phospholipid molecule isS. M. Gruner, V. A. Parsegian and R. P. Rand 31 Fig. 1. Parameters used to describe the inverted hexagonal phase. where v, is the volume of a weight-averaged lipid molecule and its complement of hydrocarbon. One important dimension is d,,,, the longest length the lipid molecule has to stretch in forming the hexagonal phase. Samples X-rayed under the stress of osmotic or vapour pressure yielded the funda- mental hexagonal repeat distance d.Toheir composition and structural parameters were derived by referring to the equivalent spacing of the gravimetrically prepared samples. Results We describe results obtained for the hexagonal phases formed by pure DOPE (PE), DOPE plus 5 wt% tetradecane (PE-TD) and DOPE-DOPC mole ratio 3 : 1 plus 20 wt% tetradecane (PE-PC-TD). We omit from this paper the many instances of the coexistence of lamellar and hexagonal structures that result after several days' equilibration with different PE-PC-TD ratios. DOPE-DOPC 3 : 1 without TD forms only a lamellar phase. At 20% TD this mixture shows only the HII phase at 25 "C and at all osmotic stresses applied over extended times. Fig. 2 shows the dependence of the radius of the aqueous cylinders on the osmotic stress (plotted as log P ) applied to the lipid phase.The PE and PE-TD data sets32 HI, Phase Deformation Energy X. xo 0 0 X. R X. * (a) 77 \ \ (b) \ \ I \ \ I I I I I I I I I I 0 0 $ 0 0 a 0 0 10 20 30 10 20 30 rw,,,*/A rw,t,r/ A Fig. 2. Osmotic stress, as log P us. pore radius for three different lipid mixtures. ( a ) 0, Pure DOPE, X, DOPE plus 5% tetradecane; (b) DOPE-DOPC 3: 1 plus 20% tetradecane, with the dashed line for comparing the data from (a). [fig. 2(a)] coincide over the whole pressure range and d,,, remains constant. On the other hand, while all three materials give strikingly similar results at the high pressure, more dehydrated, end of the phase diagram, the PC-containing HII phase [fig. 2(b)] swells to larger dimensions at low pressures.In the stress-free state ( P = 0, corresponding to pure water), it takes on an equilibrium radius and aqueous volume far larger than do the other two sets. The dependence of the aqueous volume per mass of polar group on applied stress for these three data sets is shown in fig. 3. (This is an aqueous volume per average mass of polar group, taken as molecular weight 213 for pure DOPE and molecular weight 255 for DOPC to give an average molecular weight 223.5 for the 3 : 1 PE-PC mixture.) Note that at sufficiently high osmotic stress, all three systems tend toward the same variation in the applied osmotic pressure to the water/lipid polar mass ratio. In the analysis that follows, we attribute this behaviour to the work of dehydration of a solution of polar groups as solutes independent of their kind or arrangement.Without tetradecane, the energetic cost of stretching chains to a length d,,, prevents the formation of HI1 phases with large diameter lipid tube^.^-^ In the presence of tetradecane this hindrance is removed and the spontaneous curvature of the lipid monolayers is allowed to drive the formation of HII structures with large water cores. Remarkably then, the fully hydrated PC-containing hexagonal phase achieves water per polar group far in excess of the others. Analysis and Discussion We have chosen to look at the stability of the structures measured here in terms of tension among three conflicting factors: the drive for a spontaneous radius of curvatureS. M. Gruner, V.A. Parsegian and R. P. Rand 33 E I TE C C 6 C K E l C C C C I I I 0 500 1000 (water volume/mass)/A3 Fig. 3. Osmotic stress us. volume of water per average mass lipid polar group. Volumes are normalized to the mass of pure DOPE polar groups. E, pure DOPE; T, DOPE + 5% tetradecane; C, DOPE : DOPC 3 : 1 plus 20% tetradecane. ro of the phospholipid monolayer, the work of progressive dehydration, and the obviation of stress in hydrocarbon regions by the addition of alkanes. Acyl Chain Stress This is the simplest to describe. Added tetradecane has no significant effect on the dimensions of those inverted hexagonal phases that already form in its absence (fig. 2). Neutron diffraction of DOPE mixed with deuterated alkane (Gruner and Huang, in preparation) demonstrates clearly that at least some and probably all of the alkane does enter the HII lattice. The clear implication is that the entry of alkane does not significantly relax stress in a pre-existing HI* structure.This point has been anticipated in earlier measurements of DOPE-dodecane mixtures' where the addition of alkane caused only slight differences in the Bragg spacings of HII lattices. It has been suggested8,' that many PE-PC mixtures normally form La phases because of a prohibitively high cost of stretching acyl chains to fill the hydrocarbon volume of an HI, phase of the proper curvature. Added alkane induces PE-PC mixtures to form HII phases of very large lattice size. Addition of the PC component apparently shifts the spontaneous radius to a correspondingly large value requiring acyl chains to stretch to energetically prohibitive lengths.Added alkane removes this constraint. In the pure DOPE HII phase studied here d,,, is a constant 20.3 A at all hydrations; the addition of 5% tetradecane results in d,,, = 21.2 A, again stable with varying hydration. However, in the PE-PC 3 : 1 mixture, d,,, varies between 22.12 and 23.7 A. This is close to the maximum extent of oleate chains plus polar groups, presumably not easily spanned in the liquid chain state (R. Pastor, personal communi- cat ion).34 HI, Phase Deformation Energy These data suggest to us that the chain free energy rises steeply with chain stretch at the dimensions and lateral pressures in these structures. This free energy cost largely determines whether a curved structure, such as in an HII phase, can occur.Recent theoretical models of lipid chains seem to come to similar conclusions.27~28 Spontaneous Curvature The concept of a bilayer spontaneous curvature has been used to explain the elastic bending of b i l a y e r ~ . ~ ~ , ~ ~ , ~ ~ ~ ~ ~ Following Kirk et aZ.; we choose to define a spontaneous curvature of the individual monomolecular lipid layers which, back to back, form bilayers (ignoring back-to-back interaction) or form HrI phases when rolled into cylinders. We postulate, then, that the lipid monolayers of an osmotically stress-free HrI phase assume a curvature of radius r, for its aqueous region and that deviations in dimension encounter a quadratically varying free energy g(r) = WO/2)[(1/~) - (1/r0)I2.Differentiating this free energy per phospholipid molecule with respect to the volume v of water per phospholipid molecule, we can write the osmotic pressure relation for the Hrr phase in the vicinity of ro P = -dg/dv 1 - (dg/ d r ) / (d U/ d r ) where A is the area per phospholipid at the polar interface. Not only does this expression allow direct comparison with data, but it is prefaced by a factor, 2Ko/A, that should be the bending modulus K , of a b i l a ~ e r . ~ (To see this, recall that KO is normalized to the energy per molecule. Division by Ao, the area per molecule at ro, converts this to a modulus per unit area. The factor of two includes contributions from both sides of a bilayer, each of whose faces should be subject to equal energy of deformation if the quadratic form has meaning and r, is infinite.This relation ignores any back-to-back monolayer interaction within the bilayer.) By plotting r2P us. l / r we find that this relation fits fairly well in the vicinity of r,, but predicts too high an energy at smaller radii. The system goes to even higher curvatures than predicted from a purely quadratic form for the energy in the vicinity of its minimum at r,. The values of KO, K , and ro so obtained, as well as the range of radii over which the quadratic energy minimum seems to hold, are listed in table 1. One cannot help but be struck by the consistency among K , despite differences in KO and ro, as well as the fact that the range of 3 to 8 kT is just at the lower end of the range of bending moduli measured in planar PC bilayers.22-26 Hydration Energy Following Marcelja and Radic3' and Kirk et aL7 we write the hydration energy per unit length of a water cylinder of radius r in the form G(r) = gs(2.rrr) w l ( r / A ) / 1 o ( ~ / A ) l where g, is related to the free-energy density of water at the cylinder surface, A is a characteristic length of decay of the perturbation of the water by the surface (2-3 A inS.M. Gruner, V. A. Parsegian and R. P. Rand 35 Table 1. Bending moduli and spontaneous curvatures case Kc/ kT ro/ A r rangea / A PE PE-TD PE-PC-TD 4.6 f 2.5 5.6 f 2.2 5.8 f 1.5 23.8zt2.4 22.5 f 1.1 29.3 f 1.7 22-25.8 15-23.1 20-30 a Range of r over which the data was fitted as r2P vs. 1/ r. lamellar ~ystems’~,~*) and I are Bessel functions of the first kind.Since (r/ A ) is typically much greater than one, we expand the Bessel functions to derive G(r) + g, ( 2 7 4 w - [ A m r)H. If one divides G(r) by 27r r / A , the number of phospholipid molecules per unit length, one may write the corresponding free energy per molecule g(r, A) = gsAW - [ M 2 r)Ih It is immediately clear from either of the last two expressions that this kind of hydration energy has a very weak explicit dependence on the dimensions of the water cylinder. Similarly for its contribution to the osmotic pressure taken as the rate of change of energy with respect to volume, -dg( r, A)/dv, or dG( r)/d( m2). Again, for r much greater than lambda Po,,= [1/(27rr)] dG/dr=g,(A/r). The observed dependence of Po,, on r between 7 and 18 A is nearer to an exponential than to a reciprocal dependence on radius. If a hydration energy of the type used here is the cause of the observed osmotic pressure, then it must be through a change in the coefficient g, with change in radius or, more likely, with molecular area A.A similar argument seems to dispose of any significant contribution from a constant interfacial energy term of the type Gint( r ) = y27rr per unit length. This will contribute to the osmotic pressure only as - y / r, again much too slow a variation to matter over most of the range of observed radii. Again, from r = 19 8, down to 9 A, pressure changes by more than an order of magnitude, far more than to be expected from a Laplacian pressure. The convergence of all three curves at higher pressures suggests that in this limit the work of dehydration does not depend strongly on differences in polar group.A similar convergence was seen in earlier measurements on lamellar phases where egg PE and egg PC exerted comparable pressures at low hydration.32 In fact, the highest pressures reported in fig. 3 are very close to what one sees in the lamellar DOPC multilayers at comparable water contents. What might one make of the remarkable overlap in the data for osmotic stress vs. the volume of water per lipid molecule? It appears that at high osmotic stress the work of extracting water becomes independent of polar group identity (PE vs. PC) and of the shape of the aqueous-polar group region. Perhaps one might think of the aqueous region as a ‘mash’ of water and polar groups without making a serious effort to seek arbitary lipid-water bondaries. An obvious test of this idea is to examine osmotic pressures of concentrated a-glycerophosphorylcholine and a-glycerophosphoryl- ethanolamine solutions.36 HI, Phase Deformation Energy The unimportance of added alkane at high pressures again suggests that the primary work in deformation is from dehydrating the polar mash in the HII cores rather than in chain packing or stretching. Chain energetics might be so stiff as to prohibit significant chain energies in deformed HI, structures.Chain packing in the presence of alkanes obviously enables the occurrence of HII phases where spontaneous radii of curvature of the monolayer are too large to be accommodated by the phospholipid acyl chains alone.This demand for a large radius results in a water content beyond that needed for polar group hydration in a correspond- ing 3 : 1 PE-PC lamellar phase, suggesting that all the initial work of dehydration goes into forcing the tubes to curve to smaller radii and not into dehydrating headgroups. We note the following: For a quite unrelated system, the voltage-dependent anionic channel from mitochondrial membrane (VDAC or mitochondrial porin), it has been possible to measure the work of channel opening and closure as well as the concomitant change in the internal aqueous volume.33 This was done again using osmotic stress. A channel of 30-40 A diameter34 ‘closes’ to a diameter of some 10-15 A with a work of the order of 10 kT. There, as here in the HII phase, the work of closure is viewed as one of dehydration.If we integrate the pressure us. volume curve of fig. 2 over a similar range of radii, the work of compressing HI1 cylinders comes to ca. 20 kT for the 60 A length of VDAC,34 in surprising qualitative agreement with the ionic channel energy. Conclusions As indicated by the very small effects of added alkane on HI, lattice dimensions, little strain energy is stored in the hydrocarbon chain region of the HII lattice. Still, alkane is absolutely necessary to form certain HII phases. In the presence of excess water and alkane, the aqueous region of the HI, structure has a most favoured or spontaneous radius of curvature. This radius is characterized by a locally quadratic bending energy minimum whose bending modulus is of the same magnitude as the bending moduli measured in planar bilayers.At high osmotic stress, the work of lattice deformation goes largely into removing water from the polar core of the HII tubes. This work appears to depend only weakly on the chemical identity of the two polar groups examined here. These three features of packing and deformation should simplify thinking about phospholipid polymorphism and assembly. This work was supported in part by NIH grant GM32614 and DOE grant DE-AC02- 765V03 to S.M.G. as well as an NSERC grant to R.P.R. References 1 V. Luzzati and F. Husson, J. Cell. Biol., 1962, 12, 207. 2 V. Luzzati, in Biological. Membranes, ed. D. Chapman and D. F. H. Wallach (Academic Press, New 3 D. Marsh and J. M. Seddon, Biochim. Biophys.Acta, 1982, 690, 117. 4 J. M. Seddon, G. Cevc and D. Marsh, Biochemistry, 1983, 22, 1280. 5 J. M. Seddon, G. Cevc, R. D. Kaye and D. Marsh, Biochemistry, 1983, 23, 2634. 6 S. M. Gruner, P. R. Cullis, M. J. Hope and C. P. S. Tilcock, Annu. Rev. Biophys. Biophys. Chem., 1985, 7 G. L. Kirk, S. M. Gruner and D. L. Stein, Biochemistry, 1984, 23, 1093. 8 G. L. Kirk, and S. M. Gruner, J. Phys. (Paris), 1984, 46, 761. 9 S. M. Gruner, Proc. Natl Acad. Sci. USA, 1985, 82, 3665. York, 1968), pp. 71-123. 14, 211. 10 A. J. Verkleij, C. Mombers, W. J. Gerritsen, L. Leunissen-Bijvelt and P. R. Cullis, Biochem. Biophys. 11 S. W. Hui, T. P. Stewart, P. L. Yeagle and A. D. Albert, Arch. Biochem. Biophys., 1981, 207, 227. Acta, 1979, 555, 358.S. M. Gruner, V. A. Parsegian and R.P. Rand 37 12 R. P. Rand, T. S. Reese and R. G. Miller, Nature (London), 1981, 293, 237. 13 D. P. Siegel, Biophys. J., 1984, 45, 399. 14 D. P. Siegel, Biophys. J., 1986, in press. 15 R. P. Rand, R. P., Annu. Reu. Biophys. Bioeng., 1981, 10, 277. 16 W. Helfrich, Z. Naturforsch., Teil A, 1978, 33, 305. 17 E. A. Evans and V. A. Parsegian, Proc. Natl Acad. Sci. USA, in press. 18 D. M. LeNeveu, R. P. Rand and V. A. Parsegian, Nature (London), 1976, 259, 601. 19 D. LeNeveu, R. P. Rand, D. Gingell and V. A. Parsegian, Biophys. J., 1977, 18, 209. 20 V. A. Parsegian, N. L. Fuller and R. P. Rand, Roc. Natl Acad. Sci. USA, 1979, 76, 2750. 21 V. A. Parsegian, R. P. Rand and D. C. Rau, in Methods in Enzymology. Vol, 127. Biomembranes; Protons 22 R. M. Servuss, W. Harbich and W. Helfrich, Biochim. Biophys. Acta, 1976, 436, 900. 23 E. A. Evans and R. Skalak, in Mechanics and Thermodynamics of Biomembranes (CRC Press, Boca 24 M. B. Schneider, J. T. Jenkins and W. W. Webb, Biophys. J., 1984, 45, 891. 25 M. B. Schneider, J. T. Jenkins and W. W. Webb, J. Phys. (Paris), 1984, 45, 1457. 26 H. Engelhardt, H. P. Duwe and E. Sackmann, J. Phys. Lett., 1985, 46, L-395. 27 A. Ben-Shaul, I. Szleifer and W. M. Gelbart, J. Chem. Phys., 1985, 83, 3597; I. Szleifer, A. Ben-Shaul 28 D. W. R. Gruen, J. Phys. Chem., in press. 29 E. A. Evans, Biophys. J., 1974, 14, 923. 30 H. J. Deuling and W. Helfrich, J. Phys. (Paris), 1976, 37, 1355. 31 S. Marcelja and N. Radic, Chem. Phys. Lett., 1976, 42, 129. 32 L. J. Lis, M. McAlister, N. Fuller, R. P. Rand and V. A. Parsegian, Biophys. J., 1982, 37, 657. 33 J. Zimmerberg and V. A. Parsegian, Biophys. J., 1984, 45, 59a. 34 C. A. Mannella, M. Radermacher and J. Frank, in Proc. 42nd Annu. Meeting of Electron Microscopy and Water, Structure and Translocation, ed. L. Packer, (Academic Press, New York, 1986). Raton, Florida, 1980). and W. M. Gelbart, J. Chem. Phys., 1985, 83, 3612. Society ofAmerica, ed. G. W. Bailey (S. F. Press, San Francisco, 1984), pp. 644-645. Received 13th January, 1986

ISSN:0301-7249    

DOI:10.1039/DC9868100029

出版商:RSC    

年代:1986

数据来源: RSC

摘要:

Faraday Discuss. Chem. SOC., 1986,81, 39-48 Peter Fromherz," Carlheinz Rocker and Diether Ruppel Abteilung Biophysik der Universitat Ulm, D- 7900 Ulm-Eselsberg, Federal Republic of Germany The edge energy of a lipid bilayer is considered to be the crucial parameter controlling the formation of closed vesicles. It is considered to be modulated by amphiphiles accumulating at the edge as described by a Gibbs isotherm. The approach is tested for the system egg-lecithin - taurochenodesoxycho- late. The parameters entered into the Gibbs' isotherm are determined by dynamic light scattering in the regime of mixed micelles (vanishing edge tension). Adjustment of an appropriate finite edge tension then leads to the discovery of metastable discs after sonication which close to vesicles spon- taneously.This disc-vesicle transition is observed by electron microscopy. It is described in terms of a phenomenological potential profile. Estimates of the intrinsic edge tension and of the elastic modulus of an open lipid bilayer are obtained. The idea of a 'protocellular vesicle' made up of a fluid assembly of amphiphilic molecules was introduced by Krafft in 1896.' After the proposal of a bimolecular layer of lipid to be the backbone of biomembranes in 19252 it took almost fifty years until the existence of vesicles made up of a single bilayer was demonstrated definitel~.~ A consistent physical picture for their formation and stability, comprising a theoretical concept and systematical experimental tests, is not yet availabie, despite the numerous recipes available for their p r e p a r a t i ~ n .~ It has been attempted to consider vesicles as micellar aggregates, assigning their size and stability to the geometric constraints of the amphi- philic molecules5 and to the thermodynamics of their assembly.6 In that approach the material making up the bilayer is characterized by the average ratio of an optimal area of the hydrophilic headgroups and of the volume occupied by the hydrophobic g r o u p ~ . ~ ~ ~ In the present paper we describe a novel approach to the rationalization of the formation of vesicles. We take into account a partial segregation of two different amphiphiles in a micellar assembly, of a lipid and of a detergent.'-'' Let us consider a finite fragment of a bilayer. We expect that detergent molecules accumulate at the open edge, as their geometry is better matched there than in the bulk of the bilayer (fig.1): the detergent is 'edge-active' with respect to the bilayer. In order to obtain a quantitative description of this effect we chose the thermodynamic approach of Gibbs' isotherm, such that the accumulation at the edge is related to a reduction of the tension of that edge.' Fluid fragments with a finite edge tension tend to lateral fusion, i.e. to coalescence. In addition the edge tension drives these 'two-dimensional droplets' to escape into the third dimension, to collapse to vesicles (fig. 1). Although the feasibility of such a disc-vesicle transition was considered in early studies on vesicle formation,12 the postu- lated transient discs have never been found. The changing packing energy in the rearrangement of lipid from a disc to a vesicle may be described in a phenomenological approach by a modulus of bending elasti~ity,'~.'~ such that the transition is governed by the counterbalance of edge energy and elastic Within such a framework it is not the final vesicular assembly alone which determines its f ~ r m a t i o n ; ~ ' ~ the transient edge plays a crucial role.3940 Disc- Vesicle Transition t Fig. 1. ( a ) Illustration of doping of a finite lipid bilayer by an edge-active amphiphile. The amphiphile is solubilized in the bulk membrane with a binding constant KWM and adsorbs to the edge with a higher binding constant KWE. The accumulation at the edge is related to a reduction of the edge energy according to the Gibbs isotherm.( b ) Illustration of the shape transformation of an isotropic fluid bilayer from a planar disc of radius RD into a spherical vesicle of radius RD/2 with transients shells of radius R. We propose that the dominant role of various amphiphiles in the existence of lipid vesicles is their effect upon the edge, that this effect is due to a modulation of the edge tension and that this modulation may be described by a Gibbs isotherm.’ An additional effect on the packing is not excluded of course. In the present paper we first consider the concept of edge activity, i.e. we derive the relation between the edge tension and the concentration of an edge-active agent. The parameters in this relation are determined by dynamic light scattering for the system egg lecithin-taurochenodesoxycholate for vanishing edge tension.In the regime of finite edge tension the existence of transient discs after sonication is shown by electron microscopy. The spontaneous transformation of these discs to vesicles is demonstrated and rationalized in terms of a phenomenological potential profile. The paper closes with some remarks on possible generalizations of these concepts with respect to vesicle preparation and to the stability of membranes. Edge Activity We consider an aqueous solution of bilayer fragments made of a lipid and of another amphiphile A. The Gibbs free energy G = G ( n A , L ) depends on the number nA of molecules A, on the length L of the edge and on other variables, such as the amount of lipid, the temperature and the electrical field strength.We rgplace nA by the conjugate chemical potential pA, differentiate the Legendre transform G = G - n#A with respect to pA and L and interchange the two differentiation? as usual.’7 We obtain eqn ( l ) , where we have introduced the edge tension as y = (dG/dL)pA.P. Fromherz, C. Rocker and D. Riippel 41 If an agent is to be added with increasing edge length at constant pA7 we call it 'edge-active'. Enhancing pA at constant L leads to a reduction of the edge tension. Of course, eqn (1) also holds for agents which avoid the edge. Those materials enhance the edge tension. We obtain the edge tension y(c,) as an explicit function of the concentration cw of the edge-active agent in water from eqn (1) by integration using two approximations.(i) We relate the change in pA and the change in cw as dpA = kTd(1n c,) (where kT is the thermal energy). (ii) We consider the agent to be added per unit length as an adsorbate at the edge with the density cE = (dnA/dL)pA and apply Langmuir's isotherm cE = CE/ ( 1 + 1/ KWECW) with binding constant K W E and saturation CE: y = yo - CEkT In ( KWECW + 1). (2) The fall in edge tension from the intrinsic value yo with increasing cw is determined by the two parameters EE and KWE. The edge tension vanishes at a concentration c&: c& = K &( exp - - 1). CEkT (3) If the intrinsic edge tension is large with yo >> cEkT we have KWECC >> 1 [eqn (3)], ie. the edge is saturated near y = 0. In that case the edge tension [eqn (2)] is approxi- mated by the logarithmic difference between cw and c&: y = cEkT In ( c&/cw).(4) The vanishing edge tension appears as a compensation of intrinsic edge energy and binding energy according to eqn (5), where we have introduced the standard free energy of local binding as AGkE = -kT In KWE: yo= CE(-AGkE+ kT In c&). (5) The free concentration cw of an edge-active agent is difficult to determine experi- mentally. So we express it by its total concentration cT. Contributing to C, are, in addition to c,, the material solubilized in the bulk membrane and the adsorbate at the edge: The solubilizate is described as a binding equilibrium with the lipid of concentration cL (binding constant KWM). The adsorbate is referred to the total edge length of all monodisperse discs of radius RD making up the total area of the membrane per unit volume (CLuM/2), where uM is the effective area per lipid molecule in a monolayer.(The intrinsic area of the lipid must be corrected for the contribution of the solubilized amphiphile.) Eqn (2) and (6) together express the function y(cT) with the five parameters yo, CE, KWE, KWM and aM for a dispersion characterized by cL and RD. In a dispersion without edge (e.g. vesicles) C, is proportional to c,, as the last term in eqn (6) may be disregarded. J nm-', EE = 4 nm-I, KWE = 15 500 dm3 mol-' (with c& = 0.8 mmol dm-3 and AG& = -28 kJ mol-'), KWM = 600 dm3 mol-I, uM = 1 nm2 for cL = 2.5 mmol dmP3 and RD = 25 nm. The two light lines are drawn using eqn (2) and ( 6 ) , the continuous line referring to closed membranes, the dashed line referring to open discs.(The limit of vanishing edge tension in the variable C, is above c& because of the capacity of bilayer and edge for the edge actant.) The heavy lines are drawn accordingly using the approximation of eqn (4) with c&=O.8 mmoldm-3 and CE=3.7nm-'. The relevance of this simple approximation is apparent. Fig. 2 shows y(cT) for a selected set of parameters with yo= 4.2 x42 Disc- Vesicle Transition 0 1 2 c,/mmol dmP3 3 Fig. 2. Edge tension y of a lipid bilayer (lecithin) as a function of the total concentration C, of an edge-active agent are (taurochenodesoxycholate) at a lipid concentration cL = 2.5 mmol dmP3. The continuous lines 1 and 2 refer to a dispersion without actual edge (closed vesicles), the dashed lines 3 and 4 to monodisperse discs of a radius R , = 25 nm.The thin lines 2 and 4 and calculated with eqn (2) and ( 6 ) . The thick lines 1 and 3 are obtained for the limit yo = 00 with eqn (4). For the parameters chosen see text. The right-hand part (5) of the figure shows the reciprocal radius RD-' of stable mixed micelles. The divergence of R , coincides approximately with the limit of vanishing edge tension. The arrows mark the experiment with finite edge tension. Vanishing Edge Tension: Mixed Micelles The edge tension in a lipid suspension vanishes as the free concentration of an edge-active agent reaches the limit cw = c$ [eqn (3)]. Additional agent does not enhance the free concentration further, as this would lead to a negative edge tension according to eqn (2).The additional agent must create smaller and smaller membrane fragments with their edge occupied according to the Langmuir isotherm. The limit of vanishing edge tension is identical to the well known limit of a micellar phase.18y19 The intermicellar concentration is determined by the critical concentration c$. Experimental Dispersions of egg lecithin (Sigma, type HIE) and taurochenodesoxycholate (Sigma, TCDC) were studied by photon-correlation spectroscopy. Aliquots of 20 mmol dm-3 stock solutions in methanol were mixed. The solvent was evaporated with nitrogen. 2 cm3 of water with 150 mmol dmP3 NaCl and 10 mmol dmW3 tris at pH 8 were added. The samples were sonicated (Branson sonifer, level 3) at 0 "C eight times for 5 min with intervals of 5 min.The samples were filtered through a membrane filter (Nuclepore, 0.2 pm) into a cuvette. The measurements at 0 "C were started after an incubation time of 1 h. The concentration of lipid was checked by phosphate analysis.2o The beam of an argon-ion laser (Spectra Physics 165 with Etalon at 488 nm) was focussed into the dispersion. The scattering volume was imaged onto a photomultiplier (EM1 9863B/100). The photon counts (Malvern RF 313) were correlated (Malvern KP. Fromherz, C. Rocker and D. Ruppel 43 16 14 7 1 2 ' 10 2 2 $ 0 d 6 4 0 2 4 6 0 10 12 c,/mmol dmP3 Fig. 3. Reciprocal radius R,' of discoid mixed micelles made of egg lecithin and taurochenodesoxycholate (TCDC) as measured by photon correlation spectroscopy as a function of the total concentration C, of TCDC for six concentrations cJmmol dm-3 of lecithin as indicated.The set of straight lines is obtained by matching the three parameters c&=00.8mmol dmP3, KWM = 600 dm3 mol-' and ro = 3.7 nm of eqn (7) using a least-squares procedure. 7023). The intensity correlation was transformed into the field correlation21 and evalu- ated with the cumulant method22 up to the second cumulant in a microcomputer (Apple 11). From the mean diffusion coefficient, taken from the first cumulant, an effective hydrodynamic radius RH was obtained through the Stokes- Einstein relation. The radius of the membrane discs, RD, was obtained through the relation RH = aRD+&d with a thickness d = 5 nm.I9 Results and Discussion The reciprocal radius, RE', of the discs is shown in fig.3 for a series of concentrations, cr, of TCDC and six concentrations, cL, of lecithin. We obtain a functional relation R-'( c,, c,) by rearranging eqn ( 6 ) , setting cw = c& and substituting c& according to eqn (3). Eqn (7) comprises other more restricted relations used previously: I8*l9 r, = ?,a,[ 1 - exp (-yo/ CELT)]. We evaluate the data shown in fig. 3 in terms of eqn (7) using a bilinear least-squares procedure in the variables c,' and cT/cL. Three parameters are defined by this fit: the phase limit c&, the binding constant KWM and the characteristic radius r,. We obtain c& = 0.8 f 0.02 mmol dmP3, KWM = 600 f 15 dm3 mol-' and ro = 3.7 f 0.04 nm. The result supports the model of a micelle with cholate adsorbed to the edge" and solubilized in the bilayer." The parameter ro requires some attention.If TCDC adsorbs to the rim of both monolayers with the molecules closely stacked in an edge-on position, we estimate from44 Disc- Vesicle Transition CPK models a saturation of CE = 4 nm-'. With aM = 0.7 nm2 for lecithin23 we obtain the impossible relation ro> aM& However, we have to correct aM for the contribution of the solubilized TCDC, as the molar ratio TCDC/lecithin in the bulk membrane is ca. 0.5. From CPK models we estimate an effective value aM = 1 nm2. The ratio ro/aMFE = 0.925 now being close to 1 indicates, according to eqn (7), a large intrinsic edge tension with yo >> kTCE. We may estimate yo = 4.2 x J nm-'. From eqn (3) we obtain then KWE = 15 500 dm3 mol-'.The ratio of the binding constants, KWE/ KWM = 26, indicates that TCDC is indeed edge-active. We are now in the position to calculate the edge tension y( cT, cL) using the parameters just evaluated. Fig. 2 shows for cL = 2.5 mmol dmP3 the approximation of eqn (4), i.e. with yo=^, ( K W E = ~ , FE=3.7nm-') and the complete function of eqn (2) with yo = 4.2 x J nm-', both for dispersions of discs with RD = 25 nm and for vesicles without edge. Finite Edge Tension: The Disc-Vesicle Transition Any bilayer fragments existing in the regime of finite edge tension tend to be curved in order to lower the edge energy. Isotropic fluid discs of radius RD may be bent to spheroid shells of radius R up to spheres of radius RD/2 (fig. 1). We consider the energy profile E ( 0 ) along the relative curvature Q=RD/2R as a superposition of the edge energy and of the shell energy i t ~ e l f .~ " ~ We describe the changing packing of the shell as an elasticity of bending neglecting any shear stress in the plane of the rnemb~ane.'~.'~ Applying Hooke's law with the modulus k,, we obtain the parabolic term in eqn (8): E(a) = 8nke1a2+2vRDy( 1 -a2)'l2, a = RD/2R. (8) The changing edge energy is given by the second term in eqn (8), where the geometric relation between the periphery L and the radius R is used as L = 2?rRD[ 1 - ( RD/2R)2]1/2. The two factors 87~k,l and 2vRDy are the maximum energies of the shell and of the edge, respectively. In general this profile exhibits three local minima for the disc (a = 0) and the vesicle (a = k l ) as separated by activation barriers.' The transition probability from disc to vesicle as governed by a Brownian motion in the shape coordinate RD/2R drops exponentially with increasing height E g s of the barrier24 as given by where the 'vesiculation index' VF denotes the ratio of maximal energy of the edge and the shell.We can slow down the rate of the disc-vesicle transition by adjusting the value of VF, i.e. by adjusting the edge tension y for discs of radius RD, made of a material characterized by kel. The modulation of the edge tension by an edge-active agent provides a convenient tool to control the closure of discs. We obtained a relation between the rate of the disc-vesicle transition and the concentration C, of an edge-active solute by combining eqn (9) with eqn (2) and (6).9 Considering in this relation a particular rate, i.e.a particular Egs or VF, we obtain the concentrations cT required to close discs of a certain radius RD. Describing the edge tension as shown in fig. 2 and assuming kel = 6 x low2' J, fig. 4 shows the assignment of cT to the disc size RD for the maximum rate constant, i.e. for VF = 2: the smaller the discs, the larger the edge tension required to close the disc with maximum rate and the lower the concentration of the edge actant. Experimental Dispersions of egg lecithin and taurochenodesoxycholate were studied by electron microscopy.' *Plate 1. Electronmicrographs of sonicated dispersion of 2.5 mmol dm-3 egg lecithin with 0.8 mmol dm-3 taurochenodesoxycholate in 150 mmol dm-3 NaCl at 0 "C as stained by phos- photungstic acid 35 min (left) and 180 min (right) after the end of sonication.The pattern of parallel lines in the left-hand picture is assigned to finite discs of bilayer viewed in profile as stacked by the stain; the loops in the right-hand picture are assigned to unilamellar spherical vesicles as deformed by the stain. (To face p. 45)P. Fromherz, C. Rocker and D. Ruppel 45 0 1 2 c,/mmol dmp3 3 Fig. 4. Radius R , of bilayer discs closing to vesicles at a maximal rate ( 1 and 2) and of stable mixed micelles (3) as a function of the concentration C, of an edge-active amphiphile (taurochenodesoxycholate). The figure refers to a concentration of 2.5 mmol dm-3 egg lecithin. The micellar radius is obtained from eqn (9) with eqn (2) and ( 6 ) .The dashed line (2) refers to a state with all fragments in an open-disc state, the continuous line ( 1 ) refers to all fragments in the closed state. For the parameters chosen see text. The concentration of divergence of RD at c$ ( 1 + KwMcL) = 2 mmol dm-' is indicated. The arrows mark the conditions of the experiment. Aliquots of stock solutions of lecithin (2.6 mmol dm-3) and of TCDC (3.84 mmol dmP3) in methanol were mixed. The solvent was evaporated with nitrogen. 2 cm3 of water with 150 mmol dmP3 NaCl were added. The sample was sonicated at 0 "C eight times for 5 min with intervals of 5 min and centrifuged at 0 "C at 30 OOOg for 30 min. A drop was applied onto a carbon film (5-10 nm thick) on a copper grid (exposed to a glow discharge within 2 h before use) at 0 "C and sucked off after 30 s.The sample was stained applying a drop of phosphotungstic acid (Merck, 1% solution, pH 6.9) for 30 s. The grid was dried at 40 "C. The electron micrographs were taken using a Philips EM 301 instrument. The micrographs were projected onto a reversed plotter (Watanabe). Two types of objects could be distinguished: closed loops and straight lines, both isolated and in stacks. The number and size of these two classes of patterns were evaluated by a microcomputer (Apple 11). Objects which could not be assigned uniquely, as some multilamellar liposomes and blurred patterns, were not considered. Results and Discussion Two electronmicrographs of a dispersion of 2.5 mmol dmP3 lecithin and 0.8 mmol dmP3 TCDC are shown in plate 1 as obtained 35 and 180min after the last sonication.We assign the pattern of parallel lines to stacked discs seen in profile. The stacking is assigned to the staining process: On one hand the light scattering of the sample before fixation is typical of isolated particles, on the other hand stable mixed micelles are seen in a similar pattern using the same method of preparation." The pattern of loops is assigned to closed vesicles embedded in the stain. The ratio of the numbers of discs and of vesicles drops with a time constant of ca. 1.5 h. This estimate is based on the evaluation of 19 000 objects from four suspensions46 Disc- Vesicle Transition for ten time intervals within 3 h." The (number) average of the size of the discs just after sonication is ca. RD = 20 nm, the average size of the final vesicles corresponds to RD = 30 nm as evaluated from 180 objects for both classes.The polydispersity extends from ca. 30 to 50 nm." The difference in the average size cannot be considered to be significant at the present moment, as the evaluation of the micrographs is not unam- biguous owing to the unknown shape of discs and vesicles in the stain. Similar pictures are studied for 0 and 1.7 mmol dmP3 TCDC with 2.5 mmol dmP3 lecithin. Without cholate we observe only vesicles even immediately after sonication. With 1.7 mmol dm-3 TCDC even after 24 h only discs are visible, although we are far from the limit of stable micelles at 2.3 mmol dmP3 (fig. 2). The observations demonstrate the primary formation of discs by sonication and their spontaneous closure to vesicles.It is the rate of the disc-vesicle transition which is controlled by cholate. A moderate change in the concentration of cholate leads to a dramatic change in the rate in a narrow range of concentration, which is definitely lower than the phase limit of the micelles. These qualitative features are as would be expected from the concept of edge activity combined with the phenomenological stability theory.' Thus we make an attempt at a quantitative evaluation. In the first step we estimate the edge tension for c,=2.5 mmol dm-3, c,= C.8 mmol dm-3 and RD = 25 nm using the approximation with yo = a, KWE = 00 with all fragments being open at the beginning of the experiment. For the free concentration of TCDC we obtain from eqn (6) cw = 0.17 mmol dmP3 using CE = 3.7 nm-', aM = 1 nm2 and KWM = 600 dm3 mol-'.The edge tension is calculated from eqn (5) as y = 2.3 x lop2" J nm-'. We assign the maximum rate of closure with VF = 2 as an upper limit. We then obtain from eqn (8) a lower limit of the elastic modulus as k,, = 7.2 x lop2' J. In the second step we consider the finite intrinsic edge tension. We obtain a free concentration of cw = 0.2 mmol dm-3 according to eqn (6) with yo = 4.2 x J nm-', cE = 4 nm-', KWE = 15 500 dm3 mol-' and the other parameters as above. (At this con- centration the TCDC/lecithin ratio in the bulk bilayer is 0.12 and the degree of saturation of the edge is 0.75.) The edge tension is y = 2 x J nm-' according to eqn (2). For the elastic modulus we obtain in the limit VF = 2 from eqn (8) kel = 6 x J.J nm-' is the first experimental estimate for a bilayer. It is lower then the hydrophobic energy of the open edge, estimated to be yo = 7 x J nm-'.9 The difference indicates a negative contribu- tion originating in a repulsion of the headgroups of phosphatidylcholine in the bilayer as released in the edge. The packing of such a micellar edge made of lipid molecules may be rationalized easily by the block model." The elastic modulus estimated as kel = 6 x lop2' J is lower than that obtained from flickering of closed giant vesicles as k, = 23 x lop2' J.*' Those measurements refer to a closed bilayer with forbidden exchange of lipid between the two monolayers, whereas here we consider fragments with free exchange at their open edge.With the values of yo and kel as estimated we obtain the radius of fragments of pure lecithin, which may close with maximum rate at VF = 2 from eqn (8) as RD = 11.5 nm. This value corresponds nicely to the minimal diameter of 11 nm found for vesicles made from egg lecithin.26 The value of the intrinsic edge tension of lecithin yo = 4.2 x Summary We have described the mixed dispersion of a lipid and of a detergent as an inhomogeneous distribution of the detergent in preformed bilayers of the lipid. The inhomogeneity, the accumulation at the edge, has been connected with the energy of the edge by the application of the Gibbs isotherm. This approach has lead to a novel interpretation of stable micelles as stable 'two-dimensional emulsions' beyond the limit of vanishing edge tension.From an investigation of the system with vanishing edgeP. Fromherz, C. Rocker and D. Ruppel 47 tension we have obtained the parameters necessary to calculate the finite edge tension as a function of the concentration of the 'edge actant'. By a careful adjustment of the edge tension we have discovered metastable open discs after sonication. We have shown that these discs are transformed spontaneously into closed vesicles. Using a phenomeno- logical potential profile for this transformation we have obtained experimental estimates of the intrinsic edge tension and of the elastic modulus of a bilayer. The concepts of edge activity and of the disc-vesicle transformation have been shown to be reasonable for the material egg-lecithin-taurochenodesoxycholate and for the process of closure of discs, respectively.We believe that these concepts can be general- ized in two aspects as follows. (i) The process of a vesicle bursting is the reverse of the closure of a disc. The concentrations of an edge actant required for a certain rate of the two processes are separated by a hysteresis gap.' In the regime of closed vesicles the modulation of edge tension may induce the formation of fluctuating pores which lead to a lysis of vesicles long before any burst occurs.27 Note that the concentration ranges required to induce lysis, to close discs, to open vesicles and to form stable micelles have to be distinguished clearly from each other and from the critical micelle concentration of the detergent.(ii) Edge activity may be assigned not only to other cholates, but also to other detergents such as octylglycosid, cetyltrimethylammoniumbromide and triton X- 100, to other amphiphiles such as tetracain and chlorpromazin aqd even to small alcohols and ethers. On the other hand it may be possible to consider molecules as cholesterol as edge-avoiding, enhancing the edge tension of a bilayer. It remains to be seen how far various processes of vesicle formation such as dialy~is,~"~' dilution3 1-33 and chemical modification,34735 as well as such phenomena as haem~lysis'~ and membrane per- m e a t i ~ n , ~ ~ may be rationalized by the concepts proposed. We thank the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Indus- trie for their generous support.References 1 F. Krafft, Ber. Dtsch. Chem. Ges., 1896, 29, 1334. 2 E. Gorter and F. Grendel, J. Exp. Med., 1925, 41, 439. 3 C. Huang, Biochemistry, 1969, 8, 344. 4 F. Szoka and D. Papahadjopoulos, Annu. Rev. Biophys. Bioeng., 1980, 9, 467. 5 J. N. Israelachvili and D. J. Mitchell, Biochim. Biophys. Acta, 1975, 389, 13. 6 J. N. Israelachvili, D. J. Mitchell and B. W. Ninham, Biochim. Biophys. Acta, 1977, 470, 185. 7 D. A. Haydon and J. Taylor, J. 7'heor. Biol., 1963, 4, 281. 8 J. N. Israelachvili, D. J. Mitchell and B. W. Ninham, J. Chem. Soc., Faraday Trans. 2, 1976, 72, 1525. 9 P. Fromherz, Chem. Phys. Lett., 1983, 94, 259. 10 P. Fromherz, in Reverse Micelles, ed. P. L. Luisi and B. E. Straub (Plenum Press, New York, 1984), p. 55. 1 1 P. Fromherz and D. Ruppel, FEBS Lett., 1985, 179, 155. 12 E. G. Finer, A. G. Flook and H. Hauser, Biochim. Biophys. Acta, 1972, 260, 49. 13 F. C. Frank, Discuss. Faraday SOC., 1958, 25, 19. 14 W. Helfrich, 2. Naturforsch, Teil C, 1973, 28, 693. 15 J. L. Fergason and G. H. Brown, J. Am. Oil. Chem. Soc., 1968, 45, 120. 16 W. Helfrich, Phys. Lett., 1974, 50A, 115. 17 C. Wagner, Nachr. Akad. Wiss. Gottingen, 1973, 37. 18 D. M. Small, Gastroenterology, 1967, 52, 607. 19 N. A. Mazer, G. B. Benedek and M. C. Carey, Biochemistry, 1980, 19, 601. 20 P. S. Chen, T. Y. Toribara and H. Warner, Anal. Chem., 1956, 28, 1756. 21 Photon Correlation and Light Beating Spectroscopy, ed. H. Z . Cummins and E. R. Pike, (Plenum Press, New York, 1974). 22 D. E. Koppel, J. Chem. Phys., 1972, 57, 4814. 23 D. M. Small, J. Lipid. Res., 1967, 8, 551. 24 H. A. Kramers, Physica, 1940, 7, 284. 25 R. M. Servuss, W. Harbich and W. Helfrich, Biochim. Biophys. Acta, 1976, 436, 900.48 Disc- Vesicle Transition 26 B. A. Cornell, G. C. Fletcher, J. Middlehurst and F. Separavoc, Biochim. Biophys. Acta, 1982, 690, 15. 27 P. Fromherz, P. Forster and C. Rocker, in preparation. 28 Y. Kagawa and E. Racker, J. Bid. Chem., 1971, 246, 5477. 29 0. Zumbuhl and H. G. Weder, Biochim. Biophys. Acta, 1981,640, 252. 30 J. Brunner, J. Skrabal and H. Hauser, Biochim. Biophys. Acta, 1976, 455, 322. 31 S. Batzri and E. D. Korn, Biochim. Biophys. Acta, 1973, 248, 1015. 32 P. Schurtenberger, N. Mazer and W. Kanzig, J. Phys. Chem., 1985, 89, 1042. 33 L. Rydhag, P. Stenius and L. Odberg, J. Colloid Interface Sci., 1982, 89, 1042. 34 H. Hauser and N. Gais, Roc. Nut1 Acad. Sci. USA, 1982, 79, 1683. 35 R. Nayar and A. J. Schroit, Biochemistry, 1985, 24, 5967. 36 H. U. Weltzien, B. Arnold and R. Reuther, Biochim. Biophys. Acta, 1977, 466, 411. 37 P. Maher and S. J. Singer, Biochemistry, 1984, 23, 232. Received 19th December 1985

ISSN:0301-7249    

DOI:10.1039/DC9868100039

出版商:RSC    

年代:1986

数据来源: RSC

摘要:

Faraday Discuss. Chem. SOC., 1986, 81, 49-61 The Correlation between Molecular Orientational Order and Reorientational Dynamics of Probe Molecules in Lipid Multibilayers Gijs van Ginkel, Leo J. Korstanje, Herman van Langen and Yehudi K. Levine* Department of Molecular Biophysics, Physics Laboratory, University of Utrecht, Princetonplein 5, 3584 CC Utrecht, The Netherlands The behaviour of probe molecules in lipid systems can be characterized in terms of molecular order parameters and rates of reorientational motion. The correlation between these parameters has been investigated in oriented lipid multibilayer systems on changing the degree of unsaturation of the chains and the cholesterol content. Angle-resolved fluorescence depolarization experiments were carried out as a function of two angles: (1) the angle of incidence of the excitation beam and (2) the angle between the directions of incidence and observation.The e.s.r. spectra from nitroxide spin labels were simulated numerically with a general stochastic Liouville equation formalism. The probe molecules were assumed to undergo stochastic small-step reorientational diffusion subject to an orienting potential. Both techniques indicate that the introduction of unsaturation into the lipid chains lowers both the order parameters and the diffusion coefficients of the probe molecules. The incorporation of cholesterol reverses this trend. These findings are at odds with current ideas about the fluidity of membrane systems. Much of our current understanding of the dynamic structure of biological membranes has been derived from studies of model systems, particularly lipid bilayer systems.It is now generally accepted that the bilayer can be considered as an orientationally anisotropic fluid and that the degree of anisotropy varies across its thickness. The dynamic structure of the system is characterized in terms of order parameters and correlation times (or diffusion coefficients) for the various motions. However, the details of these properties revealed by 2H-n.m.r. seem to differ significantly from those obtained from intrusive probe techniques such as e.s.r. and fluorescence depolarization. As a result, the fidelity of the reporting of the probe molecules has been questioned in the last decade on the grounds that the molecules perturb the structure of their surroundings to such an extent that they do not monitor the intrinsic properties of the bilayer.Nevertheless, these probe techniques are particularly useful tools in membrane studies not only because of their sensitivity, but also because the specific labelling of complex molecules with 2H atoms is not a trivial task. In recent years there have been important new developments in the interpretation of e.s.r. and fluorescence depolarization experiments on liquid-crystalline materials. These treatments not only take explicitly into account the orientational and motional anisotropy, but are also valid in situations where the correlation times of the molecular motions are similar to the intrinsic timescale of the experiment, e.g. the fluorescence lifetime.One important conclusion is that the interpretation must be based on a model for reorientational motion and that the results obtained may only be valid within the context of that model. It is thus imperative to test the validity of the description used in the analysis. To this end one requires the use of macroscopically oriented membrane 4950 Order and Dynamics in Lipid Multibilayers systems which afford the determination of the anisotropic response of the reporter molecules. We shall present here a study of a number of oriented multibilayer systems differing in the degree of unsaturation of the hydrocarbon chains and cholesterol content using fluorescence depolarization and e.s.r. techniques. The results are analysed using the rotational diffusion model in which the probe molecules are assumed to undergo stochastic reorientational motions subject to an anisotropic orienting potential.This model is found to provide a satisfactory description of the behaviour of the probes in both the techniques used. The introduction of unsaturation into the lipid chains lowers both the order parameters and the rates of reorientational motion of the probe molecules. On the other hand, the incorporation of cholesterol into the bilayers has the opposite effect, enhancing the orientational order and the rates of motion. These findings are at odds with current ideas about membrane fluidity. Both techniques yield strikingly similar values for the order parameters and motional rates for probes anchored in the headgroup region of the bilayer.This indicates that the probes perturb the bilayer structure to a lesser extent than has been presumed up to now. Theory Orientation of Molecules in Membrane Systems We shall consider here the orientation behaviour of molecules relative to the bilayer normal, the local director, within the framework of the molecular field approximation. We shall further restrict the discussion to the case of a uniaxial bilayer system containing molecules with an effective cylindrically symmetric form. Thus the orientation of the molecules in the system is specified by the angle p between the molecular symmetry axes and the local director.' The orientational distribution of an average molecule in the system is then characterized by a probability distribution f( p).This distribution can be expressed2 as a series expansion of Legendre polynomials P,(cos p), each of which is weighted by an order parameter (PL) which is the ensemble average of the corresponding term This expansion defines the order parameters (P2) and (P4) as ( P 2 ) = i ( 3 cos2 p - 1) ( p4) = $(35 C O S ~ p - 30 C O S ~ p + 3). ( 2 4 ( 2 6 ) The orientational distribution function f ( P ) is fully characterized if all the order parameters (PL) are known. In practice, however, only (P2) and (P4) are accessible experimentally. For example 2H-n.m.r. and linear dichroism experiments yield ( P2) only, while e.s.r. and two-photon experiments such as Raman scattering and fluorescence depolarization yield both (P2) and (P4). The main difficulty now is obtaining an objective and realistic estimate of the form of f ( P ) from limited knowledge of its moments.This may be accomplished by an information-theoretic a p p r ~ a c h . ~ The essential point is that, given the order parameters ( P2) and ( P4), the most probable values of ( PL), L 3 6, are calculated under the assumption that the informational entropy off@) is a maximum. This corresponds to the construc- tion of the broadest possible distribution function consistent with the known values of (P2) and (P4). The resulting distribution function has the form f ( P ) = A exp [A2P*(cos P ) +A4P4(cos P)I (3)G. van Ginkel, L. J. Korstanje, El. van Langen and Y. K . Levine 51 where A is a normalization constant and h2 and h4 are determined from the known values of (P2) and (P4).If only the order parameter (P2) is known, the distribution function takes the form of eqn ( 3 ) but with h4 = 0. We note here that eqn (3) has the form of a Boltzmann distribution with an angle-dependent orienting potential. It can be seen from eqn (3) that if only ( P2) is known, the reconstructed distribution function either has a maximum at p = 0 and decreases monotonically to a minimum at /3 = 7r/2 or vice versa. Knowledge of (P4) is required for establishing the existence of a collective molecular tilt which is manifested by a maximum of the distribution function at an angle intermediate between 0 and ~ / 2 . On the other hand the observation of a minimum at such an angle may be taken as an indication for a superposition of two or more independent populations of molecules.The available information, however, is too limited to allow their resolution. Model for Reorientational Motion We shall assume that the reorientational motion of the molecule is a stochastic Markov process and neglect any inertial effect^.^ Consequently the Markov process need be described only in terms of the random angle p ( t ) , The master equation for the conditional probability P(Po/Pt) that the molecule has an orientation p relative to the director at time t, given that it had an orientation Po at t = O is (4) a - P(Po/Pt) = r,P(Po/Pt) at where rp is the stochastic operator describing the orientational motion. We shall here only consider the rotational diffusion model in which the average molecule is assumed to undergo small-step stochastic diffusion subject to an orienting potential U ( p ) . The Markov operator rP is now given by4 where D is the rotational diffusion tensor, diag (D,, D,, Dll), and M is the angular momentum ~ p e r a t o r , ~ both defined in the molecular frame.Eqn (4) is solved numeri- cally4 subject to the initial boundary condition P(pO/pO) = S(po-p). In view of the discussion above on the reconstruction of the equilibrium orientation distribution function, we shall choose U ( p ) to take the form U ( p ) = -~T[A,P~(COS p ) + A~P~(cos p ) ] . ( 6 ) We note that our choice of the orienting potential spans all the physically permissible pairs of (( P2), ( P4)) values. Angle-resolved Fluorescence Depolarization (AFD) Experiments The theory of the experiment has been discussed in detail e l ~ e w h e r e ~ - ~ and will only be summarized briefly. A macroscopically aligned bilayer system is subjected to con- tinuous illumination with light of defined wavelength and polarization direction.The geometrical arrangement is shown in fig. 1. The excitation light is incident at an angle 8 relative to the macroscopic director and is polarized in the zy plane. The fluorescence emission is observed at an angle 4 with its polarization either parallel (Ill) or perpen- dicular ( I , ) to the zy plane. In general the polarizations of the exciting light and the fluorescence emission will not be parallel in the zy plane. The polarization ratio Re = I,/ Ill is given by R , = [ I - R , + ( R ~ + R ~ ) sin2 ~ ] [ ~ + R , + ( R ~ - R ~ ) sin2 & + ( R ~ - R ~ ) sin2 e +R, sin2 8 sin2 & + R4 sin 28 sin 241-l.(7)52 Order and Dynamics in Lipid Multibilayers I X I I I I I I Fig. 1. Experimental AFD geometry for a multibilayer sample S lying in the xy plane. 6 and 4 are the angles in air between the macroscopic director z and, respectively, the direction of excitation and observation of the fluorescence. The states of polarization, either parallel or perpendicular to the zy plane, are determined by the polarizer P and the analyser A. The angles 8 and and the intensities Il and Ill in eqn (7) refer to quantities within the sample and not to those measured in the laboratory system. The angles in the sample and the laboratory can be related simply if the refractive index of the sample is known. However, the effects of the sample-air interface on the transmission of light intensity are more complex and depend furthermore on the direction of polarization.The various artefacts have been discussed in detail by Lax and Nelson.' It can be easily shown" that the effects of multiple reflections of the exciting and emitted light within the sample compensate the transmission losses of the fluorescent light at the interface. Furthermore, consideration of the +dependence of the intensities Il and 11, obtained with a normal incidence of the exciting light ( 8 = O O ) , indicate that only solid-angle expansion' distorts the experimental results. This effect cancels out on calculating the polarization ratio. Consequently no correction need be applied to the values of Re determined experimentally. The polarization ratio Re is measured for various combinations of 8 and 4 and affords the determination of five independent quantities Sp, S,, go, g , and g2 from steady-state experiments.Here Sp and S , are, respectively, the second-rank order parameters ( = ( P2)) for the absorption and emission transition moments. The quantities gk, k = 0, 1, 2 are defined by where DiI are Wigner rotation matrix elements" and .Iz, and fi, denote, respectively, the orientation of the absorption moment at time t = 0 and that of the emission moment at time t, relative to the director frame, fig. 1. F ( t ) denotes the normalized intrinsic fluorescence decay function. The time behaviour of the correlation function Gk(t) is obtained numerically from the solution of eqn (4)4,'2 and for many practical situations is found to be a mono- exponential decay.G. van Ginkel, L.J. Korstanje, H. van Langen and Y K. Levine 53 It is important to realize that the values of Gk(t) for t = 0 and t + 00 are model independent as a consequence of the assumption that the motion can be described as a stochastic process. These limits can be expressed solely in terms of the order parameters (P2) and (P4).l3,I4 The correlation functions G,( t ) and G2( t ) decay to zero at long time, whereas the function Go( t ) decays to a constant value given by ( P2)2. This behaviour is in marked contrast to that observed in isotropic systems. F ( t ) can be determined experimentally on observing the fluorescence emission in a direction normal to the sample surface ( 4 = 0') and under an angle of incidence in the sample 6 = sin-' ( l / f i ) .If the polarizer on the emission side is set with its optical axis at 45" to the vertical, then the combined signal Il + Ill is proportional to F ( t ) . Simulations of E.S.R. Spectra The analysis of e.s.r. spectra of nitroxide spin labels embedded in lipid bilayers in terms of the orientational distribution and the reorientational motions of the molecules has been reviewed in detail e1~ewhere.l~ Suffice it to say that the spin Hamiltonian describing the spectral line shapes contains contributions from the anisotropic Zeeman and hyperfine interactions and is given by where g and A are the Zeeman and hyperfine tensors respectively, S the electron spin and I the nitrogen spin operators.The interpretation of the e.s.r. spectra is not straightforward, but determined by the ratio of the time scale of the reorientational motions, expressed as a correlation time T, to the anisotropy of the hyperfine tensor A expressed in frequency units. Three regimes can be distinguished. (i) Fast-motion Regime ( lo-' < r / s < 2 x 1 0-9) In this regime the e.s.r. spectrum consists of three lines whose positions are determined by the time-averaged spin Hamilt~nian'~ and yield the order parameter ( P2). The spectral lineshapes containing the information about ( P4) and the reorientational motion can be calculated using Redfield's (ii) Slow-motion Regime (2 x In this regime the spin Hamiltonian must be considered as an explicit function of the orientation of the spin label with respect to the external static magnetic field B0.16717 The time evolution of the classical stochastic orientation is now coupled with the spin variables and the e.s.r.spectra can be simulated only on solving the stochastic Liouville equation (SLE) for the density matrix operator p ( a , t ) : l 6 > l 7 < r / s < where r p is defined by eqn (4), &(a, t ) is the Hamiltonian describing the interaction between the spins and the applied microwave field and p 0 ( a ) is the equilibrium density matrix operator. The SLE can only be solved numerically, and fast and reliable algorithms have been developed in recent years.17-19 The spectral lineshapes in this regime may become quite complex and often lose their triplet character.20y21 Spectra consisting of a superposition of five lines are fairly common.54 Order and Dynamics in Lipid Multibilayers Fig.2. Simulations of the spectrum of a CSL molecule embedded in a lipid bilayer with the applied magnetic field Bo perpendicular to the bilayer plane. The order parameter ( P2) = 0.45, while (P4) decreases from top to bottom ( a ) 0.20, (6) 0.13, ( c ) 0.10 and ( d ) 0.05; D, = 1.0 x lo7 s-' and Dll = 5D,. (iii) Powder [ Rigid-lattice Limit ( T / S > lop7 )] The reorientational motions are now too slow to affect the form of the e.s.r. spectra, which can thus be described as a superposition of lines arising from a static distribution of spin label molecules. It has not been widely appreciated that all three motional regimes can be treated by the SLE formalism.The drawbacks of this approach are its apparent mathematical complexity and the use of involved numerical algorithms for spectral simulations. Consequently it has been neglected in the past. The vast majority of people preferred the mathematically tractable Redfield theory for the analysis of e.s.r. experiments. This has often led to reports of motional rates quite clearly too slow to fall within the fast motion approximation. The question then arises as to whether the conclusions drawn in those studies are tenable. The adherence to the fast motional limit with its three line spectra, has led many workers to believe that any other spectral lineshape must be due to a superposition of signals from two or more populations of labels differing in their dynamic behaviour.However, as illustrated in fig. 2, a complex lineshape can also be obtained from a dynamically homogeneous population in the slow motion regime.20721G. van Ginkel, L. J. Korstanje, H. van Langen and Y K. Levine 55 Small changes in the order parameters (P4), keeping all other parameters constant, can be seen to induce large changes in the e.s.r. spectra. One is then forced to conclude that the interpretation of experimental e.s.r. spectra exhibiting five or more lines is ambiguous in the absence of other information about the system. The simulations of the experimental e.s.r. spectra presented below were carried out using the SLE formalism and the Lanczos a l g ~ r i t h m ’ ~ ” ~ for matrix diagonalization. In order to restrict the number of parameters, we have used the potential (6) with h4 = 0.The simulated spectra were convoluted with a Gaussian line in order to account for the broadening caused by unresolved electron-proton hyperfine interactions. Experimental Apparatus Angle-resolved fluorescence depolarization experiments were carried out with a home- built fluorimeter described Fluorescence decay measurements were carried out with a similar home-built set up, using an Edinburgh Instruments 399 flash-lamp operating with N2 gas at atmospheric pressure as a light source, and equipped with single-photon-counting detection, The fluorescence decay law as extracted from the measurements using POPOP as a reference compound following Zucker et a1.22 The intensity of the exciting light was kept as low as possible to avoid bleaching of the probe molecules and the intensity of fluorescence, measured at the same scattering geometry, remained constant during the experiments.Control experiments showed that the intrinsic fluorescence signals from the lipids and the coverslips amounted to <1% of the probe fluorescence intensity. E.s.r. measurements were carried out on a Varian E-9 X-band spectrometer equipped with a TM 110 cavity and a Varian V4540 variable-temperature accessory. Most of the spectra reported below were recorded with the applied static magnetic field normal to the plane of the sample. Materials Dimyristoylphosphatidylcholine (DMPC), palmitoyloleoylphosphatidylcholine (POPC), dioleoylphosphatidylcholine (DOPC), egg phosphatidylcholine (EPC) and cholesterol were obtained from Sigma and digalactosyldiglyceride (DGDG) was from Lipid Products.The lipids were used without further purification. The fluorescent probe 1,6-diphenyl- 1,3,5-hexatriene (DPH) was obtained from Fluka and its polar analogue trimethylamino-DPH (TMA-DPH) was purchased from Molecular Probes Inc. The probes were dissolved in absolute ethanol (AR, Baker) and stored in the dark at 4°C. The cholestane nitroxide spin label (CSL) was purchased from Syva. Doubly distilled water was used throughout. Sample Preparation Macroscopically aligned bilayer systems were prepared as described and the alignment monitored under a polarizing microscope equipped with a first-order red-plate. The sample contained ca. 25 ‘/o water unless otherwise stated. The molecular probe to lipid ratio was 1 : 250 for AFD experiments and 1 : 125 for e.s.r.experiments. Oxidation of the Lipids As the unsaturated DOPC and DGDG molecules are highly susceptible to oxidation, the sample preparation was carried out strictly under a nitrogen atmosphere. The lipids came into contact with air for a short time during the alignment procedure under the56 Order and Dynamics in Lipid Multibilayers polarizing microscope. The oxidation of the lipids was monitored after every preparative step by recording their absorption spectrum in the 200-300 nm region.z3 Samples showing traces of peroxide formation were discarded. Results and Discussion AFD Experiments The polarization ratios Re were measured for 56 combinations of 8 and $ for each sample and the parameters R,- R5 were determined from the angle-dependence following ref.(7) and (8). The quantities Sp, S,, go, g , and g , were obtained on taking appropriate linear combinations of R1-R5.6 The same values of Re within the experimental error of 2% were obtained on rotating the samples by an arbitrary angle about the normal to their surface. This indicates that the macroscopic distribution of probe molecules is uniaxially symmetric. The analysis shows that in all the systems studied S >S, in accordance with our previous and the recent finding of JohanssonZ57n other oriented lipid systems. Interestingly, we have found that the values of S, in multibilayers of egg PC increase on reducing the hydration of the bilayers. These observations indicate either that the absorption and emission moments of the DPH and TMA-DPH molecules are not mutually paralle17,R,24 or that the orientational distribution function of the molecules in the excited state differs significantly from that in the ground If the latter explanation is correct, then the analysis of the experiments becomes difficult.However, we have previously argued7 on the basis of the known photophysical properties of DPH 12,26727 that vibronic mixing of the close-lying ' B : and 'A: excited states leads to polarization borrowing effectsz8 and that consequently the two transition moments are not mutually parallel. We have therefore analysed the experimental data on assuming that the absorption transition moment, p, lies parallel to the molecular symmetry axis and that the emission moment, u, is tilted by an angle pV with respect to that axis.Furthermore we assume that all the molecules can be assigned to the same dynamic behaviour. With these assumptions we have It is important to realize that the ensemble averages implied in eqn ( 1 1 ) and (8) entail an average over the multibilayer stack and the derived order parameters and diffusion coefficients are not necessarily properties of a single bilayer. The experimental results were analysed' using the measured fluorescence decay behaviour given in table 1, with D,, h2, h4 and P2(c0s p,) as free model parameters. The derived values of the parameters for the different systems are summarized in table 1. In this connection it is important to note that the values shown change by <5% if the fluorescence decay function was taken to be monoexponential with a lifetime equal to (T): ( T ) = loa tF( t) dt.It is thus possible to analyse the experimental data without prior knowledge of the fluorescence decay and on taking F ( t ) to be a monoexponential function. In this case the model parameter D, is simply replaced by the product D,(T). It can be seen from table 1 that the angle P, for DPH varies with the composition of the system and can reach large values in unsaturated lipid bilayers. In marked contrast p, = 15 *I3" for TMA-DPH molecules embedded in all the lipid systems studied. A further marked difference between the two probes is that the orientation distribution for DPH is characterized by h2 == h4, while for TMA-DPH h2 > h4 in all the systemsC Q 3 Table 1.Model parameters and fluorescence decay obtained from AFD experiments Q 5. lipid + probe a1/a2 Tl/ns 7,/ns PZ A 2 h4 D,/ns-' f ( g / 2 > / f ( o ) T/"C "g DMPC + DPH - - 8.21 24 2.41 0.69 0.045 - 2 35 POPC + DPH 0.05 1.62 7.60 30 1.75 1.06 0.034 - 4 21 fr DOPC + DPH - - - 28 0.95 0.96 - 0.22 13 21 5 DGDG+ DPH - - - 34 1.55 0.75 - 0.04 6 21 2: EPc(24'/0 H20) + DPH 0.59 5.53 8.53 22 1.77 0.53 0.044 - 6 21 s EPC( 10% H20) + DPH - - 7.39 13 2.28 0.6 1 0.037 - 3 21 5. EPC(8.3Yo H20) + DPH 0.27 3.76 8.16 10 2.50 0.60 0.027 - 2 21 POPC/CHOL(20%) + DPH I - 8.0 14 2.93 1.22 0.042 - 0.5 21 3 DOPC/CHOL(20%) + DPH - - - 21 1.83 0.96 - 0.24 4 21 g DGDG/CHOL(20%) + DPH - - - 24 1.89 0.65 - 0.17 4 21 DMPC + TMA-DPH 0.07 0.54 5.8 12 3.24 0.06 0.035 - 0.5 35 s POPC + TMA-DPH 0.24 1 .oo 4.8 18 2.96 0.53 0.014 - 1 21 DOPC + TMA-DPH 1.33 2.45 5.42 19 2.41 0.18 0.022 - 2 21 3 DGDG+TMA-DPH 0.35 1.19 3.55 17 2.27 0.12 0.014 - POPC/CHOL(20%) +TMA-DPH - - 5.5 13 3.58 0.89 0.023 - 0.5 21 & 1 21 3 DPOC/CHOL(20%) +TMA-DPH - - 4.2 15 2.82 0.64 0.024 - 21 k 3 21 A DGDG/CHOL(20°/o + TMA-DPH - - 3.5 17 2.69 0.40 0.017 - 1.558 Order and Dynamics in L -80-60-40-20 0 20 40 '60 00 ipid Mult ibila y ers -80-60-40-20 0 20 40 60 80 P / " P / " 0.11 0 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 ) 0.0. 0. 0. 0 . -80-60-40-20 0 20 40 60 80 -80-60-40-20 0 20 40 60 80 P / " P l " Fig. 3. The orientational distribution functions of DPH ( a ) and (c) and TMA-DPH ( b ) and ( d ) molecules in multibilayers of POPC.(c) and ( d ) show a blown-up plot to emphasize the form of the distribution for angles near to * ~ / 2 . h @l IL v 4.0 ,&I 3.21 0.5 0 - 4 0 . 1 4.01 1 1 I I I 1 I I 1 I 1 1 I I I I 1 1 3.2 2.4 ;;;' 1.6 0.0 0 -80-6e40-20 0 20 40 60 80 Pl" P l " -80-60-40-20 0 20 40 60 80 P l " -80-60-40-20-0 20 40 60 80 P l " Fig. 4. The orientational distribution functions of DPH ( a ) and (c) and TMA-DPH ( b ) and ( d ) molecules in multibilayers of DOPC. (c) and ( d ) show a blown-up plot to emphasize the form of the distribution for angles near to * t / 2 .G. van Ginkel, L. J. Korstanje, H. van Langen and Y. K. Levine 59 studied. Fig. 3 and 4 show the distribution functionf(P) for the two probes in POPC and DOPC bilayers, respectively. The main difference can be seen to be the appearance of a distinct minimum at p - 60" for DPH molecules and their more pronounced tendency to lie with their axes parallel to the bilayer surface.This is also the case for the other lipid systems. The addition of cholesterol, however, suppresses this effect. Note (table 1) that in EPC bilayers, the population of DPH molecules lying parallel to the bilayer surface increases with increasing hydration. These results strongly suggest a heterogeneous distribution of DPH molecules in lipid bilayers, consistent with their lipophilic nature. On the other hand, TMA-DPH molecules, which are expected to be anchored at the headgroup region of the bilayer appear to form a homogeneous population in all the systems studied. We consider it therefore inappropriate to use DPH molecules as probes of membrane dynamic structure especially in unsaturated systems.The results for TMA-DPH molecules exhibit the known trends of lower order parameters in unsaturated systems than in saturated systems above their phase transition and the increase in molecular orientational order induced by cholester01.~~ However, the results show unambiguously that the diffusion coefficients D, are lower for the unsaturated systems at 21 "C, well above their phase transition, than for DMPC at 35 "C. Furthermore D, increases in the presence of cholesterol. This latter behaviour has been observed earlier using 2H-n.m.r. techniques." These findings run counter to established ideas about membrane fluidity29 and may simply indicate that the only role of unsatur- ation in lipid bilayer systems is the lowering of the phase transition temperature of the chains, E.S.R.Experiments E.s.r. experiments complement AFD measurements on DPH and TMA-DPH in that the spectra of CSL molecules embedded in lipid bilayers reflect not only the orientational ordering and D,, but also Dll characterizing the reorientational motion of the molecules about their long axes.16-19 As CSL molecules are known to be anchored in the head-group region of the bilayers, their behaviour should be analogous to that of TMA-DPH molecules. The three-line e.s.r. spectra of CSL in bilayers of DMPC and POPC above their phase transition were simulated numerically with the general SLE formalism employing the simple potential U ( P ) = -~TA~P*(cos p). This approximation provides an adequate description of the AFD results with TMA-DPH as probe.The temperature dependences of the order parameter (P2) and the diffusion coefficients D, and Dll are shown in fig. 5 and 6, respectively. The values of the diffusion coefficients found fall quite clearly within the slow-motion regime for both systems. This conclusion is supported by the observation of a significant asymmetry in the positions of the low- and high-field lines relative to the centre of the spectrum. We have previously shown that still lower values of the diffusion coefficients are derived from simulations of the spectra of CSL in DGDG bilayer systems.2' The results show that the ordering of CSL molecules in DMPC bilayers is higher than in POPC bilayers, yet in the 30-50 "C temperature range similar values for D, are found.Nevertheless, the CSL molecules appear to rotate faster about their long axes in the DMPC bilayer over the same temperature range. Table 2 shows that the results obtained for the same lipid system with CSL and TMA-DPH probe molecules are strikingly similar. This agreement supports our approach to the interpretation of the experimental results and the conclusions reached.60 1.0 0.9 0 . 8 0 . 7 0 . 6 0 . 5 0.4; Order and Dynamics in Lipid Multibilayers I I 1 I I I - - - 8 - 0 D - 0 0 8 - ,,no c1 - 8 . - 8 " - - I I I I I I Fig. 5. The temperature dependences of (P2) obtained from numerical simulations of the spectra from CSL molecules embedded in multibilayers of DMPC (0) and POPC (W) above their respective phase transitions.1 o9 1 o8 1 o7 1 o6 1 o5 3.0 3.1 3.2 3 . 3 3 . 4 3 . 5 3.6 3.7 lo3 KIT Fig. 6. The temperature dependences of the diffusion coefficients obtained from numerical simula- tions of the spectra from CSL molecules embedded in multibilayers of DMPC (0, Dil; 0, Dl) and POPC (0, Dll; ., Dl) above their respective phase transitions.G. van Ginkel, L. J, Korstanje, H. van Langen and Y. K . Levine Table 2. A comparison of the parameters obtained with TMA-DPH and CSL molecules embedded in different multibilayer systems 61 label POPC, 22 "C CSL 0.60 0.24 1.7 x 10' TMA 0.66 0.35 1.4 x lo7 CSL 0.68 0.26 3.5 x lo7 TMA 0.64 0.29 3.5 x 10' TMA 0.50 0.18 1 . 4 ~ lo7 DMPC, 35 "C DGDG, 22°C CSL" 0.40 0.10 8 x lo6 " From ref. (21).We gratefully acknowledge the contributions made by G. Deinum, D. Engelen and F. Mulders to the experimental work and thank Dr A. J. Dammers for developing the e.s.r. simulation programs. References 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 I C. Zannoni, in The Molecular Physics of Liquid Crystals ed. G. R. Luckhurst and G. W. Gray (Academic Press, London, 1979), chap. 3, pp. 51-83. L. B-A. Johansson and G. Lindblom, Q. Rev. Biophys., 1980, 13, 63. The Maximum Entropy Formalism, ed. R. D. Levine and N. Tribus (M.I.T. Press, Boston, 1979). P. L. Nordio and U. Segre, in The Molecular Physics of Liquid Crystals, ed. G. R. Luckhurst and G. W. Gray (Academic Press, London, 1979), chap. 18, pp. 411-426. E. Merzbacher, Quantum Mechanics (Wiley, New York, 2nd edn, 1970), chap. 9, pp.172-190. B. W. van der Meer, R. P. H. Kooyman and Y. K. Levine, Chem. Phys., 1982, 66, 39. R. P. H. Kooyman, M. Vos and Y. K. Levine, Chem. Phys., 1983,81, 461. M. J. M. van de Ven and Y. K. Levine, Biochim. Biophys. Acta, 1984,777, 283. M. Lax and D. F. Nelson, in Coherence and Quantum Optics, ed. L. Mandel and E. Wolf (Plenum, New York, 1973), pp. 415-445. J. NauB, AUPO, Fac. r. Nar. vol. 76, Physica XXII, 1983, pp. 9-18. M. E. Rose, Elementary Theory of Angular Momentum (Wiley, New York, 1957). C. Zannoni, A. Arcioni and P. Cavatorta, Chem. Phys. Lipids, 1983, 32, 179. P. L. Nordio, in Spin Labelling, Theory and Applications, ed. L. J. Berliner (Academic Press, New York, 1976), chap. 2, pp. 5-52. J. Seelig, in Spin Labelling, Theory and Applications, ed. L. J. Berliner (Academic Press, New York, 1976), chap. 10, pp. 373-410. Spin Labelling, Theory and Applications, ed. L. J. Berliner (Academic Press, New York, 1976). J. H. Freed, in Spin Labelling, Theory and Applications, ed. L. J. Berliner (Academic Press, New York, 1976), chap. 3, pp. 53-132. A. J. Dammers, Ph.D. Thesis (University of Utrecht, 1985). G. Moro and J. H. Freed, J. Chem. Phys., 1981, 74, 3757. A. J. Dammers, Y. K. Levine and J. A. Tjon, Chem. Phys. Lett., 1982, 88, 198. E. Meirovitch, A. Nayeem and J. H. Freed, J. Phys. Chem., 1984, 88, 3454. P. Koole, A. J. Dammers, G. van Ginkel and Y. K. Levine, Biochim. Biophys. Acta, 1984,777, 297. M. Zuker, A. G. Szabo, L. Bramall, D. T. Krajcarski and B. Selinger, Rev. Sci. Instrum., 1985, 56, 14. R. A. Klein, Biochim. Biophys. Acta, 1970, 210, 486. M. Vos, R. P. H. Kooyman and Y. K. Levine, Biochem. Biophys. Res. Commun., 1983, 116, 462. L. B-A. Johansson, Chem. Phys. Lett., 1985, 118, 516. P. C. Alford and T. F. Palmer, Chem. Phys. Lett., 1982, 86, 248. B. E. Kohler and T. A. Spiglamin, J. Chem. Phys., 1984, 80, 5465. A. C. Albrecht, J. Chem. Phys., 1960, 33, 156. 29. D. Chapman and G. Benga, Biological Membranes 5 (Academic Press, London, 1984), chap. 1, pp. 1-56. 30 R. Ghosh and J. Seelig, Biochim. Biophys. Acta, 1982, 691, 151. Received 5 th December, 1985

ISSN:0301-7249    

DOI:10.1039/DC9868100049

出版商:RSC    

年代:1986

数据来源: RSC

摘要:

GENERAL DISCUSSION Prof. J. F. Nagle (Carnegie-Mellon University, Pittsburgh, PA) addressed Dr Gersh- feld: These are interesting experimental results. Your characterization of T" as being a critical temperature seems a most plausible suggestion, but one that I find difficult to reconcile with your characterization of the state at T" as being a surface' bilayer. As you have anticipated in your publications, a surface bilayer seems so counterintuitive that perhaps other interpretations should be entertained. I would like tentatively to suggest a different model. In this model for temperatures slightly lower than T" there is a monolayer with occasional fluctuating infolding to form bilayers underneath part of the monolayer. As T" is approached the infolding becomes critically unstable, so that there is one (or more) bilayer beneath all the monolayer. Above T* the accumulated subsurface bilayers peel of€ the monolayer owing to their own weight.This infolding model predicts a surface coverage at T* greater than the equivalent of three monolayers ( i e . one monolayer and one bilayer) over a very narrow temperature range; owing to the difficulty of precise temperature control this higher surface coverage could easily have been missed in your surface coverage measurements that reported a maximum surface coverage near two. Surface potential measurements might be a better way to distinguish between the two interpretations. A single symmetric bilayer would have no net potential across it, whereas a monolayer, or a monolayer with one or more bilayers beneath it, would have typical monolayer potentials of ca.500mV. Therefore, as a function of temperature the surface potential would change by ca. 500 mV according to your surface bilayer interpretation. In contrast, the surface potential would change much less according to the infolding monolayer model sketched above. Dr N. L. Gershfeld (National Institutes of Health, Bethesda, MD) replied: The model you propose has some appealing features because it may provide a reasonable model for films with densities between one and two monolayers at temperatures above and below T". However, at T* the model is inconsistent with a number of our observations. ( a ) Within the experimental error of the radioisotope measurement (*2%) the film density does not fluctuate after equilibrium has been attained, but remains at the bilayer density.Even after stirring the dispersion vigorously, the bilayer density is recovered after equilibrium is re-established.' ( b ) Resistance to the evaporation of water occurs only when the dispersion is at T" (k0.1 K); the monolayer gives little or no resistance to evaporation.2 In your model the properties of the monolayer would dominate over the entire temperature range, and we would not have observed any evaporation resistance at T*. We view the surface bilayer to be a continuous bilayer structure with water on both surfaces. Since the film density falls fairly steeply to values between one and two monolayers at temperatures above and below T", the surface potential may very well approach the monolayer value, particularly if the temperature is not carefully controlled at T".We have estimated that the temperature should be controlled to at least kO.001 K to observe pure bilayer properties at T".2 1 K. Tajima and N. L. Gershfeld, Biophys. J., 1985, 47, 203. 2 L. Ginsberg and N. L. Gershfeld, Biophys. J., 1985, 47, 211. Dr D. Marsh (Max-Planck-Institut Gottingen, West Germany) turned to Dr Gersh- feld. I would like to ask you to comment on the extremely high aqueous solubilities which you observe for DMPG. At the approximate position of the Krafft point in fig. 3 of your paper, the critical micelle concentration (c.m.c.) is ca. mol dme3. Interpola- tion of the logarithmic chain-length dependence of the c.m.c.s measured for diacyl 6364 0 General Discussion I I I I , i+-J m phosphatidylcholines' suggests that the c.m.c.for dimyristoyl phosphatidylcholine is ca. mol dm-'. Dr King and I have measured the c.m.c.s of a short-chain, spin-labelled phosphatidylcholine and phosphatidylglycerol using electron spin resonance spectros- copy. The salt dependence of the c.m.c.s at pH 7 is given in fig. 1. In the absence of added salt, (at a counter ion concentration of 10-4moldm-3) the c.m.c. of phos- phatidylglycerol is ca. 10 times greater than that of phosphatidylcholine. Hence I would expect the c.m.c. of DMPG to be of the order of ca. lo-* mol dm-3. 1. R. J. M. Tausk, J. Karmiggelt, C. Oudshoorn and J. Th. G. Overbeek Biophys. Chem., 1974,1, 175; 184; R. Smith and C. Tanford, J. Mol. Biol., 1972, 67, 75. Dr N .L. Gershfeld (National Institutes of Health, Bethesda, MD) said: Dr Marsh's estimate of the c.m.c. for DMPG is based on the assumption that the value of 10 for the ratio of c.m.c.s that he measured for the spin-labelled compounds was applicable to the dimyristoyl analogues. Solubility vs. temperature data for DMPC suggest that it is not (unpublished results). The solubility data indicate that the Krafft temperatures, where the solubility of the lipid increases markedly with temperature, are approximately the same for DMPG and DMPC; however, the solubility, and the c.m.c., for DMPC at this temperature is ca. 10-7moldm-3. Thus, a factor of ca. 1000 rather than 10 might be anticipated for the c.m.c. ratios of the dimyristoyl analogues. Moreover, the extrapolation used to obtain the c.m.c.of DMPC may not be valid. Indeed, the extrapolation underestimates the c.m.c. of DPPC obtained by Smith and Tanford by at least an order of magnitude. Given these two corrections to Dr Marsh's estimate of the c.m.c. of DMPG, our value for the solubility of DMPG appears to be reasonable.General Discussior 65 Dr D. P. Siege1 (Proctor and Gamble, Cincinnatti, OH) posed two questions: ( a ) I wonder if the existence of the ‘jelly’ phase may be due in part to the strong electrostatic repulsions between bilayers in these nearly salt-free systems. Such interac- tions could make the dispersion quite viscous because either the Debye length may be significant compared to the average inter-bilayer separation in the dispersion, or because the electrostatic repulsion is so strong (i.e.the surface electrostatic potential in the system is so large) that the repulsive forces between bilayer aggregates is significant even when they are many Debye lengths apart. If this were true, the effective viscosity of the system would increase with increasing lipid concentration, as you observed, because of the corresponding decrease in average inter-bilayer separation in the disper- sion. Have you observed this viscous phase in the presence of added electrolytes? ( b ) One of the most interesting aspects of your work is the implication that the spontaneous production of unilamellar vesicles at temperatures around T* is a higher- order phase transition in the La phase. Higher-order phase transitions are accompanied by discontinuities in the corresponding derivatives of the free energy with respect to temperature and pressure ( e.g.discontinuities in the heat capacity and isothermal compressibility accompany second-order phase transitions). Is there any experimental evidence for discontinuities in these derivatives in the L, phase at T*? Dr N. L. Gershfeld (National Institutes of Health, Bethesda, MD) replied: ( a ) Preliminary studies indicate that the ‘jelly’ phase persists in mol dm-3 NaCl solutions, although the viscosity of this phase appears to decrease at this ionic strength. At still higher NaCl concentrations, e.g. 0.1 mol d ~ l l - ~ , the ‘jelly’ phase does not seem to form. ( b ) Heat-capacity measurements in the region of T* have not been reported, but we have begun to examine this temperature interval very carefully with a high-precision adiabatic calorimeter.However, we anticipate that the effects are likely to be very small because the transformation is between two liquid-crystalline bilayer states, the jelly and the unilamellar vesicle. Dr M. N. Jones (University of Manchester) remarked: We have found that sonicated vesicles of DPPC or DPPC+PI are disrupted by glass surfaces in the form of either solid or porous glass beads. This disruption catalysed by a glass surface results in the formation of a more turbid lipid suspension, the precise structure of which has yet to be established. I note that you suggest that glass beads facilitate the formation of vesicles from crystals of DMPG. Would you care to suggest a mechanism for the process and comment on the size of the vesicles formed in the presence of the glass beads? Dr N.L. Gershfeld (National Institutes of Health, Bethesda, MD) replied: In the absence of glass beads the diameter of the vesicles formed at T* is of the order of 0.5 pm, while in the presence of beads the vesicles are ca. 5-50 pm. The former estimate is based on preliminary light-scattering measurements. Given the paucity of data regarding this effect I hesitate to suggest any mechanisms for the process, except to note that the glass beads appear to increase the size of the vesicles in both of our systems. Prof. D. G. Hall (Unilever Research Port Sunlight Laboratory) said: My comments are concerned with fig. 1 of the paper: me vs T. (1) I would expect a change in slope at T,, but there are too few data points for this to be apparent, do you agree? (2) The break at T* appears to be quite sharp.A break of this kind suggests the occurrence of a first-order phase transition. Indeed it is hard to see how a higher-order transition can be responsible. Since there is no evidence of a first-order transition of T* in the bulk phases could the effect be due to a first-order transition in the interface? If so should there not be a sharp change in the surface excess r? Has such a change been observed? If the transition is blurred the methods developed in ref. ( 1 ) may be useful. 1. D. G. Hall, J. Chem. SOC., Furuduy Trans. 2, 1972, 68, 668.66 General Discussion Dr N. L. Gershfeld (National Institutes of Health, Bethesda, MD) replied: The major focus of this paper was to establish that at T", where the surface pressure is a maximum, the phospholipid dispersion forms a suspension of unilamellar vesicles.We were primarily concerned with identifying T* for DMPG, and therefore obtained only the minimum number of data sufficient to establish this temperature. The points you have raised about the relationship between the (rIe, T ) data of fig. 1 and lipid transitions have been addressed in detail with other phospholipids. Thus, for dispersions of dimyristoylphosphatidylcholine (DMPC), whose values of T", T, and AH, are similar to those for DMPG, we have observed a change in slope at T,, and were able to evaluate the latent heat of the transition from the slopes at this temperature [ref.(7) of the paper]. The Il, vs. T curve for DMPC shows a sharp break at T* similar to the one exhibited in our fig. 1. However, independent measurements of the surface concentration using radiotracers shows that the film density is a continuous function of the temperature with a maximum at T* [ref. (7) and (8) of the paper]. Since a first-order transition requires that the surface concentration be discontinuous at the transition temperature, the transition at T* is assumed to be of higher order. For DMPG, T* coincides with the temperature where the 'jelly' is transformed to the unilamellar vesicles (fig. 4 of the paper). This bulk phase transformation occurs over several degrees, and we have therefore concluded that it is a higher-order transition. We assume that an equivalent phenomenon occurs in the surface film at T*.Mr F. A. M. Leermakers, Dr J. M. H. M. Scheutjens and Prof. J. Lyklema (Agricultural University of Wageningen, The Netherlands) said: Data on the free lipid concentrations in equilibrium with a membrane are very useful for thermodynamical analyses. Our statistical-thermodynamical theory predicts the equilibrium volume fraction of the lipids for given head-solvent, solvent-tail and tail- head interaction parameters.' It appears that these concentrations are very close to the critical micelle concentration (c.m.c.). The interaction parameters used are enthalpies per kT, Le. inversely proportional to the absolute temperature. Hence, from our theory the temperature dependence of the critical volume fraction can also be obtained. Fig.2 shows the result for a series of non-ionic lecithin-like molecules. The curves look very similar to the experimental data given in fig. 3 of the paper by Gershfeld et al. which are replotted in our fig. 2. Only at high temperatures do significant deviations between theory and experiment occur. As sug- gested in the paper, these deviations might be due to the formation of micelles which can not be separated by centrifugation. Fig. 2 also shows that the lines for different tail lengths are parallel with respect to each other, both above and below T,. The vertical distance between the curves is ca. 1.2, i.e. 0.3 per added tail segment, which correlates well with known c.m.c. data for small ionic lipids.* Furthermore, the molecular-weight dependence of the discontinuities agrees with experimental T, data.Does Dr Gershfeld have experimental indications that for different tail lengths the lines are indeed parallel to each other and shifted as predicted above? 1 F. A. M. Leermakers, J. M. H. M. Scheutjens and J. Lyklema, Biophys. Chem., 1983, 18, 353. 2 Solution Behavior ofSurfactants, ed. K. L. Mittal (Plenum Press, New York, 1982), vol. 1 and 2. Dr N. L. Gershfeld (National Institutes of Health, Bethesda, MD) replied: The only T us. solubility data presently completed are for DMPG. However, the slope of the solubility curve depends on the latent heat of the gel-liquid-crystal transition. Since this heat varies with chain length, we would not expect the solubility vs. temperature curves for the homologues of DMPG to be parallel. Dr G.Cevc (Essen University, West Germany) said: Dr Parsegian told us that the addition of tetradecane to phosphatidylethanolamine in the inverted hexagonal phaseGeneral Discussion 67 10 30 50 T / "C Fig. 2. Dependence of the volume fraction 4* in equilibrium with a membrane on temperature. The theoretical curves apply to membranes composed of nonionic lecithin-like molecules with variable tail length x. The experimental curve is replotted from fig. 3 of the paper by Gershfeld et al. The interaction parameters at 300 K are: xAB = 1.2, xAS = 1.6, xBS = 0 for tail-head, tail- solvent and head-solvent, respectively. The trans-gauche energy is (275/300)kT. does not change the radius of the water cylinders appreciably, whereas partitioning of this alkane into the phosphatidylcholine hydrocarbon matrix increases the swelling of the latter lipid in such phase substantially.However, the samples of phosphatidy- lethanolamine that he was comparing contained 5 YO and those of phosphatidylcholine 20% of the alkane. Would the conclusion be different if the tetradecane concentration was in both cases the same? Secondly I would like to ask where Dr Parsegian envisages that the tetradecane molecules sit in the lipid hydrocarbon region? Does he think that different localization of the alkane molecules in the lipid hydrocarbon cores might provide a partial explanation for the large differences in the behaviour of phosphatidylethanolamine and phos- phatidylcholine upon the tetradecane-induced perturbation? Dr V.A. Parsegian (National Institutes of Health, Bethesda, MD) replied: (1) We have in fact looked over a range of alkane concentrations. The two concentrations used in the text were above saturating values in their respective systems. It may be that extra alkane is simply not incorporated into a regular lattice but sits aside in a separate pool invisible to low-angle X-ray diffraction. We expect further therefore no difference in our conclusions if both preparations be at 20% tetradecane. (2) We suggest favoured accumulation of deuterated alkane in the directions of d,,, . Alkanes can go elsewhere, but not to the same concentration. Certainly there can be differences in alkane distribu- tion in PC and PE systems, since each of these species is expected to have different spontaneous curvatures.Prof. D. A. Haydon (Cambridge University) said: I note that n-tetradecane was used to test for the existence of stress in the alkyl chain region of the H phase. In lipid68 General Discussion bilayers the penetration of n-alkanes into the hydrocarbon chain region declines as the size of the n-alkane increases’ and n-tetradecane lies in the chain-length range in which this effect is becoming important. Thus n-tetradecane is not the best choice of hydrocar- bon. Have the authors investigated the effects of shorter n-alkanes-on the H phase? 1 R Fettiplace, D. M. Andrews and D. A. Haydon, J. Membr. Biof., 1971, 5, 277. Dr V. A. Parsegian (National Institutes of Health, Bethesda, MD) answered Prof. Haydon as follows: This is a good question! Ideally one would like to use an alkane small enough to penetrate all hydrocarbon regions, but small substances are too volatile to be practical.Dodecane acts much as tetradecane on the phase structure, but we have used no alkanes smaller than dodecane. It might help to remember that we are looking at alkanes in HII lattices not in bilayers. Difficulties of tetradecane entry need not be the same here. The neutron- diffraction observation that alkane is indeed entering the lattice together with the null result of penetrant alkane on the osmotic stress as a function of lattice spacing should suffice to support our present conclusion. Having said this, it would not surprise us if slightly different behaviour were indeed seen with different medium-length alkanes ( n = 10-16, say).For very small alkanes we expect the entropy of distribution to be relatively important; for longer-chain alkanes packing constraints on their incorporation should be more important. Dr D. Marsh ( Max-Planck-Institut Gottingen, West Germany) addressed Dr Parsegian. First I have a comment regarding acyl chain stress in the HII phase. We have measured the dimensional changes at the L,-HII phase transition for two saturated phosphatidylethanolamines of differing chain-length.’ A geometrical calculation of the water dependence of the hexagonal chain dimension, d,,, , for diarachinoyl phosphatidylethanolamine, based on these dimensional measurements, is given in fig. 3. At the water content corresponding to the limiting hydration of the HII phase at a temperature just above the L,-HII transition, the value of d,,, is equal to the lipid thickness in the fluid lamellar phase immediately below the transition.This also holds approximately true for the hexagonal phase transition in didodecyl phosphatidy- lethanolamine. Thus the L,-HII transition takes place without any increase in hydrocar- bon chain extension. This result seems to be in full agreement with your experiments on the effects of added alkane and may constitute a guiding principle for hydrocarbon chain packing in the HI, phase. My question relates to your interpretation of the relative hydration properties of PE and PC at very low water activity. From your fig. 3 (and cf. also the accompanying fig. 3) it is clear that the simple ‘water cylinder’ structure of the HI, phase cannot hold at these very low water contents. What is known about the detailed molecular structure of the HII phase under these conditions, and is it possible that the osmotic stress properties are dominated by the structural rearrangement which must take place in the water/ headgroup/chains at these low water compositions, and not by the headgroup hydration properties per se ? 1 J.M. Seddon, G. Cevc, R. D. Kaye and D. Marsh, Biochemistry, 1984, 23, 2634. Dr V. A. Parsegian (National Institutes of Health, Bethesda, MD) replied: Yes, a water cylinder picture is certainly too idealized at low water contents. That is why we referred to the polar aqueous region there as a ‘mash’. Our suspicion is that the hydration in this regime depends only weakly on the constraint that the polar groups are attached to acyl chains.We hope to test this suspicion by measuring the hydration of polar group preparations. We will then see whether there are large PE/ PC differences.General Discussion - 0 69 I 1 1 1 I I lipids per perimeter water content, (1 - c) Fig. 3. Variation of the maximum length of a lipid molecule, d,,,, as a function of fractional water content, (1 - c) w/w in the HI* pbase, calculated using purely geometrical considerations and the values of area per molecule = 49 A2 and partial specific volume 6, = 1.043 cm3 g-' measured for diarachinoyl phosphatidylethanolamine in the HI, phase at maximum hydration.' The horizon- tal dashed line corresponds to the lipid thickness in the fluid lamellar La phase immediately below the L,-HII transition and the full line to that in the crystalline lamellar L, phase.' Dr J.M. Seddon (University of Southampton) asked Dr. Parsegian. (1) What values of area per molecule (at the water/lipid interface) did you measure in the HII phase, and are these values lower than those measured in La? (2) Do you consider it likely that a tighter headgroup packing in the HII phase of PE would permit more extensive and/or stronger hydrogen bonding between the headgroups than in the L, phase? (3) Could this mean that the apparent similarity between the pressure curves for DOPE- DOPC-tetradecane and DOPE at low hydration is actually spurious, in the former case the principal contribution to the work coming from the dehydration of the relatively polar headgroups, whereas in the latter case the work arising partly from dehydrction, and partly from a breaking of hydrogen bonds upon the structural rearrangement of the PE headgroups that must occur below some minimum radius of the water cylinders? Dr V.A. Parsegian (National Institutes of Health, Bethesda, MD) replied to p r Seddon's three questionsoas follows: ( 1 ) The HII areas are as folloys: DOPE, 53.5 A3 (full hydration) to 23.9 A2; DOPES;;S% tetradecane, 55.3 to 24.3 A2; DOPE: DOPC 3 : 1 + 20% tetradecane, 71.3 to 30.9 A2. Only the last can be easily compar$d with La, where for DOPE: DOPC 3: 1 (no tetradecane) areas ran from 66 to 61.4 A. The HII values are lower than for L, only after some dehydration. (2) Smaller areas should certainly allow tighter head-group packing.The continuous change in area will make it impossible to maintain a particular head-group packing over much of a hydration70 General Discussion range. Specific arrangements of hydrogen bonding are beyond us now. (3) Perhaps the similarity is spurious, but we think not. A similar convergence is seen with egg PC and egg PE lamellae at high osmotic stress [ref. (32) of our paper]. Dr G. J. T. Tiddy (Unilever Research Port Sunlight Laboratory) asked Dr Parsegian. How critical is the position of the lipidlwater boundary in the calculation of ro and r? What is the change in K , if the headgroups are included in the aqueous region? Dr V. A. Parsegian (National Institutes of Health, Bethesda, MD) replied: Not To see this, recall that we originally got K , from critical; there is an increase in K , to bring it even closer to the planar bilayer values.-_- rl ro If we use another measure of radius p = r(1 + q) and use the same definition P m P1) lim - PI+PO 1 1 -_- P1 Po where and we have --- rl ro 1.e. the estimate of K , increases by a factor (1 + q)3. molecule, we obtain (1 + q ) from Now if we add a volume v, per polar group to the volume of water v, per phospholipid 2, + v , v p 2 P--- - v r 2 - (1 + T I 2 VW and a correction factor to K , , [ 1 + ( u p / v,,,)]~’~. From our measured density of DOPE polar groups we have 215.2 0, = x 0.66 = 236 A3. 0.602 x loz4 From the data for DOPE HI* phase at zero stress v, = 600 A3, to give a maximal correction of 1.64 or 64%. So 4.6kT becomes 7.6kT, in better accord with the (7-17)kT expected from planar membranes [ref.(26) of our paper]. Prof. B. de Kruijff (Utrecht, The Netherlands) said: I wish to make one comment and pose one question to Dr Parsegian.General Discussion 71 (1) First, I would like to comment that I, with you, argue that it is essential to obtain detailed insight into the phase behaviour of lipids under equilibrium conditions. However, since biological membranes are not at equilibrium, insight into (intermediate) non-equilibrium lipid structures in model membranes is likely to provide a greater understanding of the possible biological significance of lipid polymorphism. (2) Concerning your studies on the effect of alkanes on phase structure of hydrated PE and PC, I would like to ask whether you have any experimental data on the localisation of these molecules in either the bilayer or the hexagonal Hrr phase.This in view of a possible discrimination between an acyl chain disordering and ‘space-filling’ type of mechanism for HII phase formation by these molecules. Dr V. A. Parsegian (National Institutes of Health, Bethesda, MD) replied: (1) Of course, as long as you really know how to transfer to biological situations non-equilibrium information gathered in vitro. (2) Your excellent question is related to that put by Prof. Haydon. As analysed so far, the neutron diffraction studies mentioned in our paper show only that alkanes are entering the lattice in a way that corresponds to HII symmetry. We are now analysing the data to assign a more specific alkane location.We do tend to think of the alkane as filling a space where, lattice dimensions suggest, the phospholipid acyl chains cannot reach without a large expenditure of energy. (However, the idea of a volume that acyl chains cannot fill is an idealization that breaks down on a molecular scale.) There can indeed also be some disordering of acyl chains by diffusely distributed alkane. To go further into the matter, we are investigating a wider range of alkane sizes. Prof. G. Lindblom (University of Ume6, Sweden) said: I have a comment on Dr Parsegian’s paper. We have investigated the ternary phase diagram (for the temperature range 25-50 “C) for the system dioleyl-lecithin-dodecane-water.’ This system exhibits several liquid-crystalline phases, e.g.lamellar, cubic and hexagonal. The most interesting feature of the phase diagram is that there is a transition from the lamellar phase to a reversed hexagonal phase with increasing water content. the phase transition can be conveniently followed by 31P n.m.r. as shown in fig. 4. These results are in good agreement with Dr. Parsegian’s theory for the formation of HII phases in excess water. These findings also have important implications for some of the functions of the biological membrane. Thus for example we have found2 that the bacterium Acholeplasma laidlawii changes the membrane lipid composition in a remarkable way upon addition of dodecane to the growth medium. This is necessary in order to keep the bilayer membrane intact. 1 M. Sjolund, G.Lindblom, L. Rilfors and G. Arvidson, Biophysics, submitted for publication. 2 A. Wieslander, L. Rilfors and G. Lindblom, Biochemistry, in press. Dr V. A. Parsegian (National Institutes of Health, Bethesda, M D ) said to Prof. Fromherz. How can you be sure of a well defined chemical potential after the shattering disturbance of sonication? The question is especially worrying in view of the polydisper- sity shown in your plate 1. Your ‘discs’ presumably have different perimeter-to-area ratios with necessarily different resulting mid-disc cholate concentrations and ‘chemical potentials’ for each disc. Prof. P. Fromherz ( University of Ulm, West Germany) replied: We have to distinguish two processes of quite different time constant. Sonication creates fragments of finite edge tension.This is a non-equilibrium situation with respect to the state of aggregation of the bilayer. In a relatively slow process the system relaxes to the metastable state of closed vesicles. (In a further slow process the system would relax by fusion to extended bilayers, which would be the equilibrium situation again.) We assume that during the slow relaxation the distribution of cholate between edge, interior of bilayer and bulk water is in equilibrium, because the exchange of surfactants is a fast relaxing reaction.72 General Discussion Fig. 4. 101.3 MHz 31P n.m.r. spectra recorded at 25 "C from DOPC-'H,O-n-dodecane mixtures with a DOPC-n-dodecane molar ratio of 1 : 2 and with (a) 14% (w/w), (b) 44% and ( c ) 54% 2H20. Then in the non-equilibrium situations of the dispersion during relaxation from discs to vesicles the chemical potential of cholate is a well defined quantity as determined by the concentration of cholate in bulk water.Owing to the destruction of edge during the closure of discs to vesicles the chemical potential may change slightly during the slow relaxation of the dispersion, implying a slow shift of the distribution equilibrium. Dr L. Fisher ( CSIRO, Sydney, Australia) said: The system egg-lecithin-taurocheno- desoxycholate is known to form large rod-shaped micelles. Could the objects shown in your plate 2 be related to such micellar structures? Prof. P. Fromherz (University of Ulm, West Germany) answered: Considering the periodicity of the pattern in plate 1 we interpret the pattern as stacks of bilayer discs.The stacks are formed by the stain. Before staining these large aggregates do not exist, as indicated by dynamic light scattering. At high concentration of cholate stable rod-shaped micelles may exist. I do not know whether they appear as similar patterns in the electron microscope after negative stain. Prof. J. K. Thomas (University of Notre Dame, ID) said: Sometime ago' we investi- gated the effect of bile-acid surfactants on lecithin vesicles. We found that the bile acid tends to form domains in the vesicle and eventually leads to vesicle rupture and to the formation of mixed micelles. Would these effects in any way affect your model of edge-bile acid stabilization? 1 J. K. Thomas, D. A. N. Moms, F. Castellino and R. McNeil, Biochim.Biophys. Acta, 1980, 599, 380.General Discussion 7 3 Prof. P. Fromherz (University of Ulm, West Germany) replied: Titration of vesicles by cholate leads to a drop of the edge tension, even if no actual edges are present. The edge tension governs the probability of the formation of transient pores. If the edge tension is further lowered by addition of edge actant a pore may grow above a critical size such that rupture occurs. This process of rupture appears before the edge tension vanishes, depending on the elastic modulus and the osmotic pressure. Systematic experiments on the formation of transient pores and on burst in relation to the concentra- tion of emulsification (vanishing edge tension) and to the c.m.c. of various edge actants are in progress.Dr R. Schubert (Chimrgische Klinik, Tubingen, West Germany) said: On the basis of our investigations on detergent-lipid interactions, a few remarks should be made concerning vesicle formation via dialysis and the effect of detergent binding on vesicular membranes. If cholesterol or sphingomyelin is partially substituted for egg-yolk lecithin or if temperature is decreased but still above T, , dialysis of lipid-cholate mixed micelles results in larger vesicles. This effect may be due to changes (i.e. reduction) in the edge tension of the mixed disc micelles. On the one hand, however, we found an increase in vesicle size with increasing dialysis temperature, when C,EO, was used as detergent. Whether the concept of edge activity is also valid for such non-ionic detergents, therefore, remains to be clarified.On the other, the kinetic control of detergent dialysis by reduced cutoff of the dialysis membrane as well as a higher lipid concentration result in larger vesicles, both of which are independent of changes in the edge binding constant. These observations tend to indicate that, before disc-vesicle transition, the size of the largest discs that finally close to vesicles is determined by the frequency of disc fusion. Moreover, the concept of shape transformation from a flat bilayer to a closed vesicle does not adequately explain the higher amount of lipid in the outer vesicle leaflet, postulated on the basis of sterical considerations. Our studies on cholate-membrane interaction clearly demonstrate that the size of the fluctuating vesicle pores induced by membrane-bound detergent can only be small.Large hydrophilic molecules are only released from vesicles shortly after cholate addition by transient pores arising during a sudden membrane foldover.' The rapid closure of these larger pores suggests that the inner edges in membrane pores formed by bile salts are far less stable than the other edges in mixed disc micelles. 1 R. Schubert, K. Beyer, H. Wolburg and K-H. Schmidt, Biochemistry, 1986, 25, 5263. Prof. P. Fromherz (University of Ulm, West Germany) commented in reply: ( a ) The sensitivity to temperature of the structure of dispersions of detergents with headgroups of oligo-oxyethylene is well known. Thus a modulation of edge-activity by temperature is possible. ( b ) During dialysis two processes occur.( 1 ) A reduction of the cholate concentration with concomitant enhancement of edge tension. (2) A growth of the discs with finite edge tension by lateral fusion. In the case of fast dialysis the enhancement of edge-tension is fast as compared to growth. That is to say the discs become unstable at a relatively small size owing to the high edge tension. In the case of slow dialysis the discs have time to grow at a relatively low edge tension. There the instability occurs at a relatively large size. The model of edge activity leads thus to the first rationalization of the correlation slow dialysis/large vesicles and fast dialysis/ small vesicles. The relation has been drawn explicitly in the size us. concentration diagram in our first paper on the subject [fig.4 of our ref. (9)]. ( c ) The theory of disc-vesicle transition developed in our paper and in our ref. (9) treats the bilayer as a two-dimensional liquid with bending elasticity. A phenomenologi- cal model describing the energetics of bending by an elastic modulus cannot explain,74 General Discussion of course, the molecular mechanism of the process. In molecular terms the redistribution of lipid from the inner to the outer monolayer is a natural mechanism to lower the elastic energy. At the open edge this redistribution is a fast process. ( d ) Concerning the formation of pores in closed vesicles, the concept of edge activity apparently describes the instability of edges in closed vesicles with a finite edge tension as compared to the edges of mixed micelles with vanishing edge tension. Dr D.S. Dimitrov (Bulgarian Academy of Sciences, Sofia, Bulgaria) said: There are at least three important findings in Prof. Fromherz's work: (1) demonstration of open discs as intermediates of vesicle formation, (2) estimates of the edge energy and the curvature elasticity and (3) effects of cholate on the rate of vesicle formation. I would like to add that membrane viscosity can contribute to the kinetics of vesicle formation, especially in the cases where there are no activation barriers. This may be the case for formation of giant vesicles, where the curvature elasticity effects can be neglected. We have made a simple estimate for the rate of liposome formation assuming that the driving force is the edge tension and that there are no activation barriers.The forces which resist the edge tension are due to the membrane viscosity and curvature elasticity. Then an approximate balance of forces leads to the following expression for the time of liposome formation time = viscosity x radius/ (edge tension - curvature elasticity/ radius). Using values for the two-dimensional membrane viscosity, curvature elasticity, edge tension and liposome radius on the order of erg, lo-* dyn and 10 pm, respectively, we get times on the order of 100 s. It is seen that the curvature elasticity effect can be neglected for giant liposomes. For small vesicles it is important and determines a minimum radius of the order of curvature elasticity/line tension = cm. Unfortunately, there is no theory for the size distribution.It should be pointed out that for small vesicles the edge energy per disc, which eventually will transform to a liposome, is of the order of thermal energy kT. For giant liposomes, however, if we assume that the mechanism of liposome formation is that of disc-vesicle transformation and the disc edge has the same edge energy as for small vesicles, then the total edge energy per liposome is much larger than the thermal energy. In this case the size distribution should be very narrow which is apparently not the case. Therefore, the effective edge tension is probably very small. This leads to long times of liposome formation. The above formula indicates that the kinetics of liposome formation should depend on the mem- brane viscosity. This dependence may be responsible for the increased rate of liposome formation at high temperatures.In addition, liposomes do not form from lipids in solid state because their viscosity is very high. s.P., Prof. J. F. Holzwarth (Fritz-Haber Institut, Berlin, West Germany) said to Prof. Fromherz: In your paper you discuss the influence of amphiphiles on the stabilisation of bilayer systems with 'open' edges (i.e. the hydrophobic part in contact with water). You also mention briefly the paper by Batzri and Korn [ref. (31)] which gives the first description of the so-called injection method for preparing vesicles. In this method, which was further refined by Kremer et al.' as well as Eck and Holzwarth? an alcoholic solution of lipids is carefully injected into a tenfold volume of buffer solution over a period of several minutes.During the injection process the buffer solution is kept 5-10 "C above the phase transition temperature (T,) of the lipid and carefully stirred. Afterwards the mixture is dialysed against buffer at a temperature of 5-10 "C above T' for at least 8 h to remove the alcohol from the bilayer (vesicle) preparation. In this way very stable preparations of small unilamellar vesicles (SUV) are formed, the size of which varies between 20 and 100nm depending on the final concentration of lipids (typically 1-10 mmol drnp3). If one measures the turbidity of such preparations in thePlate 1. See caption to fig. 6. (To facep. 75)General Discussion I/] h Y d .- * 2 - v 0 c -E 0, 2 1 - 75 ----- - - ---- - 2 - -. -__ ' --- -: -.r\ ' '\ d \ \ I 1 1 I > l o o 90 F 80 70 4 60 D 5 50 40 30 20 10 10 20 30 40 50 60 70 80 90 100 sizelnm Fig. 6. Size distribution of unilamellar vesicles from 2.5 mmol dm-3 dipalmitoylphosphatidyl- choline (DPPC) lipids derived from negatively stained preparations in an electron microscope, shown in plate 1. Total number of vesicles = 121 1; maximum number = 80.3; size at maximum = 23.3 nm; direct width parameter = 1.33. temperature range of the phase transition one gets the results given in fig. 5 . My question is: Can you explain the hysteresis which is found in the presence of 10% ethanol in contrast to the dialysed vesicles by a stabilization of aggregates with edges caused by the alcohol? If such aggregates are dialysed they might lose the alcohol from the edges and stabilize themselves by forming unilamellar vesicles.The second question is: Can the size distribution, an example of which is given in fig. 6 and plate 1, be explained76 General Discussion by the amount of alcohol which is available to stabilize intermediates with edges? An indication of this is that higher concentrations of lipids in the alcoholic solution obtained by keeping the volume ratio buffer/alcohol constant results in larger vesicles. 1 J. M. Kremer, M. W. Eskev, C. Pathmamanocharan and C. Wiersema, Biochemistry, 1977, 16, 3932. 2 V. Eck and J. F. Holzwarth, in Surfactants in Solution, ed. K. L. Mittal and B. Lindman (Plenum, New York, 1984), vol. 3, pp. 2059-2079. Prof. P. Fromherz (University of Ulm, West Germany) replied: From the chemical structure of ethanol one would expect it to be an edge actant.’ However, I have no estimate of its edge activity as compared to the cholates.The size of vesicles is governed by the kinetics of the growth of the lipid aggregates and by the rate of change of the edge tension for a particular recipe of vesiculation. The data available on the method of ethanol injection233 and our own preliminary studies are too incomplete to provide a solid basis for an interpretation. 1 P. Fromherz, Chem. Phys. Lett., 1983,94, 259. 2 S. Batzri and E. D. Korn, Biochim. Biophys. Acta, 1973, 298, 1015. 3 J. M. Kremer, M. N. Eskev, C. Pathmamanochoran and C. Wiersma, Biochemistry, 1977, 16, 3932. Dr R. E. Dale (Christie Hospital, Manchester) addressed Prof.Levine. (1) For DPH in the multibilayer systems you report a qualitative change in the derived orientational distribution function between interpretation in terms of P2( cos p ) and both P,(cos p ) and P~(COS p). How physically realistic can the latter result then be considered, since presumably inclusion of P,(cos p), if it were possible, would result in a further qualitative change in the derived distribution? (2) You have reported heterogeneity of probe lifetimes in these systems. This might arise from heterogeneity of probe environment in the membrane. Would you expect these two putative populations to have different orientational distributions and rotational diffusion coefficients, and is there any possibility of differentiating them, perhaps by time-resolved AFD? In the case of DPH at least, might they be correlated with the ‘parallel’ and ‘perpendicular’ populations indicated by the minima in the derived orientational distribution functions ? (3) More generally, is there substantially more information to be gained from time-resolved as opposed to steady-state AFD experiments? Prof.Y. K. Levine (University of Utrecht, The Netherlands) replied. (1) It is difficult to assess quantitatively the importance of the higher-order param- eters, ( PL) with L > 4, on the reconstruction of the orientational distribution function f( p ) without knowledge of the function one is trying to approximate. There are, however, some indications that knowledge of (P6) and higher terms will not change the form of the distribution substantially.It can be shown on simple mathematical grounds that if f(p) is evaluated up to the Lth order parameter ( L even), it will have at most L / 2 - 1 maxima in the interval 0 < p < ~ / 2 . On the other hand, X-ray diffraction experiments on the orientation of the hydrocarbon chains in oriented bilayer systems indicate the presence of only one maximum in this interval and that for the gel Lpr phase. This supports the view that knowledge of (PJ and (P4) is sufficient to describe the overall features of f(P). The question which remains to be answered is whether this knowledge yields a correct description of the detailed form off@) around /3 = ~ / 2 . I.e. how sure are we that the DPH molecules can lie parallel to the bilayer surface.In the paper we present arguments based on the consistency of this description for a number of systems and the chemical structure of the two probe molecules. One can say that at best this is circumstantial evidence. However, we can also note that the longer, but chemically similar, p-carotene molecules are also found to have a propensity to lie parallel to theGeneral Discussion 77 bilayer plane in many membrane systems. This behaviour is quite clearly demonstrated by linear dichroism experiments' which only yield (P2) and resonance Raman experi- ments2 which yield both (P2) and (P4). (2) We have been unable to find any correlation between the heterogeneity of probe lifetimes and a putative heterogeneity in the probe populations. In fact, our results indicate that the steady-state fluorescence depolarization experiments are essentially sensitive only to the long-lifetime component of the fluorescence decay. This arises simply from the small contribution of the short-lifetime component to the total time- integrated fluorescence intensity.Thus the heterogeneity indicated is associated in its entirety with the long-lifetime component for both DPH and TMA-DPH molecules. It is important to note in this context that our analysis is based on the assumption of a dynamically homogeneous probe population. The fact that for DPH one obtains an orientationally heterogeneous population strongly suggests that here our description of the probe behaviour is oversimplified. The point is that steady-state AFD experiments do not yield sufficient information to resolve this problem.Time-resolved experiments, however, may provide the answer. (3) In principle time-resolved AFD experiments provide means for following directly the decay of the time autocorrelation functions Gk( t ) . In this sense they are potentially more interesting than steady-state measurements, which only yield the time-integrals of these functions. For this reason we are currently carrying out such experiments with the synchrotron radiation source in Daresbury. Our expectations, however, have taken something of a knock as a result of the cumbersome analysis of the single-photon counting experiments. The main problem being the use of iterative least-squares methods for the deconvolution of the data and the setting up of statistical criteria for the goodness-of-fit.So far we have been able to show that the order parameters obtained from the time-resolved experiments are in good agreement with those obtained from steady-state ones, but are having difficulty in obtaining the kinetics of the decays in a consistent manner. This work is being continued. On the other hand, we have shown that steady-state AFD experiments coupled with measurements of the fluorescence decay afford a convenient, reliable and fairly quick method for obtaining quantitative informa- tion about the dynamic behaviour of the probe molecules. 1 B-A. Johansson, G. Lindblom, A. Wieslander and G. Arvidson, FEBS Lett., 1981, 128, 97. 2 M. Van de Ven, M. Kattenberg, G. van Ginkel and Y. K. Levine, Biophys. J., 1984, 45, 1203.Prof. G. Lindblom (University of Umea", Sweden) said: I would just like to point out that we have found that the lateral diffusion coefficient of lipids in lecithin bilayers does not depend on the molecular order (e.g. by increasing the cholesterol concentration).' I think that this supports Prof. Levine's ideas. 1 G. Lindblom, B-A. Johansson and G. Arvidson, Biochemistry, 1981, 20, 2204. Prof. J. F. Holzwarth (Fritz Haber-Institut, Berlin, West Germany) asked Prof. Levine: In your paper about e.s.r. and angle-resolved fluorescence depolarization ( AFD) experiments you mention the influence of probes on the bilayer structure. In fig. 7 we have demonstrated this influence for DMPC and could show that even at 1/250 ratios there is still a clear reduction of the transmission at temperatures above the phase- transition temperature T, , even if the normalized order-parameter/ temperature depen- dence still shows inside 0.5 "C the expected midpoint T, for pure lipids, as is demon- strated in fig.8. Another problem which is associated with the AFD measurements is that you sense the time range of nanoseconds, but the major part (80% with respect to AHHpT) of the phase transition occurs in the ps to ms time regime, as we have demon- strated' (see also the following contribution).78 General Discussion h W c -e 0 -2 a v - 0.1 1 I I I 15 20 25 T / "C Fig. 7. Turbidity us. temperature dependence of unilamellar vesicles of dimyristoylphos- phatidylcholine (DMPC) containing different concentrations of the probe molecule diphenyl- hexatriene (DPH): (---) 1/135, ( - - - ) 1/250, (-) none. 2.7 mmol dm-3; A = 300 nm. R == 50 nm; [DMPC] = I I 20 25 T / "C Fig. 8. Normalized phase transition us. temperature dependence of fig. 7: (- - -) 135/1, T, = 23.5 "C; (-) 500/1, T, = 24.2 "C. This disadvantage of fluorescence polarization lifetime measurements, and also with respect to the time range of the e.s.r. experiments, can be overcome by using the time dependence of the fluorescence anisotropy as we did between 5 ps and 100 ms.' My question is how important is the influence of the probe molecules with respect to the disturbance of the bilayer structure, especially if one senses the immediate environment of the probe? 1 A. Genz and J. F. Holzwarth, Eur. Biophys. J., 1986, 13, 323. 2 J. F. Holzwarth, V. Eck and A. Genz, in Spectroscopy and Dynamics of Molecular Biological Systems, ed. P. Bayley and R. Dale (Academic Press, London, 1985), pp. 351-377.General Discussion 79 Prof. Y. K. Levine ( University of Utrecht, The Netherlands) replied: The experiments reported in the paper were carried out at temperatures well above those of the phase transitions of the various lipid systems. Control AFD experiments yields the same fluorescence depolarization ratios for probe/lipid ratios between 1/ 1000 and 1/ 125. The ratio of 1/250 was chosen arbitrarily for experimental convenience. Similarly, no dependence of the e.s.r. lineshape on the probe/lipid ratio was observed over the same range. As we carry out our experiments above the phase transition we expect the probe molecules to be uniformly distributed in the lipid multibilayers. We have no experimental evidence of clustering or phase-separation processes which may well play a role around the phase-transition temperature of the bilayers. The point we are making is that two probe techniques, utilizing different probe molecules, yield the same information about the orientational order and dynamics in a variety of lipid bilayer systems. To my mind this strongly suggests that the perturbation of the bilayer structure by the probe molecules is much less important than has been suggested.

ISSN:0301-7249    

DOI:10.1039/DC9868100063

出版商:RSC    

年代:1986

数据来源: RSC

摘要:

Faraday Discuss. Chem. Soc., 1986,81, 81-94 Studies of Membrane Heterogeneity using Fluorescence Associative Techniques Lesley Davenport Chemistry Department, City University of New York, Brooklyn College, Brooklyn, New York 1121 0, U.S.A. Jay R. Knutson Laboratory of Technical Development, National Institutes of Health, Building 10, Room 5D-10, Bethesda, Maryland 20892, U.S.A. Ludwig Brand The Biology Department and the McCollum- Pratt Institute, The Johns Hopkins University, Baltimore, Maryland 21218, USA. Fluorescence spectra, decay times and emission anisotropy are capable of providing important information regarding the interaction of probes with biological macromolecules. Combinations of different fluorescence para- meters can provide information about heterogeneity of liposomes or biologi- cal membranes and about the character of excited-state interactions of the fluorescence probe.Decay-associated spectra (DAS) or anisotropy decay associated spectra (ADAS) can be used to resolve heterogeneous species. These methods have been applied to the study of both pyrene and a pyrene methyl cholesterol adduct (PMC) with dimyristoyl-lecithin vesicles (DML). The data indicate microheterogeneity in the distribution of the probes above the phase transition. Anisotropy decay-associated spectra (ADAS) have been used to study diphenylhexatriene (DPH) in dipalmitoyl-lecithin (DPL) and dimyristoyl-lecithin (DML) vesicles. The results indicate that DPH inhabits more than one rotational environment in the liposome preparations used. In recent years lipid vesicles have received much attention as model membrane systems.‘ , 2 It has long been recognized that biological membranes exhibit structural heterogeneity in the lipid packing akin to a ‘patchwork q ~ i l t ’ . ~ Near the phospholipid phase transition temperature, even single component lipid vesicles are expected to consist of both gel and fluid domain^.^-^ The size of these domains will naturally be dependent on tem- perature and degree of structural cooperativity. While many model membrane systems have been interpreted within a domain framework: less progress has been made in the area of directly detecting and/or quantitating the different lipid fractions. In steady-state fluorescence probe studies of model membrane systems, the domain concept has been invoked many times,*-’* yet time-resolved emission anistropy studies continue to be interpreted via homogeneous’ (‘wobbling-in-a-cone’ 12*13) models.One reason for this ambiguity of interpretation is that only a single type of measurement is being considered. Several types of measure- ments can, however, be made. These include excitation and emission spectra, quantum yields and decay times, steady-state and time-resolved emission anisotropy as well as susceptibility to quenching or other excited-state interactions. It is significant that these different types of measurements may be associated 14-16 to obtain more information than is available from individual experiments. The combination of the associative 8182 Fluorescence Associative Techniques methodology with ‘global’ I7-l9 analysis procedures provides powerful tools for un- ravelling the fluorescence of complex systems.Fluorescence spectroscopy has been used this way to subdivide the heterogeneity found in the intrinsic fluorescence of proteins. 20,21 The use of decay-associated spectra (DAS)’4,22-25 have been of value in this regard, since the association between decay times and spectral distributions greatly aids in the assignment of specific emission decay constants. There are other cases where it may be more appropriate to associate rotational decay rates with unique spectral distributions (anisotropy decay-associated spectra; ADAS) .26927 In this paper we describe applications of these new spectral association tools to the study of liposomes. Pyrene is known to form excimers both in solution and in bilayer~.~’-~l In lipid vesicles this decay can be more com lex than expected for a system undergoing a simple two-state excited-state rea~tion.~ -34 In addition, complex fluorescence decay is described for a pyrenemethylcholesterol adduct (PMC).The use of DAS leads to a reasonable interpretation of the observed results in terms of a heterogeneous compartmental model for the different excimer forming systems. Also, the use of ADAS for DPH in vesicles can provide an explanation of the ‘ra’ term in the context of microheterogeneity. !? ADAS DAS DML DPH DPL E/ M PMC SAS Tri s HCl TLC TRES Abbreviations anisotropy decay-associated spectra decay-associated spectra L-a-dimyristoyl-lecithin l76-diphenyl-1,3,5-hexatriene L-a-dipalmitoyl-lecithin ratio of excimer to monomer fluorescence intensities (480 nm/397 nm) 1 -pyrene-methyl-3/3 - hydroxy-22,23-bisnor-5-cholenate species-associated spectra tris( hydroxymethy1)aminomethane - HC1 thin-layer chromatography time-resolved emission spectra Theory Homogeneous vs.Heterogeneous Origins of Complex Excimer Kinetics In solution, complex formation in the excited state often follows simple two-compartment kinetics, as shown in fig. 1. The lifetimes in such a system are given by 71 , 72 = 2/{ (x + Y ) f [ (x - Y)’ + ~ ~ B A [ A ] ~ A B ] ”*}. (1) We have adopted the convention T~ < 72 for later identification. Since only monomer is initially excited, the excimer decay is an equal-magnitude (pl = p 2 ) difference of exponentials (0 at time zero).The negative amplitude of the pair is thus always associated with T ~ , since the total steady-state intensity of B” is given by l o m I B ( A , t ) dt=-P1~1+&72>0. The details of a-p-7 linkage under various conditions has been Basically, the dominant cy term can be a1 or a’, depending on whether X > Y or Y > X, respectively. In the case of ‘irreversibility’ (kAB relatively small), the minor a willL. Davenport, J. R. Knutson and L. Brand 83 A A+A Q) -0 J 0.2 .* - a 5 -0.0 - 0 . 4 ' ' ' ' ' 380 420 460 500 540 wavelength/nm 0 . 6 r a. (11 1 4 0.4 "a 0.2 1 * .d 3 p -0.0 2 -0.2 .- Y d 1 1 . . -0.4'L ' ' ' ' ' ' ' 1 380 420 460 500 540 wavelength/nm Fig. 1. Left-hand side: kinetic scheme for monomer and excimer fluorescence. The monomer species in the ground-state, excited-state and the excimer species are indicated by A, A* and B*, respectively.Radiative rate constants for A* and B* are given by kA and k,, respectively. The rate of conversion between A* and B* is given by kBA[A] and dissociation of B* to A* by /CAB. The combined rate constants, X and Y , for depletion from the A* and B* states are indicated. The decay of the fluorescence for species A* and B* is expressed as a biexponential law with pre-exponential terms a l , a2 and P I , P2: respectively, and lifetimes T~ and T~ for both species. Right-hand side: simulated decay-associated spectra describing two contrasting schemes for depletion from the A* and B* states for reversible excited-state excimer formation. The DAS are defined here as pre-exponential terms ( a l , a2, P I , p2) for a given fluorescence lifetime ( T ~ , T ~ ) as a function of the emission wavelength.In case I the combined rate constant Y describing depletion from the B" state is greater in magnitude than X, the combined rate constant for depletion from the A* state. This leads to the characteristic decay-associated spectra shown with a biexponential fluorescence decay function across both the A* and B* regions of the emission spectrum. In case I1 X > Y. Under these conditions the longer-lifetime component has a smaller absolute contribu- tion in the A* region of the spectrum. For both models the negative pre-exponential term is associated with the shorter-lifetime component. [(-) long T, (-O-) short 7.1 essentially vanish.The summation of (Y and p across the spectrum to form DAS [here DAS, = (Y,(A) +p,(A)] is shown in fig. 1 for two different, slightly reversible, cases. Under conditions of increased viscosity, one must consider the appearance of transient (diff usion-limited) kinetics, since the average concentration gradient around an excited monomer takes some time to e ~ t a b l i s h . ~ ~ The effect of transient (non- equilibrium) rates is a time dependence in kBA[A] that can be approximated as kgA= a + b / d t . This result for a three-dimensional fluid might take on a different character for situations where a two-dimensional surface geometry of diffusion might dominate.29 A more complex hybrid (tensorial) diffusion process is probably at work, given the substantive thickness and depth-dependent dynamics37 of the bilayer hydrocarbon slab.The homogeneous approach, while useful, is dominated by the concept of a uniform vesicle. We have chosen to examine the competitive concept of a 'patchwork'84 Fluorescence Associative Techniques (heterogeneous) system. Homogeneous sources of decay complexity have been pre- viously studied in detail. Thus, we chose to decompose the vesicle dynamics into a superposition of simple compartments. Theory of Anisotropy Decay-associated Spectra The definition of emission anisotropy, r, is a linear average: where y = Iparallel - Iperpendicular is the polarized difference decay, di is the total intensity decay function (if di is in fact independent of i, the system is said to be non-associative, since the index i relates to rotating species).The ai are spectra of each rotating species, and ri are their corresponding anisotropy decays, e.g. ai = ai(A) and ri = ro,i exp ( - t / & ) (3) where and +i are initial anisotropy and rotational correlation of the ith species, respectively. Although ADAS can be derived for associative decays [with r + 4 linkage, see ref. (25) and (26)], we will concentrate here on the simpler, non-associative case. In that case, even if a slowly rotating (large, bound or ‘immobile’) and rapidly rotating (small, free or ‘mobile’) fraction coexist in the measurement volume, total intensity decay -will be wavelength-independent: The difference decay, however, will be heterogeneous: Y(A7 t ) = Iparallel- Iperpendicular Instruments have a finite response function, so the actual observed difference decay is determined by the convolution of time response with lamp function L: Y(A, t ) = C ( Y ~ ~ I = C C U ~ ( A ) d ( t - t ’ ) r i ( t - t ’ ) L ( t ’ ) d t ‘ . (6) I 1 I: An example of two different Y, is shown in fig.2. The mobile difference is all but absent after channel 123, while the immobile probes remain partially aligned and thus exhibit a longer difference decay. Note that the ‘slowest’ difference decay curve possible is that of the total decay (when 4 + 00, yi -+ r&). The division of the difference decay into ‘time windows’ provides an easy way to obtain ADAS (spectra of each rotating species). An anisotropy-decay analysis from one (or a few) wavelengths provides ri and d (and hence yi and K).Knowing Y, functions (like those in fig. 2) we can predict how much of each species is present in a windowed (kth window) difference TRES, i.e. whereL. Davenport, J. R. Knutson and L. Brand 85 0 .o MCA channels (0.205 ns per channel) Fig. 2. Procedure for the determination of time windows a + b and c + d used to extract ADAS from time-resolved polarized emission spectra. The time-resolved emission anisotropy for DPH incorporated into a mixture of separately labelled DML and DPL vesicles ( 1 : 1 mixture) is defined by a biexponential decay law with one correlation time of ca. 2 ns and a limiting anisotropy term (roo). Yl is the synthesized difference decay profile attributed to the faster rotating species. Similarly, Y2 is the synthesized decay of the difference of the polarized intensities attributed to the ‘immobile’ fraction. A ‘time-window’ was chosen such that the late time-gated difference (TRES) derived from experimentally obtained polarized emission spectra [ Gl,(A, t) - IHv(A, t)] contain photons emitted from the more slowly rotating species only.Each channel spans 0.205 ns. Excitation was a 355 nm and emission at 430 nm. The temperature was 29 “C. is the area of each component in the window bounded by channels ak and bk. Notice the matrix form of the mixing: where Mik = [ x ] k . can solve for N unknown ADAS: If one collects N different TRES, and then calculates the area matrix Mik, one a, =I GJ,T j where G is the matrix inverse of M. In some cases an even simpler approach suffices.For the two component case seen in fig. 2, almost no mobile contribution is present in the late window, so cyslow is just a multiple of that difference TRES. For pulse fluorometers this is the easiest ADAS to observe. If +fast is small, even the steady-state difference will be dominated by aslow.26 Thus the sometimes-elusive ‘bound’ spectrum for a system of small ligands binding to macromolecules is, in principle, fairly easy to measure. Focussing on the issue of ‘rotation in a cone’ vs. heterogeneity, it is clear that the previously defined emission anisotropy, might be interpreted either way. If, however, one observes wavelength dependences in the pi, it becomes necessary to consider an ADAS approach. Segmental rotation theory,86 Fluorescence Associative Techniques whether based on cones or more general angular potentials, assumes a single homogeneous probe distribution and would not predict spectral ties to mobility.Proposed Method to Obtain Rapidly some Types of ADAS using Commercial Phase Fluorometers Suppose a system with little spectral variation of lifetime but disparate (e.g. ‘free’ us. ‘bound’) correlation times is studied. The total (Iparallel+ 2Iperpendicular or magic angle) intensity signal will decay according to d(t)=Ccf;exp(-t/Ti) (12) where C J; = 1. Since the T~ do not associate with spectra, fi: are (approximately) independent of wavelength. Even if the (N) different rotating species in the system have different spectra ai, the total decay surface N I ( A , t ) = 2 aj(A) d ( t ) = T ( A ) d ( t ) j is simply the average spectrum-average decay product.The polarized decay surface, however, is where 8 is the angle between the polarizer and vertical (for a vertical polarizer in the other channel), P2(x) = (3x2 - 1)/2, ro,j and 4j are initial anisotropy and correlation time of thejth species, respectively, and the other terms are defined above. The extremes Iparallel and Iperpendicular correspond to P2 = 1 and P2 = - 1/2. Suppose one observes this system with a phase fluorometer capable of lock-in suppression. For Isso (magic angle) the value of P2=0 implies that a phase angle t,b characteristic only of d will be observed (approximately wavelength-independent). If one locks onto a signal orthogonal to this (I) f 90’) the ‘total’ spectrum will be suppressed.The rotation of the polarizer away from this null at magic angle should then provide spectral contributions from probes that are rotating during the decay process. Immobile probes, however, retain the same t,b set at magic angle and are still suppressed. Continued rotation of the polarizer will alternate the sign of these mobile contributions, as P2 can range from - 1/2 to I . ~ * Interestingly, this means the phase system provides a sort of ‘complementary capabil- ity’, since in experiments with pulse instruments the immobile ADAS is usually the easier of the two to obtain. Experimental L-a-Dimyristoyl-lecithin (DML) and L-a-dipalmitoyl-lecithin (DPL) were purchased from the Sigma Chemical Co. and used without further purification after t.1.c. analysis. Pyrene (Eastman Kodak) and PMC (Molecular Probes) were used without further purification after fluorescence intensity decays in hexane revealed a single-exponential decay law.Vesicles were prepared as described in detail Briefly, sonicated vesicles were centrifuged at 108 OOOg for 1 h prior to labelling with DPH as described elsewhere. 27i40241 Pyrene and PMC labelled single bilayer vesicles (along with cholesterol additions) were carried out via probe-lipid cosonication prior to centrifugation. Cosoni- cation of DML and DPL (unlabelled) provided a vesicle fusion control (data not shown). Lipid concentration was determined by lipid pho~phorus.~~L. Davenport, J. R. Knutson and L. Brand 87 Steady-state measurements of fluorescence anisotropies and spectra were carried out as described elsewhere.27y40941 Excitation and emission bandwidths of 4 and 3 nm, respectively, were used for collection of emission spectra vs.3 and 4 nm for excitation spectra. All spectra were carried out using ‘magic-angle’ polarizer geometries to avoid rotation artifacts in the measured total intensity c011ected.~~ Time-resolved measurements of fluorescence decays, emission anisotropies and emission spectra were carried out as previously d e ~ c r i b e d . ~ ~ ~ ~ ~ , ~ ~ Total-intensity (‘magic- angle’) DAS of pyrene and PMC were obtained from a ‘global’ data analysis Complete decay curves at four emission wavelengths (397, 410, 480 and 520 nm) were analysed together to obtain the common set of exponentials adequate to satisfy all of the data curves.Curves with different times per channel calibration were also analysed together to enhance the overall resolution. The amplitude spectra (DAS) were then obtained from partial decay curves ( 5 min acquisition time) at 2 nm intervals. The lifetimes in this last analysis were fixed to the previously determined ‘global’ values. An emission bandwidth of 13 nm was employed. Corrections for colour shifts of the detection system and the instrumental G factor were assessed as described in detail elsewhere.43 In contrast, polarized TRES were obtained by a ‘time-windowed’ scanning procedure described earlier. 14326 Basically, the photon counts accumulated in a polarized decay curve, collected for a brief period (typically ~ O S ) , were bunched according to a time channel interval and accumulated across an emission scan (in 1 nm steps).This process provides simultaneous acquisition of several different gated spectra. These windowed TRES are multiplied by appropriate to mix them properly into ADAS (see Theory section). The G-factor c o r r e ~ t i o n ~ ~ ’ ~ ~ was close to one across the emission wavelength regiofi of interest and was determined as described in detail elsewhere.27 Results and Discussion This section is divided into two subsections. In the first we consider different possible probe locations based on differences in lifetimes. Here we have looked at the nanosecond time-resolved fluorescence decay profiles of both pyrene and a pyrenemethyl cholesterol adduct incorporated into DML vesicles. The latter fluorophore has further utility as a probe of cholesterol heterogeneity.From these systems decay-associated spectra giving insight into the heterogeneity of probe location were extracted. In the second part we discuss heterogeneity of probe location based on different rotational constraints placed on the environs of the probe. Here we describe studies we have performed using the fluorescent probe diphenylhexatriene, which is known to have more than one rotation rate from previous time-resolved emission anisotropy studies on vesicles. Evidence for at least two probe ‘fractions’ based on differences in emission spectra of each rotational motion is presented. Lifetime Associations Typical fluorescence emission spectra for both PMC and pyrene separately incorporated into DML vesicles (1 : 50, probe to phospholipid molar labelling ratio) both above and below the phospholipid phase transition temperature are shown in fig.3. For both probes incorporated into bilayers a monomer emission spectrum is observed at shorter wavelengths that has detailed vibronic structure. At longer wavelengths a broad, struc- tureless emission band arising from the excimer is detected. For both excimer-forming probes, an increase in temperature gives rise to an enhanced excimer fluorescence. In the case of pyrene, minimal shifts and changes between intensity of the vibronic peaks (monomer emission) are observed for pyrene with increasing temperature. Vibronic shifts are observed for PMC when embedded in fluid vs. gel phases (fig. 3). Intensity and spectral changes in the 0-0 vibronic band have previously been reported for pyrene88 Fluorescence Associative Techniques 360 440 520 6 OC wavelength/nm wavelength/ nm Fig.3. Emission spectra for ( a ) pyrene and ( b ) PMC incorporated into L-a -dimyristoyl-lecithin vesicles in 0.01 mol dm-3 Tris- HCl at pH 8.5 containing 0.1 mol dm-3 NaCl at 6 (-) and 24.5 (-.---) "C. The vesicles were labelled 1 : 50 (PMC to phospholipid molar labelling ratio). Excita- tion and emission slits were 4 and 3 nm, respectively. The excitation wavelength was 340 nm. Background vesicle scatter was subtracted using an unlabelled vesicle sample of identical phos- pholipid concentration. incorporated into both pure egg lecithin and DPL vesicles,46 at a higher spectral resolution than used here.They have been attributed to an altered polarity of the environment around the probe. The excitation spectra for both pyrene and PMC systems at this temperature show superimposition when using monomer and excimer emission wavelengths. With increasing temperature, the ratio of excimer to monomer (E/ M) fluorescence intensities shows an increase for both pyrene and PMC labelled DML samples. In all cases, however, the integrated fluorescence intensity decreases with increasing tem- perature. Such effects may arise from an enhanced quenching by oxygen at higher t e m p e r a t ~ r e . ~ ~ Addition of increasing mole fractions of cholesterol (20% and 33 mol%) results in a decreased E/M intensity ratio for pyrene labelled vesicles, the effect being the greatest for the highest cholesterol concentration. In contrast, for PMC labelled DML/ cholesterol samples, increasing cholesterol concentrations give rise to an enhanced E/ M intensity ratio, indicating a higher effective concentration of excimer species.This may reflect a non-homogeneous distribution of PMC in the bilayer, with local concentra- tions of the probe occurring within 'cholesterol-rich' regions. The distribution of cholesterol is thought to be non-random within the bilayer m a t r i ~ . ~ ~ - ' ~ Time-resolved fluorescence measurements were performed for PMC and pyrene- labelled DML vesicles in the absence of additional cholesterol. Fluorescence decay curves for the samples (15 000 counts at the peak) at both monomer (397 annd 410 nm) and excimer (480 and 520 nm) emission wavelengths were collected. Collection of the decay data at two timing calibrations (0.2 ns per channel and 2 ns per channel) ensured accurate estimates of both short- and long-lived components.Using a 'global' non-linear least-squares fitting procedure, l 7 - I 9 a consistent set of decay parameters were obtained for the eight data sets, across the emission bands of both monomer and excimer species. The results are reported in the legend to fig. 4. For pyrene the decay data were best expressed using a triple-exponential decay function. For PMC-labelled samples, a four-exponential decay was required. At longer wavelengths (480 and 520nm) in the spectral region of excimer emissions, it is noted that for pyrene a negative pre-exponential term is associated with the 34 ns componentL.Davenport, J. R. Knutson and L. Brand 89 W a U .- - a s 0.40 r ! '. 0.20 ! ...... ,*..a. .... .... ............... ....' I -0.101 ' 1 1 1 1 1 ' 1 1 1 1 I I I ' I J 380 420 460 500 540 wavelength/nm 0.40 W .: 0.00 - 2 380 420 460 500 540 wavelength/ nm Fig. 4. Decay-associated fluorescence emission spectra (DAS) for (a) pyrene and ( b ) PMC-labelled DML vesicles (1 : 50 probe to phospholipid molar labelling ratio) at 30 "C. For pyrene the decay components were determined to be (-.---) 128.2, ( - - - . -) 33.9 and (-) 8.4 ns. For PMC a minimum of four decay constants were required: (-.-.-) 82.2, (- - - - -) 27.2, (- - -) 8.7 and (-) 120.5 ns. For both molecules the negative pre-exponential terms are associated with the short decay components.Excitation was 340 nm, with excitation and emission bandwidths of 13 nm each, respectively. Global ,y2 values were 1.05 for pyrene and 1.05 for PMC. with a magnitude almost equal (except for the presence of spectral overlap) to the pre-exponential associated with the 128 ns component. These decay characteristics are diagnostic of a two-state excited-state (excimer) reaction.29730 In the case of PMC, in the excimer band, two negative pre-exponential terms are obtained from the decay profiles (the other two are positive). Decay-associated spectra (DAS) were obtained from decay curves collected at 2 nm intervals (ca. 2000 counts at the peak) across the emission wavelength region of interest. A 100-curve global analysis was used to extract the pre-exponential associated with a particular lifetime as a function of emission wavelength (DAS).Decay times, accurately determined from the eight-curve analyses of higher peak count, were also fixed in this multi-curve analysis. The DAS thus determined are shown in fig. 4. For pyrene-labelled vesicles [fig. 4(a)] DAS for three lifetimes are shown. For pyrene-labelled vesicles, the derived DAS for the 34 and 128 ns components suggest a reversible Y > X mechanism (fig. 1). In the excimer band of the emission the amplitude components are equal in magnitude but opposite in sign, characteristic of an excited-state population which builds up (initially zero) and then decays away. The shorter lifetime is always associated with the negative pre-exponential term. At shorter wavelengths, the pre-exponential terms for both lifetimes are evident. This indicates reversibility of excimer formation.30y35 Interestingly, the third DAS, associated with the very-short-lifetime component (8.4 ns), reveals a characteristic monomer spectrum and no excimer emission at longer wavelengths. The strongly quenched lifetime of this second pyrene population may be the limit to excimer formation.One may speculate as to why the lifetime of this fraction is so short. It is likely that a transverse distribution of pyrene, like other small hydro- phobic molecule^,^^ will span the bilayer. Under such circumstances the probability of locating some pyrene molecules at the polar head group is high. In a polar environment the lifetime of this apolar probe is likely to be quenched.In addition, the molecular motions of the fatty acyl chains are reduced near the head group region.37 Consequently, the diffusion of pyrene molecules to form excimers might be further hindered. Hence, this separate 'quenched' monomer population seems reasonable. A more complete study90 Fluorescence Associative Techniques of this fraction vs. lipid composition and temperature (especially for gel/ fluid redistribu- tion) is clearly in order. For PMC, the DAS [fig. 4(b)] reveal added complexity. Analogous with pyrene, one pair of the DAS (for the lifetimes of 82.2 and 27.211s) reflect excimer formation with the kinetics Y > X (i.e. rate of loss from the monomer species is less rapid than from the excimer complex). In addition, however, a second negative DAS is seen, associated with a very short (8.411s) lifetime. To reconcile these kinetics, we suggest that excimer formation proceeds at two different rates in two different ‘domains’ or classes of binding sites.In one region, the PMC have a high ‘local concentration’ (clustering or proximity), so excimer formation (hence X ) would be very fast. In the second region X would be smaller, since excimer formation would occur via the same random-walk that pyrene performs. If the dissociation/deactivation rate ( Y ) were the same for both regions, three exponentials would be predicted and the two negative DAS (in the B” emission band) would mirror the sole positive DAS there. Further, the small X and the common Y would resemble those seen for pyrene. All of these features are, in fact, present in three of the DAS (fig.4). In addition to the two different excited-state reactions suggested by these DAS, a fourth lifetime component is associated with a ‘monomer’ spectrum. Unlike the pyrene-labelled system, the lifetime for this spectrum (120.5 ns) is not quenched. Since the pyrene group is conjugated through the alkyl chain of the 17 position on the amphiphilic cholesterol, it might be expected that the pyrene group of this probe is ‘anchored’ in the hydrophobic part of the bilayer and is not affected by the polar-region quenching suggested for pyrene. It is pertinent to note that we have preferred to interpret the time-resolved lifetime data for both pyrene and PMC in terms of a heterogeneous or multicompartment distribution of molecules.We realize that this concept represents only one possible view of the true process. A combination of X > Y and Y > X excimer forming kinetic mechanisms might equally be represented by transient kinetics, where the fast term (equivalent to the X > Y flow) originates from the early transient (fast non-equilibrium diffusion) term and the ‘slower’ rate of excimer formation ( Y > X ) arises from the equilibrium diff usion-controlled excimer formation. An interpretation for the additional term (an isolated, long-lived, monomeric DAS) using this transient kinetic approach is more difficult. A more appropriate model is probably a combination of these two possible views. Further information will be required to distinguish between these two kinetic schemes. Species associated (species DAS) may prove useful in such studies.For a uniform transient kinetic mechanism it is expected that the SAS for the transient and equilibrium-diff usional species will be totally superimposed. In contrast, and as a result of the environmental sensitivity of the pyrene group of PMC, heterogeneity of probe location is likely to result in vibrational shifts of the highly resolved monomer emission (cJ fig. 3). In this case derived SAS for the probe subpopulations should reveal shifts. Higher spectral resolution DAS will be required in order to conclude this issue. While a variety of compartmental models could be used to explain the four exponen- tials observed in this study, the association between decay times and spectral distribution favours the microdomain interpretation we have presented here.Rotational Heterogeneity A rotationally heterogeneous model system was prepared from a 1 : 1 mixture of DML and DPL vesicles separately labelled with DPH. At 29 “C, DML vesicles are in the fluid phase (transiton temperature is 23 0C52), whereas DPL is below its phase-transition temperature (41 0C53,54), so DPH in the latter lipid is located in a more restricted phase. The DPL-DML vesicle mixture showed a biphasic phospholipid phase transition on measuring the steady-state emission anisotropy of DPH as a function of increasing temperature. This suggested that there is minimal fusion of the single bilayer vesiclesL. Davenport, J. R. Knutson and L. Brand 91 1.0 0.8 0.6 0.4 0.2 0.0 370 410 4 50 49 0 530 570 wavelength/nm Fig.5. Anisotropy decay-associated spectrum for a DPH labelled 1 : 1 mixture of DML and DPL vesicle preparations at 29 "C. The steady-state emission spectra for the individual (- - -) DPL and (---.-) DML vesicles and for the 1 : 1 vesicle mixture (-) are shown. The anisotropy decay-associated spectrum (-A-) corresponding to the 'immobile' probe fraction shows superim- position with the DPL steady-state spectrum. The obtained parameters for r( t ) for DPH mixed vesicles were recovered in terms of a single-exponential decay plus a constant ( p = 0.13,4 = 1.56 ns and r,=0.21, x2 = 1.53). The parameters obtained for d ( t ) for the DPH mixed-vesicle system best fit a double-exponential decay function ( fl = 0.233, T~ = 5.46 ns, fi = 0.767, T~ = 10.26 ns, x2 = 2.34). Excitation was at 355 nm and emission was 430 nm with bandwidths of 14 and 9.9 nm, respectively, for ADAS measurements and 14 and 7 nm, respectively, for time-resolved anisotropy measurements.The molar labelling ratio was 1:500 (probe to phospholipid). A value of 0.23 was measured for steady-state emission anisotropy of DPH in the lipid mixture. during the time course of the experiments. In contrast, a co-sonicated phospholipid sample revealed a broad transition with a phase transition at ca. 32°C. This is in agreement with previous studies.55 It is known that small unimolecular vesicles undergo a spontaneous change in size owing to fusion and/or aggregation, especially when incubated below the phase-transi- tion t e m p e r a t ~ r e .~ ~ - ~ ~ It is not known whether macroheterogeneity, if present in our preparations, is related to the microscopic heterogeneity discussed here. The steady-state emission spectra for DPH in the two separate vesicle samples are shown in fig. 5 . There are subtle spectral changes between the two vesicle samples.6o The DPL-DML mixture provides an emission spectrum which lies intermediate between the component spectra. The decay of the emission anisotropy for DPH incorporated into the mixed DML-DPL vesicle sample was best expressed in terms of one correlation time plus a constant or residual (roo) term (see legend to fig. 5 ) . As described more fully in the Theory section, a late window was chosen such that the difference decay associated with the more mobile fraction was essentially excluded.Only photons associated with the immobile fraction will be seen in that difference TRES. This ADAS extracted for the 'immobile probe fraction' shows excellent superimposition with the component DPL spectrum, as expected. In the case of DPH labelled DML vesicles at 25 "C, in the region of the phospholipid phase transition, where both solid and fluid phases are expected to coexist, the ADAS obtained from the early and late time windows (fig. 6 ) are also significantly different. In contrast, the total intensity spectra, obtained within the same broad windows, showed no spectral differences (data not shown). Hence lifetime associations with rotation rates cannot account for the observed spectral shifts. The ADAS corresponding to the92 Fluorescence Associative Techniques 1.0 - 0.8 - 0 0.6 - 2 .- ti2 0 5 0.4 - .- m c Y .- 0.2 - 0.0 370 410 450 490 530 57 0 wavelength/nm Fig.6. Anisotropy decay-associated spectrum for a DPH-labelled DML vesicle sample at 25 “C. The ‘late’ difference spectrum (-A-) represents directly the ADAS of the ‘immobile’ probe fraction in the system. The early difference (-) includes contributions from both mobile and immobile probes. The decay of the total fluorescence emission fits best a double-exponential decay function (fi = 0.106, T~ = 1.18 ns,f2 = 0.894, T~ = 9.09 ns, x 2 = 1.64). The parameters obtained for r( t ) were best expressed in terms of a single-exponential decay function plus a constant term ( p = 0.21, 4 = 3.03 ns, roo = 0.15, x2 = 1.20).Excitation was at 355 nm and emission was at 430 nm, with bandwidths of 14 and 9.9. nm, respectively, for the ADAS measurements and 14 and 7 nm, respectively, for the single-curve measurements. The molar labelling ratio was 1 : 500 (probe to phospholipid). The value of the steady-state emission anisotropy was 0.20. ‘immobile’ probe fraction is of lower intensity at longer wavelengths. This provides clear evidence that DPH, even within one-component phospholipid vesicles, exhibits spectral heterogeneity associated with mobility. Conclusion This paper describes the application of fluorescence associative spectral techniques for studying membrane heterogeneity. We present here two methods for spectral association with either lifetime or rotational motions, although other associations are po~sible.’~ Our studies of PMC, which we have used as a probe of cholesterol heterogeneity, reveal a non-random distribution of probes.The associative data have been interpreted in terms of a heterogeneous model, although a transient kinetic model must also be considered. A more rigorous test of our proposed heterogeneous model may be effected by varying concentrations and rates of reaction, e.g. temperature and addition of probe and cholesterol. This will result in an altered partitioning of the probes into the different domains or phases. In addition we have considered a microheterogeneous model for the restricted rotational motions ( rm) of DPH measured in bilayer vesicles. By associating emission spectral contours with differently rotating fractions, ‘immobile’ and ‘mobile’ fractions can be assigned.Thus it appears that the rotationally homogeneous model, previously adopted to describe the anisotropic motions of DPH, is an inadequate interpretation of these data. In all we feel that the fluorescence of systems known to be heterogeneous requires modelling capable of recognizing and quantitating that heterogeneity. Associative tech- niques are well suited to this task.L. Davenport, J. R. Knutson and L. Brand 93 We thank R. E. Dale, I. Z. Steinberg, M. Ameloot and J. M. Beechem for helpful discussions. We acknowledge technical assistance from D. G. Walbridge and thank also J. M. Beechem and M. Ameloot for use of the global analysis programs. We are especially grateful to Julie Kang and Hazel Ward for their help in preparing the manuscript and to Nancy Beechem for graphic arts assistance.We are extremely grateful to Angela Fish of the Royal Society of Chemistry for her patience. This work was supported by National Institutes of Health grant no. GM11632. Contribution no. 1338 from the McCollum-Pratt Institute. References 1 Ann. NYAcad. Sci., 1978, 308. 2 M. K. Jain, in Introduction to Biological Membranes, ed. M. K. Jain and R. C. Wagner (Wiley, New York, 1980), chap. 4, pp. 53-86. 3 D. Chapman, J. Urbina and K. M. Keough, J. Biol. Chem., 1974, 249, 2512. 4 M. C. Phillips, H. Hauser and F. Paltauf, Chem. Phys. Lipids, 1972, 8, 127. 5 M. C. Phillips and E. G. Finer, Biochim. Biophys. Acta, 1974, 356, 199. 6 L. A. Sklar, B. A. Hudson and R. D.Simoni, Biochemistry, 1977, 16, 819. 7 E. J. Shimshick and H. M. McConnell, Biochern. Biophys. Res. Commun., 1973, 53, 446. 8 R. D. Klausner, A. M. Kleinfeld, R. L. Hoover and M. J. Karnovsky, J. Biol. Chem., 1980, 255, 1286. 9 R. D. Klausner and D. E. Wolf, Biochemistry, 1980, 19, 6199. 10 B. R. Lentz, Y. Barenholz and T. E. Thompson, Biochemistry, 1976, 15, 4521. 11 G. Lipari and A. Szabo, Biophys. J., 1980, 30, 489. 12 K. Kinosita Jr, S. Kawato and A. Ikegami, Biophys. J., 1977, 20, 289. 13 K. Kinosita Jr, A. Ikegami and S. Kawato, Biophys. J., 1982, 37, 461. 14 J. R. Knutson, D. G. Walbridge and L. Brand, Biochemistry, 1982, 21, 4671. 15 L. Brand, J. R. Knutson, L. Davenport, J. M. Beechem, R. E. Dale, D. G. Walbridge and A. A. Kowalczyk, in Spectroscopy and the Dynamics of Molecular Biological Systems, ed.P. Bayley and R. E. Dale (Academic Press, New York, 1985), pp. 259-305. 16 J. R. Knutson, L. Davenport, J. M. Beechem, D. G. Walbridge, M. 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Brand, Biochemistry, 1986, 25, 1811. 28 H-J. Galla and E. Sackmann, Biochim. Biophys. Acta, 1974, 339, 103. 29 J. M. Vanderkooi and J. B. Callis, Biochemistry, 1974, 13, 4000. 30 J. B. Birks, in Photophysics of Aromatic Molecules (Wiley-Interscience, New York, 1970), pp. 301-371. 31 Th. Forster and K. Kaspar, 2. Phys. Chem., 1954, 1, 275. 32 L. Davenport and L.Brand, Biophys. J., 1984, 45, 330a. 33 L. Davenport and L. Brand, Photochem. Photobiol., 1984, 39, 415. 34 L. Davenport and L. Brand, Biophys. J., 1985, 47, 367a. 35 L. Davenport, J. R. Knutson and L. Brand, Biochemistry, 1986, 25, 1186. 36 W. E. Ware and J. C. Andre, in Time-resolved Fluorescence Spectroscopy in Biochemistry and Biology, 37 L. Tilley, K. R. Thulborn and W. H. Sawyer, J, Biol. Chem., 1979, 254, 2592. 38 J. R. Knutson, L. Davenport and L. Brand, Biophys. J., 1982, 37, 203a. 39 Y. Barenholz, D. Gibbes, B. J. Litman, J. Goll, T. E. Thompson and F. D. Carlson, Biochemistry, 1977, 16, 2906. 40 L. A. Chen, R. E. Dale, S. Roth and L. Brand, J. Biol. Chem., 1977, 252, 2163. 41 R. E. Dale, L. A. Chen and L. Brand, J. Biol. Chem., 1977, 252, 7500. 42 C. W. F. McClare, Anal. Biochem., 1971, 39, 527. 43 M. G. Badea and L. Brand, in Methods in Enzymology, ed. C. H. W. Hirs and S. N. Timasheff (Academic Press, New York, 1979), vol. 61, pp. 378-394. Szabo and L. Masotti (Plenum Press, New York, 1987), in press. ed. R. B. Cundall and R. E. Dale (Plenum Press, London, 1983), pp. 363-392.94 Fluorescence Associative Techniques 44 P. Lianos and S. Georghiou, Photochem. Photobiol., 1979,30, 355. 45 A. K. Mukhopadhyay and S. Georghiou, Photochem. Photobiol., 1980, 31, 407. 46 P. Lianos, A. K. Mukhopadhyay and S. Georghiou, Photochern. Photobiol., 1980, 32, 415. 47 3. Fischkoff and J. M. Vanderkooi, J. Gen. Physiol., 1975, 65, 663. 48 R. A. Haberkorn, R. G. Griffin, M. D. Meadows and E. Oldfield, J. Am. Chem. SOC., 1977, 99, 7331. 49 B. Dekruyff, P. W. M. Van Dijck, R. A. Demel, A. Schuijff, F. Brants and L. L. M. Van Deenen, 50 E. J. Shimshick and H. M. McConnell, Biochemistry, 1973, 12, 2351. 51 L. Davenport, R. E. Dale, R. H. Bisby and R. B. Cundall, Biochemistry, 1985,24,4097. 52 D. Chapman, Q. Rev. Biophys., 1975, 8, 185. 53 T. N. Estep, D. B. Mountcastle, R. L. Biltonem and T. E. Thompson, Biochemistry, 1978, 17, 1984. 54 P. I. Lelkes, A. Kapitkovsky, H. Eibl and I. R. Miller, FEBS Lett., 1979, 103, 181. 55 B. R. Lentz, Y. Baranholz and T. E. Thompson, Biochemistry, 1976, 15,4529. 56 E. L. Chang, B. P. Gaber and J. P. Sheridan, Biophys. J., 1982, 39, 197. 57 B. P. Gaber and J. P. Sheridan, Biochim. Biophys. Acta, 1982, 685, 87. 58 C. F. Schmidt, D. Lichtenberg and T. E. Thompson, Biochemistry, 1981, 20, 4792. 59 N. 0. Peterson and S. I. Chan, Biochim. Biophys. Acta, 1978, 509, 111. 60 C. Zannoni, A. Arcioni and P. Cavatorta, Chem. Phys. Lipids, 1983, 32, 179. Biochim. Biophys. Acta, 1974, 356. Received 18th March, 1986

ISSN:0301-7249    

DOI:10.1039/DC9868100081

出版商:RSC    

年代:1986

数据来源: RSC

摘要:

Faraday Discuss. Chem. SOC., 1986,81, 95-106 A Single Spectroscopic Probe for the Determination of Both the Interfacial Solvent Properties and Electrostatic Surface Potential of Model Lipid Membranes Calum J. Drummond,* Franz Grieser and Thomas W. Healy Colloid and Surface Chemistry Group, Department of Physical Chemistry, The University of Melbourne, Parkville, Victoria 3052, Australia 2,6-Diphenyl-4-(2,4,6-triphenyl-l-pyridinio)phenoxide, &(30), has been investigated in order to ascertain its suitability as a probe for both the effective interfacial dielectric constant ( ceE) and the electrostatic surface potential (&,) of model lipid membranes in aqueous solution. This work establishes that the solvatochromic visible absorption band for b ( 3 0 ) can be used to provide a good estimate of the eeff for cationic micelles.It is also shown that the acid-base dissociation of E-,-(30) can be utilized to obtain a quantitative measure of the I/+, in the case of cationic micelles. There are problems and uncertainties associated with the use of E,,-(30) in aqueous solutions of other types of charged self-assembled surfactant aggregates, and these are discussed. The interfacial region between an aqueous solution and a self-assembled lipid phase possesses physicochemical properties which are, in general, dissimilar to both that of bulk water and the interior of the lipid self-assembled unit. The importance of the interfacial region to the function of biological membranes has long been recognized. Indeed, in order to fully understand phenomena such as the surface reactions which occur at or in biological membranes, the transport of species across membranes, and the adsorption of species onto membranes it is necessary to have an intimate knowledge of the nature of such interfacial microenvironments.Consequently, a wide range of spectroscopic probes have been employed to investigate the solvent properties (effective dielectric ~0nstantsl-l~ and m i ~ r ~ ~ i ~ ~ ~ ~ i t i e ~ ~ ~ ~ ~ ~ ~ - ~ ~ ) of the interfacial microenvironments and to determine the electrostatic surface potential^^'*^-^^ of model lipid membranes. Unfortunately, one of the spectroscopic methods currently employed which relies on the use of acid-base indicators to calculate the electrostatic surface potential generated at a charged i n t e r f a ~ e ~ - ~ ~ - ~ ~ also involves some ~ncertainty.~~ If the influence of specific molecular interactions and salt effects on the acid-base equilibrium of an interfacially located acid-base indicator can be neglected, the observed pK, (pKZbs) is dependent upon two principal factors, namely the lower effective dielectric constant ( E , ~ ) of the interfacial region and the electrostatic field at the surface.2 This dependence is shown in the following relationship: 2926-33,38339 where pKZb" is the apparent pK, value for the acid-base indicator at the charged surface, pKL is the apparent pK, value for the indicator at the interface if the surface potential (&) is zero and F, R and T are the Faraday constant, the universal gas constant and the absolute temperature, respectively.In the absence of any extra contribution from specific molecular interaction andlor salt effects to the acid-base equilibrium at the interface, the magnitude of the pKg value for an acid-base indicator is believed to be 9596 Probe for the Surface Region of Model Membranes directly related to the E , ~ value characterizing the interfacial microenvironment.2 Until now, since there are two unknowns in eqn (1) (pKa and t,bo), it has been the practice to assume, for micellar systems at least, that the pKa value can be equated with the pKZbs value determined for the acid-base indicator in non-ionic micelles of surfactants with poly( ethylene oxide) h e a d g r o ~ p s . ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ However, for a number of acid-base indicators this latter assumption is highly q ~ e s t i o n a b l e .~ ~ In addition, in the case of the acid-base indicators, such as 4-heptadecyl-7-hydroxycoumarin and 4-octadecyloxy-l- naphthoic acid, where it does appear to be an acceptable assumption to make for charged micelles, it is not clear whether it is also valid for charged vesicles.27 To avoid the uncertainty in the value of pKa and consequently Go, we have investi- gated the use of 2,6-diphenyl-4-(2,4,6-triphenyl-l -pyridinio)phenoxide, ET(30), in a dual role as a probe for both the effective interfacial dielectric constant and the electrostatic surface potential of model lipid membranes. The molecule ET( 30) possesses a phenolic oxygen atom capable of being protonated, with a pK, value in the middle region of the pH scale, and a strongly solvatochromic ( T + T") absorption band with intramolecular charge-transfer c h a r a ~ t e r .~ ~ Zachariasse et aL7 have previously utilized the solvato- chromic absorption band of ET(30) to estimate the ceff values for the interfacial microen- vironments of a number of micelles, vesicles and microemulsions. In the present work we have assumed that the E , ~ value obtained from the solvatochromic band of the ET(30) visible absorption spectra can provide the appropriate organic solvent- water mixture to best approximate or calibrate the solvent properties of the interfacial region of a charged surface. It is shown how the pK, value obtained for ET(30) in this organic solvent-water mixture can then be converted to a pKL value and used in eqn (1). We have applied this type of analysis to a number of aqueous solutions of micelles and vesicles and have demonstrated that this procedure with ET(30) can provide a quantitative measure of the electrostatic surface potential at a number of these charged interfaces.Experimental The sample of ET(30) was generously supplied by Prof. C. Reichardt, University of Marburg. The surfactants came from a variety of sources. Hexadecyltrimethylammonium bromide (CTAB), hexadecyltrimethylammonium chloride (CTAC), dodecyltrimethyl- ammonium bromide (DTAB), dodecyltrimethylammonium chloride (DTAC), dodecyl- benzene sulphonate (DBS) and hexadecylpyridinium bromide (CPB) were all purchased from Tokyo Kasei. Specially purified sodium dodecyl sulphate (SDS) and oleic acid and Brij-35 were obtained from B.D.H.Didodecyldimethylammonium bromide (DDDAB), dihexadecyldimethylammonium bromide (DHDAB) and dioctadecyl- dimethylammonium bromide (DODAB) were kindly given to us by Dr J. Brady and Prof. D. F. Evans, University of Minnesota. Prof. B. W. Ninham, Australian National University, generously provided us with dodecylethyldimethylammonium bromide (DEAB) and dodecylbutyldimethylammonium bromide (DBAB). The Nikko Chemical Co. was the source of n-dodecyloctaoxyethylene glycol monoether (C,2E8). Dihexadecylphosphate (DHP) was obtained from Sigma. L-(Y -Dimyristoylphos- phatidylcholine (DMPC) and L-(Y -dipalmitoylphosphatidylcholine (DPPC) were puriss grade from Fluka. All the solid cationic and anionic surfactants were recrystallized prior to use.The rest of the surfactants were used as received. The inorganic reagents, NaCl, NaBr, NaOH and HCl, were all analytical grade. Tetraethylammoniu'm chloride (TEAC) and tetramethylammonium chloride (TMAC) were purum grade from Fluka. These reagents were all used without further purification. The aqueous solutions were prepared with Millipore filtered water (conductivity < 1 x lop6 LR-' cm-' at 25 "C). The 174-dioxane was U.V. spectroscopic grade from Fluka and was passed through an aluminium oxide column (active neutral Brockmann grade 1C. J. Drummond, F. Grieser and T. W. Healy 97 from B.D.H.) immediately before use. This served the dual function of removing residual water and also any peroxides present in the 174-dioxane.Spectrosol-grade methanol and chloroform from Ajax Chemicals were used as received. The solid crystalline form of ET(30) was difficult to dissolve in pure water and in aqueous surfactant solution. Therefore, a small aliquot (< 1 wt YO of the total surfactant solution) of a stock solution (50: 50 wt O/O methanol : water) of E,(30) was added to the aqueous surfactant solutions so that the concentration of ET(30) was ca. 5~ lop5 mol dm-3. Unilamellar vesicle dispersions of DDDAB, DHDAB, DODAB, DMPC, DPPC and DHP were produced by following established procedure^.^.'^^^^ These procedures have been outlined in an earlier p~blication.~' The aliquot of ET(30) stock solution was always added upon the completion of the vesicle preparation. All experiments, unless otherwise stated, were performed at 25 "C.The u.v.-visible absorption spectra were measured on a Varian Cary model 210 spectrophotometer with 1 cm quartz absorption cells. The method of performing the pH titrations in surfactant solution has been given elsewhere,l2 with the exceptions that in the present study HCl (rather than H2S04) was employed with NaOH to adjust the pH, and the lower compartment of the double-junction reference electrode contained a 1 x mol dmp3 aqueous solution of TEAC. While performing the pH titrations care was always taken to ensure that the ionic strengths of the solutions were not significant altered. The pH-meter reading in a 1,4-dioxane-water mixture is not a direct measure of the hydrogen- ion activity in the solution. Hence, for these experiments the procedure of Van Uitert and Haasa was followed to enable the determination of stoichiometric hydrogen-ion concentrations from the pH-meter readings.The calibration factors obtained were in reasonable agreement with previously calculated values and will be reported in a subsequent p ~ b l i c a t i o n . ~ ~ In pure water the positively charged protonated form of ET(30) was found to be completely soluble at a concentration of 5 x lop5 mol dmp3, while the zwitterionic phenoxide form of E,(30) was found to have a solubility limit of ca. 2 x lop6 mol dmp3. Consequently, for the pH titration of ET(30) in pure water, the solution was first filtered at pH 10, pH 6 and then pH 10 again, in that order, to ensure the complete removal of any undissolved ET(30).An example of the change with pH in the u.v.-visible absorption spectrum of the solvatochromic band of ET(30) is shown in fig. 1. Not shown in fig. 1 is the isosbestic point, which occurs at a shorter wavelength (334nm in pure water). The spectra of fig. 1 are representative of the type of changes observed in the u.v.-visible absorption spectrum of ET(30) as a function of pH in pure water, 174-dioxane-water mixtures and every aquebus micellar and vesicular solution studied in this work with the exception of the SDS, DBS, DHP and oleate solutions. Fig. 2 illustrates the kind of spectra obtained for ET(30) when the pH was varied in SDS, DBS, DHP and oleate solutions. The pK, values for ET(30) in the various media investigated were determined from the spectra with the aid of eqn (2): where a is the percentage of E,(30) phenolic hydroxy groups that have been ionized.Values for a were calculated as a function of pH at the solvatochromic band maximum, A,,,, with a fixed concentration of ET(30). The maximum absorbance obtainable for ET(30) at A,,,, by varying the pH of the solution, was taken as the 100% ionization value and the minimum absorbance as the 0% ionization value. At least six different a values between zero and 100 were examined for each 174-dioxane-water mixture or self-assembled surfactant solution. This protracted procedure established that the PK:~" values for ET( 30) residing in the interfacial microenvironments of the lipid self-assembled98 Fig. 1. U.v.- Probe for the Surface Region of Model Membranes 0.L 0.3 8 2 2 0.2 3 rd 0.1 0 LOO 5 00 600 700 wavelength/nm -visible absorption spectrum of 7.2 x mol dmP3 E,,-(30) in a 50 wt % water mixture ( E = 35.85) as a function of pH at 25 "C. The structure of b ( 3 0 ) is also shown. wavelength/nm Fig. 2. U.v.-visible absorption spectrum of 5.0 x mol dmP3 b ( 3 0 ) in 0.069 mol dm-3 SDS solution as a function of pH at 25 "C. an aqueous phases were well defined. All the pK, values quoted in the present work are the average values calculated by using the results of the pH titrations and eqn (2). The magnitude of the error associated with each pK, value indicates the maximum deviation from the average value. Results Fig. 3 and 4 illustrate how the A, value for the solvatochromic band of ET(30) varies with the dielectric constant of a number of organic solvent-water mixtures and neat n-alcohols.The A,,, values in the isopropanol-water, ethanol-water, methanol-water,C. J. Drummond, F. Grieser and T. W. Healy 60 01 I I I I I I I 1 t t 1 dielectric constant 99 Fig. 3. A,,, values for the solvatochromic band of E-,-(30) in 0, isopropanol-water: M, ethanol- water; 0, methanol-water and 0, ethylene glycol-water mixtures as a function of the dielectric constants of the mixtures at 25 "C. dielectric constant Fig. 4. A, values for the solvatochromic band of b ( 3 0 ) in 0 , acetone-water mixtures; 9, 1,4-dioxane-water mixtures and 0 , neat n-alcohols as a function of the dielectric constants of the media at 25 "C.100 Probe for the Surface Region of Model Membranes dielectric constant Fig.5. ApKF (0) and ApKa (0) values for G(30) in 1,4-dioxane-water mixtures as a function of the dielectric constants of the mixtures at 25 "C. ethylene glycol-water, acetone-water and 1,4-dioxane-water mixtures were acquired from the works of Dimroth and Reichardt& and Kosower et al.47 The dielectric constants of the mixtures were interpolated from the data of Akerlof4* and Critchfield et al.49 The A,,, values in the neat n-alcohols were obtained from the compilation of Reichardt and Harbusch-GonertS0 and the dielectric constants for these neat solvents were taken from ref. (51) and (52). Also included in fig. 4 are our own results for 1,4-dioxane-water mixtures. We obtained a A,,, value of 454nm for ET(30) in pure water, which is in close agreement with the value of 453 nm found by Dimroth and R e i ~ h a r d t .~ ~ The changes in the pK, value for ET(30) in 1,4-dioxane-water mixtures relative to its pK, value in pure water (i.e. ApKr) are shown in fig. 5 as a function of the dielectric constant of the mixtures. The ApKF data points shown in fig. 5 refer to pure water and 10, 20, 30, 40, 50, 60, 70 and 80 wt '/o 1,4-dioxane-water mixtures. The values of Critchfield et al.49 for the dielectric constants of 1,4:dioxane-water mixtures were used for fig. 5. The pK, value found for ET(30) in pure water was 8.63k0.03. The explanation of how, in the absence of distortion of the acid-base equilibria by any specific molecular interactions and salt effects, the p K a values can be derived from the pKF values has already been clearly and comprehensively covered by Fernandez and Fromherz,2 and only a very brief account will be given here.The basic premise is that for a 1,4-dioxane-water mixture which has a dielectric constant identical to the E~~ characterizing a charged interface the difference between the pK," and pKH value is solely due to the medium effect on the activity coefficient of the proton, i.e. pK: = pKF - log my"+. The medium effect for the proton, cannot be measured. Hence we have followed the procedure of Fernandez and Fromherz2 and have used the values of the medium effect for HCl in 1,4-dioxane-water mixtures, ,y+, to approximate the ,yH+ values. In the present work, the log my* values were obtained from the change in the standard potential, E", for the cell Pt I H,(g), HCl in 1,4-dioxane-water mixture, AgCl I Ag usingC.J. Drummond, F. Grieser and T. W. Healy 101 Table 1. The A,,,, E , ~ , pK:, pK",' and (clo values obtained with b(30) in the aqueous self- assembled surfactant solutions investigated at 25 "C. Also included are the (clo values calculated using 4-heptadecyl-7-hydroxycoumarin, (clo( HHC) (see text for explanation) concentration A,,, $0 +o(HHC) surfactant /mmol dm-3 /nm Eeff PK', p K zbs /mV /mV C12E8 C12E8 CTAC CTAB DTAC DTAC" DTAB DTAB DTAB DTAB DTAB' DEAB DBAB CPBf SDSg SDSg SDSg SDSg DBSg Oleateg DHPg DMPC DPPC 10 204 50 50 50 50 50 65 162' 324d 50 50 50 50 50 69 173' 347d 50 50 2.5 5 5 540 542 532 534 524 537 528 528 530 532 542 530 532 533 493 493 493 493 490 505 503 545 517 30 29 34 33 39 31 36 36 35 34 29 35 34 33 56 56 56 56 58 49 51 27 43 9.61 9.61 9.60 9.60 9.57 9.61 9.59 9.59 9.59 9:60 9.61 9.59 9.60 9.60 9.32 9.32 9.32 9.32 9.27 9.45 9.41 9.61 9.53 9.06 f 0.04 9.31 f 0.21 6.93 f 0.06 7.22 f 0.01 7.39 f 0.03 8.82 f 0.03 7.59 f 0.04 7.67 f 0.02 7.93 f 0.02 8.07 f 0.04 9.30 f 0.06 7.79 f 0.03 7.78 f 0.03 7.08 f 0.06 10.72 f 0.03 10.70 f 0.03 10.68 f 0.06 10.48 f 0.06 10.29 f 0.03 10.91 f 0.05 10.96 f 0.08 10.55 f 0.02 10.48 f 0.10 0 0 +158 + 141 +129 +47 +118 +114 +98 +91 +18 +lo6 + 108 + 149 -83h -82h -80h -69h -60h -86h -92h -56h -56h 0 +154 +139 +127 +37 +116 +114 + 102 +85 +19 - - - - - - - - - - - -115h - 173h 4 mol dm-3 NaCl.2 wt '/o surfactant. ' 5 wt '/o surfactant. Experiment performed at 30 "C. 10 wt '/o surfactant. 4 mol dm-3 Spectra as a function of pH differed from that obtained (clo value is not the electrostatic NaBr.in pure water and 1,4-dioxane-water mixtures (see text for details). surface potential (see text for details). the r e l a t i ~ n s h i p ~ ~ where is the standard potential for the cell when pure water is the solvent and is equal to 0.222 34 V at 25 0C54 and ' E o is the standard potential of the cell in a 1,4-dioxane- water mixture. The ' E 0 values were taken from the compilation of Feakins and French,55 with the exception of the value in the 82 wt% percent 1,4-dioxane-water mixture which was taken from the work of Danyluk et aL56 Fig. 5 illustrates the change in the pKL value for ET(30) relative to its pK, value in pure water (i.e. ApK!J as a function of the dielectric constant of the 1,4-dioxane-water mixtures.Presented in table 1 are the A,,, and pKib" values determined for ET(30) in the self-assembled surfactant solutions investigated. The corresponding E , ~ values based on the 1,4-dioxane-water curve of fig. 4 are also given. E , ~ values based on other reference systems can also be easily determined with the aid of fig. 3 and 4. These E , ~ values are in reasonable agreement with those that have been determined by Zachariasse et al.,' with the exception of the DMPC and DPPC results. For both DMPC at 25 "C and DPPC at 50 "C they obtained an eeff value, based on the 174-dioxane-water reference system, of 14. We were unable to ascertain the reason for this discrepancy between the102 Probe for the Surface Region of Model Membranes two studies.The pKa values contained in table 1 were obtained from fig. 5 by using the eeff values for the particular self-assembled surfactant systems. These p K a values and the PK;~” values were then substituted into eqn (1) to gain the qb0 values. Not included in table 1 are the results for E,(30) in the unilamellar vesicle dispersions of DDDAB, DHDAB and DODAB, since at the surfactant concentrations employed in this study, (2.5 x lop3 and 5.0 x mol dm-3), both acidic and basic forms of ET(30) were found to partition only slightly, if at all, into the interfacial region of the vesicles, i.e. the same A, and pKZb” results were obtained for ET(30) in these vesicular solutions as were obtained in pure water. In micellar Brij-35 solutions it was not possible to reproduce the spectrum of ET(30) as a function of pH, as the absorption maximum of the phenoxide form of ET(30) disappeared rapidly with time.Once the maximum had vanished it was impossible to regain with the addition of NaOH. Similar behaviour was also observed when unpurified 1,4-dioxane was used as a solvent component and in micellar CI2E8 solution, but to a lesser extent and over a much longer time span in this case. We believe this behaviour is due to a reaction occurring between the phenoxide form of ET(30) and possibly peroxides, or other oxidants present in the sample of Brij-35. Similar behaviour for ET( 30) in unpurified cyclohexanol, 2,4-dimethylpentan- 1 -one, 1 -phenylethanol and 3- phenylpropan-1-01 has also been observed by Aslam et aL5’ Discussion The fundamental assumptions inherent in the type of treatment proposed in this study to determine +b0 values from the pKzd” values for ET(30) in aqueous solutions of charged self-assembled surfactant aggregates are that: (i) the ionizable phenolic hydroxy group of ET(30) resides, on average, in the plane of the charged headgroups of the self- assembled surfactant aggregates; (ii) the position of the solvatochromic band maximum for ET(30) in the self-assembled surfactant aggregates gives a true indication of the eeff of the interfacial region; (iii) the interfacial solvent properties characterized by the eeff value have the same influence on the pKa value for ET(30) as does a 1,4-dioxane-water mixture of equivalent dielectric constant; (iv) the differences between the pK, in pure water and the pKZbS values are solely due to the surface potential and the lower E , ~ at the charged interface; i.e.both acidic and basic forms of ET(30) have fully partitioned into the interfacial region and specific molecular interactions and/or interfacial salt effects do not influence the PK:~” values. The forthcoming discussion will primarily consist of an assessment of the validity of each of these assumptions when dealing with the different kinds of self-assembled surfactant aggregates. Cationic Micelles Two very different types of n.m.r. have established that the average location for the phenolic oxygen atom of ET(30) in cationic micelles is in the plane of the positively charged surfactant headgroups. Thus assumption (i) appears to be justified for cationic micelles.A large electrochromic component60961 to the A,,, values in self-assembled surfactant solutions would invalidate assumption (ii). However, the A, results contained in table 1, especially those for the CI2E8 micelles and DTAB and DTAC micelles with up to 4moldm-3 electrolyte, suggest that the shifts in A,,, are not the result of a large electrochromic response. Since there is a large effective counter-ion concentration present within the interfacial region of m i ~ e l l e s ~ ~ . ~ ~ assumption (ii) would be unjustified if there was a major com- ponent in the ET(30) A,,, values which was due to electrolyte interaction. We tried toC. J. Drummond, F. Grieser and T. W. Healy 103 gauge if there was any possibility of a large electrolyte induced A,,, shift by attempting to determine the A,,, of ET(30) in a number of aqueous electrolyte solutions.We found, however, that the phenoxide form of ET(30) precipitated on the addition of electrolyte, 10 mmol dm-3 NaCl and TMAC, and consequently we were unable to obtain any Amax values in this type of media. Nevertheless, the results for l-methyl-4-[(oxo- cyclohexadienylidene)ethylidene]- 1,4-dih~dropyridine,~~ a more water soluble molecule that possesses solvatochromic behaviour similar to that of ET(30), suggest that it is unlikely that there would be a large electrolyte effect on the A,,, values for ET(30). Furthermore, the E , ~ values determined for the interfacial microenvironments of the cationic micelles are also similar to estimates that have been obtained by employing other solvatochromic spectroscopic One way to test assumptions (iii) and (iv) would be to locate ET(30) in an interfacial microenvironment where the surface potential is zero or close to zero.In principle, this can be achieved by using either non-ionic micelles or by using a high electrolyte concentration to ‘screen’ the surface charge density of a charged interface. Unfortunately, owing to the problems associated with the use of ET(30) in non-ionic micelles comprised of surfactants with poly( ethylene oxide) headgroups, which have been mentioned in the Results section, it was not possible to obtain an accurate pKzbs value for ET(30) in CIZEs micelles. The pK:bs results for C&, given in table 1, also indicate that it is necessary to have a very high concentqation of C,,E, micelles in order to ensure that most of the protonated form of ET(30) has partitioned into the micellar phase.In addition, from the results for DTAC micelles in the presence of 4 mol dm-3 NaCl and DTAB micelles with 4 mol dmP3 NaBr it is evident that the surface charge density of these micelles is still not fully ‘screened’ even at this high electrolyte concentration. A range of added electrolyte concentrations, between 0 and 4 mol dmP3, were investigated for DTAC and DTAB micelles. There is a monotonic decrease in the t,bo as electrolyte is added for both DTAC and DTAB m i ~ e l l e s . ~ ~ A 6 mol dm-3 NaBr solution of DTAB micelles was also investigated but was found to be extremely viscous, and it was not possible to perform reliable pH titrations with ET(30) in this medium.Although pKgb” values for ET(30) in C& micelles and DTAB/4 mol dmP3 NaBr micelles do not prove that the magnitude of the pKa values are quantitatively correct they certainly suggest that the pKa values are at least within 0.3 pK, units of being correct. with 4-heptadecyl-7-hydroxycoumarin (HHC) t,bo values for man of the self-assembled aggregates of table 1 were calculated by assuming that the pK$ value for HHC in C&8 micelles could serve as the pKg value in eqn (1). Since it is now clearly evident that ceff is not the same for each of the different cationic micelles, we have used the ceff values obtained with ET(30) and the curve of pKg as a function of the dielectric constant of 1,4-dioxane-water mixtures for the 7-hydroxycoumarin chromophore, which was determined by Fernandez and Fromherz,* to correct our earlier estimates of the t,bo values.These corrected values are given as the t,bo (HHC) values in table 1. For cationic micelles the agreement between the t,bo values calculated with ET(30), and the t,bo values calculated with HHC is extremely good. Both the un-ionized and ionized forms of HHC fully partition into the micellar phase and both forms of the 7-hydroxycoumarin chromophore reside within the interfacial region of cationic m i ~ e l l e s . ~ ~ It has been shown27 that there is probably little, if any, influence from specific molecular interaction and interfacial salt effects on the acid-base equilibria of HHC at charged interfaces.For 1,4-dioxane-water mixtures, the pKg behaviour of the HHC with changing dielectric constant is not the same as the ET(30) pKL behaviour with changing dielectric constant. Therefore, the close agreement between the t,bo value calculated with ET(30) and the t,bo value calculated with HHC for a particular cationic micelle is a clear indication that assumptions (ii), (iii) and (iv) must be valid in the case of cationic micelles. In an earlier104 Probe for the Surface Region of Model Membranes Anionic Micelles The n.m.r. study of Plieninger and B a ~ m g a r t e l ’ ~ ~ ~ ~ has shown that in the case of SDS micelles the N+ centre of ET(30) is aligned on average in the plane of the sulphate headgroups and the ionizable hydroxy group is positioned some distance out from the plane of the anionic headgroups. This should also be the case for E,(30) in the anionic DBS and oleate micelles. Consequently the acid-base dissociation of ET( 30) in these anionic micelles will not be influenced by the electrostatic surface potential but by the electrostatic potential at the average position of ‘sit’ for the hydroxy group.As can be seen by comparing fig. 1 and 2, the behaviour of the visible absorption spectrum of ET(30) in anionic micelles as a function of pH is unlike that seen in pure water, 1,4-dioxane-water mixtures or cationic micelles. This is probably a result of specific molecular interaction between the N’ centre of the ET(30) molecule and an anionic headgroup which affects the optical properties of the ET(30) molecule.Because of this difference in the optical properties of ET(30) in anionic micelles we are uncertain whether or not the E,(30) A,,, values in these micelles can be compared with the A,,, results of fig. 3 and 4 to obtain eeff values for the interfacial microenvironments. Table 1 contains E , ~ values based on the assumption that this type of comparison can be made. Note that the E , ~ values are consistent with the known location of the probe, i.e. higher ceff values would be expected for ET(30) in anionic micelles than for ET(30) in cationic micelles because ET(30) is on average located further out from the interface in the former case. Interestingly, Plienir~ger~~ observed that the addition of a long alkoxy chain to the ET(30) molecule, at the opposite end of the molecule to the phenolic oxygen atom, altered the A,,, values obtained with this probe in SDS micelles in a manner that was consistent with the long alkoxy chain ‘dragging’ the chromophore in closer to the micelle interior.For CTAB micelles P l i e ~ ~ i n g e r ~ ~ found very little difference between the results obtained with ET(30) and the alkoxy ET(30). The magnitude of the t,bo values, which were calculated by assuming that the eeff values given in table 1 are correct, are also consistent with the ET( 30) molecule ‘sensing’ an electrostatic potential out from the plane of the headgroups in the case of anionic micelles. It should be emphasized, however, that the different optical properties found for ET(30) in anionic micelles introduces a large element of uncertainty into the analysis of.these systems. Vesicles No studies on the average location of ET(30) in vesicles have been reported. As discussed in the Results section, at the concentration of surfactant employed in this investigation, ET(30) does not appear to partition into the cationic DDDAB, DHDAB and DODAB vesicles to any great extent. The spectrum of ET(30) as a function of pH in the anionic DHP vesicles is similar to that seen for ET(30) in the anionic micelles. Therefore, it is highly likely that the N+ centre of ET(30) is aligned on average in the plane of the phosphate headgroups with the ionizable hydroxy group situated on average at a position out from the plane of the headgroups. Because of the optical properties of ET(30) in DHP vesicles being different to those of ET(30) in 174-dioxane-water mixtures, the same uncertainty is involved in the calculated eeff and t,bo values, table 1, as has been discussed for the case of the anionic micelles.The optical properties of E,(30) in DMPC and DPPC vesicles suggest’that there is, on average, little or no specific molecular interaction between the Nt centre of ET(30) and the negative phosphate part of the zwitterionic phosphatidylcholine headgroups. Although it is probably situated, on average, somewhere in the vicinity of the glycerol backbone region of the vesicles, the exact location of ET(30) in DMPC and DPPC vesicles is unknown. Consequently, at this stage it is not possible to assess whether the different eeff values found for the DMPC and DPPC vesicles at 25 “C are due to these types of vesicles possessing different interfacial properties above their gel-liquid-crystal-C.J. Drummond, F. Grieser and T. W. Healy 105 line phase transition temperature, T,, to what they do below their T, (DMPC, T, = 24 "C and DPPC, T, = 41 "C) or to the ET(30) molecule being located at a different position in the two vesicles. The $o results for the DPPC and DMPC vesicles indicate that the ET( 30) molecule 'senses' a local negative electrostatic potential, which is consistent with ET(30) residing on average in the glycerol backbone region of these vesicles. Work with HHC has indicated2' that a local negative electrostatic potential exists in this region. The differences between the & (HHC) values and the E,(30) values (table 1) can be attributed to the two chromophores having different average sites of residence in the glycerol backbone region.The finding that local electrostatic potentials exist in the interfacial microenvironments of vesicles with zwitterionic phosphatidylcholine head- groups, and that they influence the acid-base dissociation of molecules, is interesting in view of the fact that electrophoretic mobility measurements66 indicate that the net electrostatic surface potential for these vesicles is zero for the pH range studied in this paper. Conclusions It has been established that ET (30) can be employed to determine both the effective interfacial dielectric constant and the electrostatic surface potential of a cationic micelle. The optical properties of ET(30) in anionic micelles and vesicles have been found to be dissimilar to those of ET( 30) in pure water, 174-dioxane-water mixtures, cationic micelles and vesicles with zwitterionic phosphatidylcholine headgroups.As a consequence of this we are uncertain whether or not the A,,, values obtained for ET(30) in anionic micelles and vesicles can be compared with the reference 174-dioxane-water mixtures to determine tzeff values. However, if it is assumed that the comparison can be made, the ceff and values calculated are consistent with the known average location and orientation of the ET( 30) molecule. At 25 "C the ceff values determined for the DMPC and DPPC vesicles are not the same. However, the average location of ET(30) in these vesicles is unknown.Therefore, it is not clear whether this difference is a result of the ET(30) molecule being situated at a different average site of residence in the two types of vesicles or due to the ceff value at the one site being different. The I,!J~ results for the DMPC and DPPC vesicles clearly indicate the presence of a local negative electrostatic potential within the inter- facial microenvironment of these zwitterionic vesicles. We are currently investigating other kinds of solvatochromic acid-base indicators in the hope that some of the problems that have been associated with the use of ET(30) in anionic micelles and in all types of vesicles may be resolved. We thank Prof. C. Reichardt, University of Marburg, for his generous gift of the E,(30). We thank Dr J Brady and Prof.D. F. Evans, University of Minnesota, and Prof. B. W. Ninham, Australian National University, for supplying some of the surfactants. 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Chem., 1976, 80, 207. 62 P. Mukerjee, J. Phys. Chem., 1962, 66, 943. 63 F. M. Menger, H. Yoshinga, K. S. Venkatasubban and A. R. Das, J. Org. Chem., 1981,46, 415. 64 S. J. Davidson and W. P. Jencks, J. Am. Chem. SOC., 1969, 91, 225. 65 A. Haase, Ph.D. Dissertation (University of Giessen, 1980). 66 D. Papahadjopoulus, Biochim. Biophys. Acta, 1968, 163, 240. 1982, 104, 1800. Sci., 1984, 97, 380. 2167. Received 12th December, 1985

ISSN:0301-7249    

DOI:10.1039/DC9868100095

出版商:RSC    

年代:1986

数据来源: RSC

摘要:

Faraday Discuss. Chem. SOC., 1986, 81, 107-116 Physicochemical Studies of Vesicles and Biomembranes Spectroscopic Studies and Phospholipid Polymers Dennis Chapman,* David C. Lee and James A. Hayward Department of Biochemistry and Chemistry, Royal Free Hospital School of Medicine, Rowland Hill Street, London NW3 2PF Recent biophysical studies using a range of physical and spectroscopic techniques of both biomembranes and various lipid-water (vesicle) systems are described. The nature of lipid-protein interactions in some natural and model biomembranes has been examined with regard to the extent of lipid perturbation and protein conformational changes. The application of second- derivative and fourth-derivative infrared spectroscopy to these systems indi- cates clearly the bands associated with the secondary structure and also the weak bands associated with the minor amide components of the membrane proteins.The technique also provides evidence for the effects of intrinsic proteins on the lipid carbonyl groups when the lipid is below its T, transition temperature. Studies of phospholipid polymers in vesicle form and in Langmuir- Blodgett films are discussed. The introduction of new biomembrane-mimetic surfaces are also described. The latter are used to study protein and cell adsorption to particular phospholipid polar groups. In recent years we have been examining lipid vesicles and biomembranes using a variety of physical techniques. We have been particularly interested in examining lipid-protein interactions, i.e. how the lipid molecules are perturbed by the presence of intrinsic or integral proteins and also seeking methods to provide information about membrane protein structures.We have also set out to create new polymers, i.e. phospholipid polymers, so as to produce polymeric vesicles, thereby increasing their stability for certain applications. We have also developed derivatisation methods for treating the surfaces of glass and metals producing in some cases surfaces which mimic (in a simple form) the outer-surface character of the lipid portion of red blood cells. We describe some of our recent studies in this paper. I.R. Studies of Protein-Lipid Interactions and Protein Structure in Natural and Reconstituted Membranes Fourier transform infrared (f.t.i.r.) spectroscopy has been applied extensively in recent years to probe the structure and dynamics of biological membranes. Variations in band frequency, linewidth and intensity are sensitive to structural transitions of both lipid and protein components.The vibrations of individual groups provide structural informa- tion on highly localised regions of the lipid bilayer. Thus, the C-H stretching absorp- tions of the lipid acyl chains are readily distinguished from the carbonyl stretchings of the interfacial region and the phosphate stretchings of the polar headgroup. Of particular importance in the study of lipid-protein interactions is the non-perturbing nature of the technique. The addition of an external probe molecule is not required and the absorp- tions of the lipid and protein groupings reflect their genuine environments.The principal impediment to the study of aqueous membranes and their models, i.e. the strong and 107108 Spect roscopic Studies and Phospholipid Polymers broad i.r. absorptions of water, has been removed with the use of microcomputers for digital subtraction of the background. 192 Studies of the i.r. spectra of phospholipids have provided much information on the structure of the acyl chain, interfacial and headgroup regions of aqueous bilayer~.~ The main endothermic phase transition of aqueous phospholipid bilayers results in pro- nounced alterations in the methylene band parameters. The band maximum frequencies of the CH, asymmetric and symmetric stretching bands are sensitive to the static order of the acyl chains.The introduction of an increased proportion of gauche conformers above the phase transition causes a shift of these bands to higher freq~encies.~ The width of these i.r. absorptions is determined by rotational, translational or collisional effects. Thus, the CH, bandwidths are sensitive to the degree of motional freedom of the CH, groups. Most i.r. studies of protein-lipid interactions have concentrated on measurement of the C-H bands. In an early study the effects of the incorporation of the intrinsic proteins Ca*+-ATPase and bacteriorhodopsin and the intrinsic polypeptide gramicidin A on the acyl chain order of saturated phosphatidylcholine bilayers were r e p ~ r t e d . ~ It was found that below the phase-transition temperature, these molecules behaved in a similar manner to cholesterol, causing a disordering of the chains.Above the phase transition, the perturbations of the acyl chains differed from the progressive ordering effect observed with cholesterol. At high lipid protein molar ratios, an ordering of the acyl chains was observed. However, when the concentration of the intrinsic protein was increased this effect was removed and the static order of the chains was essentially the same as in the pure lipid system. More recently, we have repeated this type of study using dimyris- toylphosphatidylcholine in which the acyl chain hydrogen atoms have been completely replaced by deuterium atoms.6 This allows examination of the acyl chain C-'H stretching modes without overlap from the C-H stretching modes of the intrinsic protein.Our study indicated that bacteriorhodopsin, at low concentration, has no effect on the static order of the lipid racy1 chains above the phase-transition temperature (fig. 1). At high protein concentration there was some disordering of the lipid acyl chains, indicated by an increase in band frequency compared to the pure lipid. 1.r. spectroscopy may be used to probe the interfacial region of lipid bilayers via examination of the C=O stretching bands between 1750 and 1700 cm-'. Two absorption bands in this region are associated with the two ester groupings in diacyl lipids. The sn-1 carbonyl absorbs near 1740 cm-' while a band near 1725 cm-' is due to the carbonyl at the sn-2 position3 (see fig. 2). After the subtraction of the aqueous background the i.r.spectrum normally reveals only a single broad C=O band envelope which shifts by 4 cm-' to lower frequencies at the phase transition. This shift arises from alterations in intensity rather then frequency of the sn-1 and sn-2 components, as revealed by second- derivative (fig. 2) or deconvolution calculations. On passing through the phase transition, we observed a decrease in the intensity of the 1740cm-' component relative to the 1726-1728 cm-' component. Values for the intensity ratio of the two components of the carbonyl stretching band are given in table 1. Intensity ratios were calculated by measuring the negative absorbance from zero at 1726 and 1740cm-' in the second- derivative spectra. For DMPC, conversion from the gel to liquid-crystalline state produced a decrease in the relative intensity at 1740 cm-'. Incorporation of bacterio- rhodopsin or Ca2+-ATPase into the bilayers caused a large reduction in 11726/11740 below T, (23 "C).Above T,, a slight increase in was observed for the bacteriorhodop- sin-containing liposomes and a slight decrease for those containing Ca2+-ATPase. However, these differences were small compared to those observed below T'. We conclude that the presence of the integral proteins in the bilayer reduces the conforma- tional inequivalence below the phase transition so that a number of the sn-2 chains are now constrained in a conformation similar to that of the sn-1 chain.D. Chapman, D. C. Lee and J. A. Hayward 109 2095 2094 2093 7 2092 2- 2091 E 2090 2089 2088 2087 Q J. I I I I I I I I 10 15 20 25 30 35 40 45 temperature/"C Fig.1. Variation of C2H2 symmetric stretching frequency with temperature for 0, dmpc A, dmpc [2H]54/bacteriorhodopsin (102 : 1) and V, dmpc-[2H],2/bacteriorhodopsin (22: 1) molar ratios. 1.r. spectroscopy is an established technique for the study of the structures of polypeptides and proteins. The i.r.-active amide bands are associated with the CONH grouping which these molecules have in common. Initial qualitative studies related the frequencies of the relatively strong amide I and amide 11 bands to the presence of specific types of secondary structure in various soluble polypeptides and proteins. In principle, a globular protein containing several types of substructure will give several amide I maxima. However, the large half-widths of these components prevents their resolution.A method for assessing the number and position of component peaks is derivative spectroscopy. Second-derivative i.r. spectra of water-soluble proteins have been obtained7 and peaks associated with a-helical, @-sheet and @-turn conformations together with the vibrations of some amino acid side-chains were resolved. We have recently obtained the first second-derivative i.r. spectra of membrane protein^.'^' First we concentrate on our studies of the sarcoplasmic reticulum (SR), a membrane system which is involved in the regulation of the contraction-relaxation cycle of striated muscle. The majority of the protein present in the membrane is the Ca2+ activated ATPase which causes the active accumulation of Ca2+ into the SR during relaxation of the muscle.We have studied the secondary structure of this protein in three environments: the isolated SR, vesicles of purified Ca2'-ATPase in SR lipids and reconstituted into bilayers of dimyristoylphosphatidylcholine.' In fig. 2 we present difference, second- and fourth-derivative f.t.i.r. spectra of SR membrane after the subtraction of the aqueous buffer background. Negative bands in the second-derivative correspond to positive bands in the fourth-derivative, which, in turn, correspond to positive absorption bands in the difference spectrum. Three main absorption bands are110 Spectroscopic Studies and Phospholipid Polymers 1 1 1 1 1 1 i 1 1 1 1 1 1 1 1 1 1 1 1 t I 1 I I I I I I l - l l I I I I I I I 1 19 00 1800 1700 1600 1500 1 1 wavenumber/ cm- ' I 30 Fig.2. (a) Difference, (b) second-derivative and ( c ) fourth-derivative f.t.i.r. spectra of sarcoplasmic reticulum at pH 7.4 and 20 "C after subtraction of the aqueous background absorption. seen in the difference spectrum, the amide I and amide I1 bands from the protein at 1655 and 1547 cm-', respectively, and a C=O stretching band from the lipid at 1737 cm-'. The frequency of the amide I maximum of 1655 cm-' may be assigned to the presence of a predominantly a-helical protein in the membrane. However, analysis of the second- derivative reveals the presence of p-sheet structure, with absorptions at 1632 cm-' and 1680-1690 cm-', which has been predicted from the primary sequence." Further analysis using the fourth-derivative reveals components in the amide I region which may be assigned to a-helical structure (1657 and 1643 cm-') P-sheet structure (1681 and 1630 cm-') and p-turns (1690 ~ m - ' ) .~ , ~ The band at 1531 cm-' in the amide I1 region may also be due to P-structure.' The fourth-derivative spectrum also reveals bands which may be assigned to amino acid side-chains," which were previously lost beneath the broad amide I and I1 band envelopes. These are tyrosine (1516 and 1613 cm-'), glutamate (1568 cm-') and arginine or aspartate (1581 cm-'). These bands may provide future probes for structural alter- ations in enzyme active sites in a wide range of systems. The bands at 1742 and 1727cm-' in the second-derivative are C=O stretching absorptions, assigned above, from the lipid present in the membrane.The fourth- derivative reveals a third component at 1709 cm-' which has also been reported by other workers using spectral deconv~lution.~ Bands in the 1468-1421 cm-' region are CH2D. Chapman, D. C. Lee and J. A. Hayward Table 1. The effects of Ca2+-ATPase and bacteriorhodopsin on the intensity ratio of the carbonyl stretching bands of DMPC" 111 DMPC/Ca2--ATPase DMPC/bacteriorhodopsin T/"C DMPC 245 : 1 135: 1 10 0.52 0.3 1 20 0.45 0.43 25 0.50 0.55 35 0.79 0.70 45 0.80 0.8 1 0.32 - - 0.83 - Data were obtained from the second-derivative spectra presented, in part, in ref. (8). The negative absorbance intensities at 1726 and 1740 cm-' were measured at each temperature. deformation modes from the lipid acyl chains. The region 1900-1800 cm-' is free from absorption bands and gives an indication of the low noise in the spectra.Fig. 3 presents difference, second-derivative and fourth-derivative spectra of purple membrane, a light-energy transducing membrane which forms part of the plasma membrane of Halobacterium halobium and other extreme halophiles. These patches contain a single protein, bacteriorhodopsin, which uses light energy to translocate protons across the membrane, thereby setting up an electrochemical gradient. This gradient is used to synthesize ATP and subsequently, to provide energy for the metabolism of the cell. The amide band maximum at 1660cm-' may be assigned to the presence of a,,-helices in the protein. A predominantly a-helical structure is also indicated by the amide I1 band frequency of 1545 cm-', in agreement with the electron diffraction study of Henderson and Unwin.12 However, second- and fourth-derivative analysis of the broad amide bands reveals components at 1684, 1635 and 1530cm-', which may be assigned to @-structure.' This @-structure has also been revealed by circular dichroism and was proposed to be transmembrane antiparallel P-sheet.13 Further studies are in progress in our laboratory using selective modification of bacteriorhodopsin by enzyme cleavage to determine the location of the structures giving rise to these i.r. absorptions.Other absorptions in the fourth-derivative f.t.i.r. spectrum of purple membrane may be assigned to amino acid side-chains in bacteriorhodopsin' as for the Ca2+-ATPase. Phospholipid Polymers In order to probe the molecular structure and motional dynamics of biological mem- branes and their models, l4 photosensitive moieties have been incorporated into lipid structures.The presence of a small photolabile group permits the generation of reactive species with minimal perturbation of the structure of the membrane in which they reside. In contrast to their use as photoaffinity labels, our approach toward photosensitive phospholipids has been for the preparation of stable polymers. We have used polymeric phospholipids as models of biological membranes and in biomedical applications. A wide variety of polymerizable functional moieties has been incorporated into an assortment of amphipathic compounds to produce stable surfactant assemblies.' Many of these compounds have contained a photolabile diacetylene group and were based upon the initial observations of Wegner, who showed that these triple-bonded structures formed crystalline polymers upon irradiation with U.V.light. l6 We have synthesised phospholipids that contain diacetylene groups in one or both of their acyl chains [fig. 4( a)]."," The physical properties of the monomeric, diacetylenic112 Spectroscopic Studies and Phospholipid Polymers 1900 1800 1700 1600 1500 wavenumberj cm- ' 10 Fig. 3. (a) Difference, (b) second-derivative and ( c ) fourth-derivative f.t.i.r. spectra of purple membrane at pH 7.4 and 20 "C after subtraction of the aqueous backgound absorption. phospholipid were studied by optical and magnetic resonance spectro~copy~~-~* and by calorimetric and monolayer techniques22923 and were found to resemble the properties of naturally occurring lipids.N.m.r. relaxation times both for the head groups and for the resolved resonances in the acyl chains were very similar in diacetylenic and non-poly- merizable phospholipids. Calorimetric studies showed that the temperature of the thermotropic phase transition, T,, is lower for the diacetylenic phospholipids than for the corresponding saturated phosphatidylcholines, but is higher than that of the cis- unsaturated homologue~.~~ In mixed-chain phosphatidylcholines, replacement of the long diacetylene-containing acyl chain with a saturated acyl chain disrupts the packing of the lipid and further decreases T,. This disruption in the packing of the acyl chains is also evident in the inability of mixed-chain diacetylenic phosphatidylcholines to form a stable monolayer at the air-water interface, even at low temperatures." These phospholipid molecules containing diacetylene groups form polymers upon irradiation with U.V.light. The incorporation of polymeric phospholipids within the bilayers of liposomes increases their resistance to precipitation and leakage, and permits the preparation of stable surfaces that mimic the surfaces of biomembranes. Phospholipid Polymerisation in Whole Cells The physicochemical similarities between diacetylenic phospholipid monomers and conventional lipids suggested that phospholipid polymers might be found in situ follow-D, Chapman, D. C. Lee and J. A. Hayward $=o $=O F=O y=o $=O :=o- ./’ where PC is ‘.CHz-C/H -$H2 r--9: - - - -1 I o=p-0 I 113 A/nm Fig. 4. (a) Formation of the polyconjugated phospholipid polymer from the diacetylenic monomer; n may be varied to produce monomers of different lengths and different phase-transition tem- peratures. The hydrocarbon chains may be esterified to different polar head groups. ( 6 ) Visible spectra of 86 layers of diacetylenic phosphatidylcholine after various irradiation times. The sample consists of 43 layers of lipid on each side of a quartz slide. The upper surface is hydrophilic and formed by the polar head groups. ing incorporation of the monomeric lipid into the membranes of living cells. The rigid structures thus formed would permit the study of ( a ) isothermal ‘freezing’ of cellular membranes; ( 6) cellular ‘capsules’ containing entrapped cytoplasmic and intrinsic membrane proteins; (c) permanently asymmetric lipid bilayers; and ( d ) lateral discon- tinuities in membrane fluidity that would accompany localized polymer formation.The biosynthetic incorporation of diacetylenic fatty acids into membrane phos- pholipids and glycolipids was accomplished with a fatty acid auxotroph of the bacterium Acholeplasma laidlawii. When A. laidlawii cells were grown in the presence of diacety- lenic fatty acids, up to 90% of the membrane acyl chains were derived from the medium.24 Shorter-chain diacetylenic fatty acids were most suitable as substrates for the growth of these cells, and resulted in the most extensive uptake of the lipid. The distribution of the C20-diacetylenic fatty acid was very similar to that obtained for cells grown on a monounsaturated fatty acid, oleic acid.Similarities in uptake, in capacity to support growth, and in distribution among lipid classes suggest that the diacetylenic lipids are arranged in the bilayer in a manner similar to the oleate-containing lipids.114 Spec? roscop ic Studies and Phospholipid Polymers Polymerization of the membrane lipid in A. laidlawii was accompanied by a loss in the activity of an intrinsic membrane protein, NADH oxidase. In contrast, the activity of an extrinsic membrane protein, ribonuclease, was unaffected by p~lymerization.~~ Blodgett and Langmuir first demonstrated that a multilayered coating of lipid could be deposited onto a solid support by successively dipping the support through a m ~ n o l a y e r .~ ~ As with most models of biological membranes, instability is the primary limitation of multilayered films. Diacetylenic phospholipids pose a significant advantage for the utilization of multilayered films; polymerization inhibits the rearrangement and decay evident with multilayers of non-polymerizable lipids. We have developed procedures for the preparation of Langmuir-Blodgett-type multi- layers of diacetylene-containing Multilayers of diacetylenic phos- pholipids consistently presented a hydrophobic surface if they were polymerized after withdrawal from the subphase. In situations involving biocompatible surfaces, however, it is sometimes important that the prosthetic surface be polar. A polar surface could be stabilized, however, by irradiating the multilayer under water.Alternatively, the diacetylenic film was replaced with stearic acid before the last upstroke. After irradiation the layer of stearic acid was washed away, exposing the underlying polar surface. Varied materials (glass, quartz, Perspex, Teflon and mica) have been coated with ordered layers of diacetylenic phosphatidylcholines in which the phosphocholine moiety formed the outer coated surface. Fig. 4(b) shows the visible spectrum of a multilayer at various irradiation times. An increase in absorption in both the visible and U.V. regions of the spectrum accompanied polymerization. The layers after polymerization were quite stable in aggressive media and, with some precautions, could be handled without damage. An alternative method for the preparation of coated surfaces has been described by Regen et al.27y28 This method involves the polymerization of vesicles composed of diacetylenic or methacryloylic phosphatidylcholine in the presence of an insoluble support.The polymerized vesicles were assumed to associate with the phase bondary between the aqueous medium and the solid support. The capacity to modify the surface properties of existing materials by deposition of polymerizable multilayers may find important biomedical applications. The mechanical and topological properties of the support can be retained while the interfacial properties are changed to mimic those of cellular surfaces. Biomimetic Surfaces Asymmetries in the distribution of phospholipid head groups have been found in a variety of cells.29 The accumulated evidence suggests strongly that in the case of blood cells, the observed lipid asymmetry serves a biological purpose in the maintenance of the delicate balance between haemostasis and thrombosis.The extracellular surfaces of the plasma membranes of blood cells are thromboresistant; in strong contrast, their cytoplasmic surfaces are highly thrombogenic. The simplest common feature of the blood-compatible surfaces is the presence of large quantities (up to ca. 90% of the total surface lipids) of phosphorylcholine-containing phospholipids. We have previously investigated the haemostatic potential of polymeric phos- phorylcholine surfaces in dispersi~n.~~*'' Polymerized liposomes of diacetylenic phos- phatidylcholines did not alter the recalcification clotting times of citrated, pooled normal plasma regardless of lipid concentration.This non-thrombogenic nature of polymeric phosphatidylcholine was not altered after incubation in human plasma in vitro for up to 1 week at 37 "C. We have now developed32 a panel of reactive species which should result in the covalent deposition of phosphorylcholine on a variety of solid substrates. The resultant monomolecular layer provides a self-assembling surface which, in theory, should resemble the lipid interface presented by blood cells. The hydrophilic, membrane-D. Chapman, D. C'Lee and J. A. Hayward 115 ( a ) parent compound ( b ) natural phosphatidylcholines X,=O--CH, 0 I II CH-0-C- [CH21n - CH, I CH,-0- C- [CH,l,-CH, II 0 ( c ) hydroxyl-reactive phosphorylcholines (I) x,= c1 CH3 I I CH3 X,= 0-CH2-CH2-O- Si-Cl x, = 0- x,= C I x,= 0- ( d ) reaction products I " m 0 CHI h + I1 1 I k* (IIP HO-Si k \ + (CH,),N-CH,-CH,-O-P-O-CH2-CH~O-Si-O-Si~ t& O CH, Fig.5. Structures of phosphorylcholine ( a ) and its natural ( b ) and synthetic ( c ) and ( d ) derivatives. In the phosphatidylcholines found in biological membranes, the phosphorylcholine head group is esterified to a diacylglycerol. Two hydroxy-reactive phosphorylcholines have been prepared ( c ) , choline dichlorophosphate (I) and phosphatidylcholine ethylene glycol dimkthylsilylchloride (11). The immobilised species formed by reaction with silicon hydroxide are shown ( d ) . mimetic character of this surface provides opportunities for the generation of new, hybrid biomaterials.The modified surfaces retain the chemical, physical and topological properties of the substrates, with the intention of mimicking the characteristics of biomembranes (extreme thinness, low antigenic potential and increased potential for haemo- and bio-compatibility) (fig. 5 ) . Evidence for the covalent deposition of a self-assembing phosphorylcholine layer has been provided for the hydroxylated surfaces of glass and silica.33 Structural integrity of the deposited group is supported by the equimolar association of phosphorus and116 choline with the reacted surfaces. Covalent modification of the treated surfaces is demonstrated by i.r. spectroscopy. The modified surfaces are thermostable at tem- peratures up to 375 "C for extended Surfaces modified by the deposition of phosphorylcholine should exhibit characteris- tics highly desirable for biomaterials.The teleological arguments for possible haemo- compatibility have already been presented and are based upon the resemblance of phosphorycholine surfaces to the thromboresistant lipid surfaces of human blood cells. Additionally, the zwitterionic character of phosphorylcholine, attributed to the presence of both a strongly basic quaternary ammonium ion and an acidic phosphate ion of approximately equal strength, may impart useful physicochemical properties to the surface. This ionic balance may alter the adhesion of proteins and cells to modified surfaces. Further studies of these treated and other biologically relevant properties are in progress.Spectroscopic Studies and Phospholipid Polymers We thank the Wellcome Trust, The Commission of the European Communities and the Humane Research Trust for financial support. References 1 D. G. Cameron, H. L. Casal and H. H. Mantsch, J. Biochem. Biophys. Methods, 1979, 1, 21. 2 D. Chapman, J-C. Gomez-Fernandez, F. M. Goni and M. Barnard, J. Biochem. Biophys. Methods, 1980, 3 H. L. Casal and H. H. Mantsch, Biochim. Biophys. Acta, 1984, 779, 381. 4 M. Cortijo and D. Chapman, FEBS Lett., 1981, 131, 245, 5 M. Cortijo, A. Alonso, J-C. Gomez-Fernandez and D. Chapman, J. Mol. Biol., 1982, 157, 597. 6 D. C. Lee, A. A. Durrani and D. Chapman, Biochim. Biophys. Acta, 1984, 769, 49. 7 H. Susi and D. M. Byler, Biochem. Biophys.Res. Commun., 1983, 115, 391. 8 D. C. Lee, J. A. Hayward, C. J. Restall and D. Chapman, Biochemistry, 1985, 24, 4364. 9 D. C. Lee, D. A. Elliot, S. A. Baldwin and D. Chapman, Biochem. SOC. Trans., 1985, 13, 684. 2, 315. 10 D. H. MacLennan, C. J. Brandl, B. Korczak and N. M. Green, Nature (London), 1985, 316, 696. 1 1 Yu. N. Chirgadze, 0. V. Fedorov and N. P. Trushina, Biopolymers, 1975, 14, 679. 12 R. Henderson and P. N. T. Unwin, Nature (London), 1975, 257, 28. 13 B. K. Jap, M. F. Maestre, S. B. Hayward and R. M. Glaeser, Biophys. J., 1983, 43, 81. 14 P. Chakrabarti and H. G. Khorana, Biochemistry, 1975, 14, 5021. 15 J. H. Fendler, Science, 1984,, 223, 888. 16 G. Wegner, Makromol. Chem., Makromol. Chem., 1972, 154, 35. 17 D. S. Johnston, S. Sanghera, M. Pons and D. Chapman, Biochim. Biophys. Acta, 1980, 602, 57. 18 D. S. Johnston, L. R. McLean, M. A. Whittam, A. D. Clark and D. Chapman, Biochemistry, 1983, 22, 19 M. Pons, D. S. Johnston and D. Chapman, Biochim. Biophys. Acta, 1982, 693, 461. 20 M. Pons, D. S. Johnston and D. Chapman, J. Polyrn. Sci., Polyrn. Lett. Ed., 1982, 20, 513. 21 M. Pons, C. Villaverde and D. Chapman, Biochim. Biophys. Acta, 1983, 730, 306. 22 0. Albrecht, D. S. Johnson, C. Villaverde and D. Chapman, Biochim. Biophys. Acta, 1982, 687, 165. 23 J. Leaver, A. Alonso, A. A. Durrani and D. Chapman, Biochim. Biophys. Acta, 1983,732,210. 24 J. Leaver, A. Alonso, A. A. Durrani and D. Chapman, Biochim. Biophys. Acta, 1983, 727, 327. 25 K. B. Blodgett and I. Langmuir, Phys. Rev., 1937, 51, 964. 26 L. R. McLean, A. A. Durrani, M. A. Whittam, D. S. Johnston and D. Chapman, Thin Solid Films, 27 S. L. Regen, P. Kirszensztejn and A. Saingh, Macromolecules, 1980, 16, 335. 28 S. L. Regen, BV. Czech and A. Singh, J. Am. Chem. SOC., 1980, 102, 6638. 29 R. F. A. Zwaal and H. C. Hemker, Huemostasis, 1982, 11, 12. 30 J. A. Hayward and D. Chapman, in Biocompatibility of Tissue Analogs, ed. D. F. Williams (CRC Press, 31 J. A. Hayward and D. Chapman, Biomaterials, 1984, 5, 135. 32 A. A. Durrani, J. A. Hayward and D. Chapman, Biomaterials, 7, 121. 33 J. A. Hayward, A. A. Durrani, C. J. Shelton, D. C. Lee and D. Chapman, Biomaterials, 1986, 7, 126. 3192. 1983, 99, 127. Boca Raton, Florida, 1985). Received 13th December, 1985

ISSN:0301-7249    

DOI:10.1039/DC9868100107

出版商:RSC    

年代:1986

数据来源: RSC

摘要:

Faraday Discuss. Chem. SOC., 1986,81, 117-126 A Study of Phospholipid Phosphate Groups in Model Membranes by Fourier Transform Infrared Spectroscopy Felix M. Goii* and Jose L. R. Arrondo Departamento de Bioqur'mica, Facultad de Ciencias, Universidad del Par's Vasco, Aptdo. 644, 48080 Bilbao, Spain The phosphate region (1000-1300 cm-') of the infrared spectrum of aqueous phospholipid dispersions has been studied by the infrared Fourier transform technique. The main features of this spectral region have been described for various phospholipids, including phosphatidylcholine, phosphatidyl- ethanolamine, cardiolipin and phosphatidylglycerophosphate (alkyl ether), the major phospholipid of Halobacterium purple membranes. No changes in the phosphate region are observed due to lipid polymorphism or as a consequence of changes in fatty acyl chain structure. Shifts in the phosphoryl stretching bands are interpreted in terms of changes in hydrogen bonding, while a contribution from R-0-P vibration is considered to reflect changes in phospholipid headgroup conformation.When added in equimolar amounts to phosphatidylcholine or phosphatidylethanolamine bilayers, sur- factants (Triton X-100, sodium cholate) modify the degree of hydration and/or the orientation of the headgroup with respect to the bilayer plane. Phospholipids containing two phosphate groups give rise to more complex spectral features. Infrared data suggest that the two phosphate groups of cardiolipin are conformationally non-identical when incorporated into a lipid bilayer.Controlled enzyme hydrolysis (e.g. with phospholipase D) may help in the study of these complex phospholipid headgroups. The study of model membranes consisting of pure phospholipid bilayers has been the object of much effort in the last two decades. In particular, the application of spectro- scopic techniques to such model systems has shed light on a number of physical properties of phospholipids in aqueous dispersions which, in turn, give us some insight into the structure and behaviour of cell membranes.' 1.r. spectroscopy was one of the first physical techniques to be applied to the study of phospholipids;2 however, the strong absorption of water in the i.r. region prevented further applications of this technique to aqueous membrane dispersions until the advent of computerized spectroscopy, when spectral subtraction of water was made possible.At that stage, both interfer~metric~ and monochromator4 i.r. spectrometers were applied to model and biomembrane studies. Recent reviews576 have summarized the salient features of these investigations. A compulsory initial step in the i.r. investigation of biomembranes consists of the unequivocal assignment of spectral absorption bands and absorption maxima from the main membrane components. However, this objective is still far from being accom- plished. Methylene stretching vibration bands have been extensively characterized, partly because of their sharp appearance, partly because of their sensitivity to ther- motropic phase transitions, the most widely studied of all phospholipid properties.Other spectral regions, however, have not been explored in so much detail. One such region corresponds to the antisymmetric and symmetric stretching vibrations of the PO 2 bond, between 1000 and 1300 cm-'. 117118 I. R. Studies of Phospholipids Phosphate stretching in oriented multilayers was studied by Fringeli and GU~~thard;~ some phosphate stretching frequencies in natural and model biomembranes have also been described.*-’’ In a recent study,12 we investigated the 1000-1300 cm-’ region of the i.r. spectrum of aqueous dipalmitoylphosphatidylcholine (DPPC) and other phos- phate-containing molecules (glycerophosphorylcholine, phosphorylcholine, L-a- glycerophosphate etc.) by the Fourier transform technique. Buffered DPPC displays two maxima, at 1086 and 1222 cm-’, corresponding, respectively, to symmetric and asymmetric PO, stretching; these values are the same above and below the gel-to-liquid crystalline T, transition temperature of the phospholipid.The present paper summarizes the main results from a series of similar studies carried out with other naturally occurring phospholipids, i.e. phosphatidylethanolamine, cardiolipin and phosphatidylgly- cerophosphate (alkyl ether), the major phospholipid of Halobacterium purple membranes. Experiment a1 1,2-Dipalmitoyl-sn-glycero-3 -phosphocholine (DPPC) and 1,2-dimyristoyl-sn-g1ycero- 3-phosphoethanolamine (DMPE) were purchased from Fluka and their purity was checked by thin-layer chromatography and differential scanning calorimetry. Cardiolipin was from Sigma, egg-yolk phosphatidylcholine (EPC) was purified according to Single- ton et d;’’ bacterial phosphatidylethanolamine (BPE), from E.coZi, was type V from Sigma. The purity of lipids from natural origin was checked by thin-layer chromatogra- phy. Purple membrane from Halobacterium halobium was prepared as described by Muga14 and their major phospholipid (I) purified, in the form of sodium salt, according to Kates et aZ.” The buffer used throughout this work was 10 mmol dm-3 Hepes, pH 7.0. Phospholipid suspensions (30 mg cm-’) were prepared, unless otherwise stated, accord- ing to Cortijo et ~ 1 . ’ ~ When required, purple membrane lipids, in the form of sonicated suspensions, were treated with phospholipase D (type 111, from Sigma) in order to obtain a phospholipid (11) containing a single phosphate group.The resulting lipid was 0 R-O-CH2 II R-0-CH 0 I I I I 11 I I II CH2-O-P-O-CH, I I 0- 0- R-0-CH2 CH20- P-0- I CH2-0-P-0- R-0-CH 0 OHCH 0- R = phytyl chain R = phytyl chain (1) (11) purified by preparative thin-layer chromatography (A. Muga, to be published), and resuspended in Hepes buffer as above. Phospholipid-surfactant mixtures were prepared in organic solvents then freeze-dried overnight. The solvent-free mixture was later resuspended in water to a final phospholipid concentration of 30mgcmP3, at a tem- perature well above T, of the pure lipid with vortexing. The samples were introduced into a thermostatted microcell (Beckman FH-O1C FT) with CaF2 windows and a pathlength of 7 pm. A 10-MX Nicolet f.t.i.r. spectrometer was used; 512 spectra were routinely accumulated and averaged with a standard resolution better than 2 cm-’.Spectral subtraction of pure buffer was performed in order to remove the scattering background.F. M. Go% and J. L. R. Arrondo 119 wavenumber/cm-' Fig. 1. F.t.i.r. absorbance spectra of aqueous dispersions of ( a ) EPC and (b) BPE, in the region between 1000 and 1300 cm-', at 19 "C. Results and Discussion Studies with Phosphatidylcholine and Phosphatidylethanolamine Various preparations of phosphatidylcholine and phosphatidylethanolamine of natural and synthetic origin have been examined in order to determine the effect of phosphate group substituents and fatty acyl chains on the phospholipid phosphate vibrations. The 1000-1300 cm-' region of the i.r.spectrum of aqueous BPE and EPC at 19 "C is shown in fig. 1. EPC shows two main absorption bands, with maxima at 1086 and 1221 cm-', and a shoulder at 1060 cm-'; these are virtually the same values found previously for DPPC,12 that were assigned, respectively, to asymmetric and symmetric PO, stretching, and to the R-0-P-0-R' group. The spectrum of BPE looks very similar, except that the symmetric band is shifted towards lower frequencies and the R-0-P-0-R' signal now appears as a well resolved band, the three maxima being located at 1026, 1076 and 1220 cm-'. Fookson and Wallach17 examined dry films of DPPC and DPPE and found a significant shift (32 cm-') between the asymmetric PO, stretching bands of both substances, DPPE being shifted to lower frequencies, but no difference is found in our case, when fully hydrated samples are observed.The shift was attributed to intermolecular hydrogen bonding of PE. It is possible that, in aqueous suspension, both PC and PE bind extensively to water through hydrogen bonds. Mendelsohn and Mantsch'* mention the preseace of a symmetric ester C-0 stretching vibration band centred at 1070k3 cm I , while the phosphate ester stretch (C-0-P) would be located at 1047* 1 cm-' for most phospholipids. In our previous paper,12 we demonstrated the contribution of the C-0-P stretching vibration to the 1060 cm-' signal in fully hydrated DPPC bilayers (1069 cm-' for anhydrous DPPC). A contribution from the C-0 stretching vibration to the same signal cannot be ruled out, h~wever.~ Mantsch et al.19 have been able to characterize both a gel-to-liquid crystalline and a liquid-to-inverted hexagonal phase transition of egg-yolk phosphatidylethanolamines by i.r.spectroscopy through changes in CH2 stretching vibrations. Fig. 2 shows the120 I. R. Studies of Phospholipids 0.59E 0 .LL 7 0 s 2 0.298 si -2 0.1.49 0.000 1300 1200 1100 1000 wavenumber/cm-' Fig. 2. F.t.i.r. absorbance spectra of aqueous dispersions of BPE at ( a ) 12, ( b ) 19 and (c) 50 "C. phosphate region of the i.r. spectrum of aqueous BPE at 10, 19 and 50°C. These temperatures correspond, respectively, to phosphatidylethanolamine in the gel phase, the liquid-crystalline phase and the inverted hexagonal phase. (Phase transitions were monitored through changes in the symmetric CH2 stretching vibration of BPE; data not shown.) No variation in the maximum wavenumber of any of the bands is observed as a result of the phase transitions.The gel-to-liquid crystalline transition of DPPC also failed to produce any change in the i.r. phosphate spectrum.12 The influence of the nature of fatty acyl chains on the PO, stretching vibration bands has also been explored. We have seen that the phosphate region of spectra from EPC and DPPC are virtually undistinguishable (fig. 1). The same is true of dimyristoylphos- phatidylcholine in aqueous dispersion. Similar results are obtained when the spectra of BPE and DMPE are compared in the temperature range between 10 and 60°C; in no case did changes in fatty acyl chains modify the phosphate vibration bands of the i.r. spectrum of aqueous phospholipid (spectra not shown).The Interaction of Surfactants with Phospholipid Headgroups Surfactants are commonly used for membrane solubilization in the process of purification and reconstitution of integral membrane proteins; they also find an application as 'activators' of lipase enzyme activities. The details of surfactant-phospholipid interac- tions are unknown, although they are important in order to rationalize many studies on membrane reconstitution and phospholipase enzyme kinetics. We have examined equimolar mixtures of DPPC or DMPE with a surfactant, either Triton X-100 (a non-ionic detergent) or sodium cholate (a bile salt) both widely used in biochemical studies. According to a variety of physical and biochemical data,20.2' the amounts of surfactantF.M. Go% and J. L. R. Arrondo 121 wavenumber/cm-' Fig. 3. F.t.i.r. absorbance spectra of aqueous dispersions of DPPC and DPPC-surfactant mixtures. ( a ) Pure DPPC; ( b ) DPPC-Triton X-100, equimolar mixture; ( c ) DPPC-sodium cholate, equimolar mixture. used in the present study are not enough to produce bilayer solubilization, although they do produce dramatic changes of bilayer fluidity and permeability, together with high increases in phospholipase activity. According to our i.r. spectroscopic studies, surfactants modify the phosphate vibra- tion modes of phospholipids. Fig. 3 shows the phosphate region of the i.r. spectrum of DPPC, DPPC:Triton X-100 and DPPC:cholate at 50°C, i.e. in the fluid state. Each surfactant acts in its own way.Triton X-100 [fig. 3(b)] shifts the two main phosphate bands of DPPC towards higher frequencies, from 1220 to 1239 cm-' and from 1085 to 1090 cm-', respectively; also, the relative intensity of the R-0-P-0-R' shoulder at 1060 cm-' is obviously decreased; finally, a sharp band with a maximum at 1014 cm-' appears. The observed phosphoryl shifts are characteristic of hydrogen bonding22 and strongly suggest a decrease in hydrogen bonding to water of the phosphate group in the presence of Triton X-100. We had previously observed similar shifts in DPPC samples with various degrees of hydration. l 2 Surfactant-dependent changes in the R-0-P-0-R' shoulder are very interesting since the orientation of this part of the molecule strongly depends on the headgroup conformation.The observed shoulder is the result of coupling between the two P-0-C vibrations, depending in turn on headgroup packing, hydrogen bonding etc.17 Different transition vectors of the two P-0-C groups as the headgroup changes from a parallel to a perpendicular orientation122 I. R. Studies of Phospholipids wavenumber/cm-' Fig. 4. F.t.i.r. absorbance spectra of aqueous dispersions of DMPE and DMPE-surfactant mixtures. ( a ) Pure DMPE; ( b ) DMPE-Triton X-100, equimolar mixture; (c) DMPE-sodium cholate, equimolar mixture. with respect to the bilayer plane would also give rise to changes in the shoulder at 1060 cm-'. In summary, changes in phosphate spectral features in the presence of Triton X-100 can be interpreted in terms of a conformational change of the DPPC headgroup, involving a decrease in phosphate hydrogen bonding to water.The effect of sodium cholate on the phosphate spectrum of DPPC [fig. 3 ( c ) ] is smaller than that of Triton X-100. The intensity of the R-0-P-0-R' shoulder is decreased, but no shifts in the phosphate stretching vibration bands are observed. If the above interpretation is correct, this would mean that cholate does induce some kind of conformational change in the DPPC headgroup without altering the degree of hydration of the phosphate groups. This is in agreement with the different structure proposed for DPPC-Triton X- 100 and DPPC-cholate mixed micelle~.~~ Our results concerning DMPE-surfactant mixtures are shown in fig. 4. These experi- ments were carried out at 50 "C, i.e.with the bilayer in the fluid state, and are therefore comparable to those in fig. 3. Both surfactants act in this case in a similar way. Their main effect consists of inducing the appearance of much fine structure in the spectral bands, of the kind seen in dehydrated phospholipid samples. Why this fine structure becomes apparent in the presence of surfactants in excess water cannot be easily explained. The surfactants are also responsible for the presence of a new shoulder at the high-frequency side of the asymmetric PO, band, at 1250 cm-'. This new band inF. M. Go% and J. L. R. Arrondo 123 0. 312 0.294 e, E .fl 2 0.156 3 0.078 0.000 1 1300 1220 1140 1060 980 w aven u m be r/ cm - Fig. 5. F.t.i.r. absorbance spectrum of an aqueous dispersion of cardiolipin in the region between 1000 and 1300 cm-'. sucn a nigniy resoivea spectrum may well correspona to tne nign-rrequency sniIt ooservea in DPPC [fig.3 ( b ) ] ; also the low-frequency side of the asymmetric PO, band of DMPE-Triton X-100, but not that of DMPE-cholate mixtures, is clearly depressed as compared to that of pure DMPE (fig. 4), as was the case with DPPC and DPPC-surfactant mixtures (fig. 3). Both surfactants decrease the relative intensity of the DMPE band at 1010 cm-', attributed to R-0-P-0-R' vibrations, as was also the case with DPPE. Therefore, surfactants appear to act in a similar way in DMPE and DPPC headgroups, changing the orientation of the P-0-C bonds and, at least in some cases, reducing the extent of hydrogen bonding between phosphate and water.Phospholipids Containing Two Phosphate Groups We have extended our study of the phospholipid phosphate vibrations to the case of DhosDholiDids containing two DhosDhate crouDs. We shall first examine cardiolioin. a I 1 I " & I Y I 1 , phospholipid containing two chemically identical phosphate groups, and then proceed to phosphatidylglycerophosphate (alkyl ether), bearing two non-identical phosphates. The 1000-1300 cm-' region of the i.r. spectrum of cardiolipin is shown in fig. 5 . As in the previous cases, two main bands are seen. The one corresponding to the asymmetric PO, stretching vibrations has a maximum at 1215 cm-'; this would correspond to a high degree of hydrogen bonding17922 and, given the structure of this phospholipid, it is tempting to speculate that even in water dispersion some degree of intramolecular hydrogen bonding may exist.The lower frequency band is very wide and complex; at least three components are resolved, with maxima at 1092, 1072 and 1044 cm-', respec- tively. The maximum at 1092cm-' reveals most probably the contribution from the PO, symmetric vibration, while the other two maxima may be attributed to the four C-0- P vibrations in this phospholipid. The details of cardiolipin headgroup confor- mation in bilayers are not known, but it is possible that, if not chemically, both phosphate groups are conformationally non-identical with respect to the bilayer plane. This would explain the complexity of cardiolipin spectral features in the region around 1050 cm-'.124 I. R. Studies of Phospholipids 0 -097 0.073 6) r: -fi 0.048 B D 0.024 o * o o o 1300 1220 1140 1060 980 wavenumber/cm-' Fig.6. F.t.i.r. absorbance spectra of aqueous dispersions of phospholipids. ( a ) Phosphatidyl- glycerophosphate (alkyl ether); ( b ) phosphatidylglycerol (alkyl ether). The spectrum of phosphatidylglycerophosphate (alkyl ether), the major phospholipid in purple membranes, (I) can be seen in fig. 6 ( a ) . In this phospholipid the two phosphate groups are clearly different, one of them being monoesterified, and the other diesterified. However, the phosphate region of the i.r. spectrum is not very different from cardiolipin, with an asymmetric PO, band (maxima at 1215 cm-') and a complex band at lower frequencies, showing a maximum at 1064 and shoulders at 1038, 1085 and 1113 cm-'.Anhydrous films of the same phospholipid allow the resolution of this band into four peaks. In this case, the complexity of the band is easily understood, since there are three obviously different C-0-P groups in the molecule. Enzymic digestion of the purple membrane phospholipid provides us with a new tool for the study of i.r. phosphate vibrations. We have recently developed a method14 for cleaving an L- a-glycerophosphate moiety from phosphatidylglycerophosphate (alkyl ether), by using phospholipase D. As a result of enzyme action, an ether analogue of phosphatidic acid (11) is produced. The phosphate region of the corresponding i.r. spectrum is shown in fig. 6 ( 6 ) . Two striking features of this spectrum are: (a) the high frequency shift of the asymmetric PO, stretching band, with a maximum at 1273 cm-', and (6) the greatly simplified low-frequency band, confirming that the wide band seen in curve (a) contained the contributions from two other C-0-P groups.It is puzzling, however, that the spectrum of phospholipid (11) looks very different from that of its diester homologue, i.e. phosphatidic acid. In fact, the two maxima in the spectrum of the latter occur at 1181 and 1076crn-'.'* Among the factors that may explain such differences we should mention the smaller possibility of hydrogen bonding when the ester carbonyl groups are absent, and the sensitivity of monoesterified phosphate groups to changes in pH or counterions. Conclusions The results summarized here constitute a further step in the application of Fourier transform i.r.spectroscopy to the study of model and biomembranes. This techniqueE M. Go% and J. L. R. Arrondo 125 Table 1. Maximum wavenumbers of the main bands appearing in the 1000-1300 cm-’ region of f.t.i.r. spectra of aqueous phospholipid dispersions compound maximum wavenumber /cm-l assignment DPPC DMPE phosphatidic acid cardiolipin phosphatidylglycerophosphate (alkyl ether) 1060sh 1086 1180sh 1222 1013 1076 1177sh 1221 1076 1181 1044 1072 1092 1136 1215 1038 1064 1085 1113 1215 R-0-P-0-R’ symmetric PO, stretch asymmetric PO, stretch symmetric PO, stretch R-0-P-0-R’ asymmetric PO, stretch symmetric PO, stretch asymmetric PO, stretch R-0-P-0-R(?) R-0-P-0-R‘ symmetric PO, stretch asymmetric PO; stretch R-0- P-0 - R’( ?) R-0-P-0-R symmetric PO, stretch asymmetric PO; stretch provides spectra of excellent quality from dilute samples in a short time and buffer, temperature and other sample conditions can be regulated easily.In our case, the technique has been applied to the description of the phosphate region of phospholipid i.r. spectra. Phospholipids have been studied in the form of liposomes. For some of them, tentative assignments of bands have been made (table 1). Our results show that phosphate stretching vibrations are not influenced by the length or unsaturation of fatty acyl chains, nor by lipid polymorphism (in excess water). However, the nature of the phospholipid headgroup, or the presence of surfactants, does influence the phosphate vibration bands. The contribution from the R-0-P-0-R’ appears to be particularly sensitive to conformational changes, C-0- P vibrational bands suggest that the two phosphate groups of cardiolipin are conformationally non-identical when the phos- pholipid is integrated in a bilayer.More complex lipids, such as the major phospholipid of Hulobucterium purple membrane, may also be studied by this technique; enzymic hydrolysis under controlled conditions provides an additional tool for the analysis of the resulting complex spectra. We thank CAICYT (grant no. 0992-84) and Diputaci6n Foral de Vizcaya (O.F. no. 2813/85) for support. References 1 Biological Membranes, ed. D. Chapman (Academic Press, New York, 5 vols, 1968-1985). 2 D. Chapman, R. M. Williams and B. D. Ladbrooke, Chem. Phys. Lipids, 1967, 1, 445.3 D. G. Cameron, H. L. Casal and H. H. Mantsch, J. Biochem. Biophys. Methods, 1979, 1, 21. 4 D. Chapman, J. C. G6mez-Fernhdeq F. M. Goiii and M. Barnard, J. Biochem. Biophys. Methods, 1980, 2, 315.126 I. R. Studies of Phospholipids 5 R. L. h e y and D. Chapman, in Biomembrane Structure and Function, ed. D. Chapman (Macmillan, 6 H. L. Casal and H. H. Mantsch, Biochim. Biophys. Acta, 1984, 779, 381. 7 V. P. Fringeli and H. H. Gunthard, in Membrane Spectroscopy, ed. E. S. Grell (Springer, Berlin, 1981), 8 D. G. Cameron and H. H. Mantsch, Biochem. Biophys. Res. Commun., 1978, 83, 886. 9 H. H. Mantsch, D. G. Cameron, T. A. Tremblay and M. Kates, Biochirn. Biophys. Acta, 1982, 689, 63. 10 H. H. Mantsch, S. C. Hsi and D. G. Cameron, Biochim. Biophys. Acta, 1983, 728, 325. 11 K. J. Rothschild, W. J. De Grip and R. Sanches, Biochim. Biophys. Acta, 1980, 596, 338. 12 J. L. R. Arrondo, F. M. Goiii and J. M. Macarulla, Biochim. Biophys. Acta, 1984, 794, 165. 13 W. S. Singleton, M. S. Gray, M. L. Brown and J. C. White, J. Am. Oil Chem. Soc., 1965, 92, 52. 14 A. Muga, M.Sc. Thesis (University of the Basque Country, Bilbao, 1984). 15 M. Kates, S. C. Kushwaha and G. D. Sprott, in Methods in Enzymology, ed. S. P. Colowick and N. 0. 16 M. Cortijo, A. Alonso, J. C. G6mez-Fernkndez and D. Chapman, J. MoZ. BioL, 1982, 157, 598. 17 J. E. Fookson and D. F. H. Wallach, Arch. Biochem. Biophys., 1978, 189, 195. 18 R. Mendelsohn and H. H. Mantsch, in Protein-Lipid Interactions, ed. A. Watts (Elsevier, Amsterdam, 19 H. H. Mantsch, A. Martin and D. G. Cameron, Biochemistry, 1981, 20, 3138. 20 M. A. Urbaneja, A. Alonso, J. L. R. Arrondo and F. M. Goiii, Surfactants in Solution (Plenum Press, 21 D. Lichtenberg, Y. Zilberman, P. Greenzaid and S. Zamir, Biochemistry, 1979, 18, 3517. 22 L. J. Bellamy, Advances in Infrared Group Frequencies (Methuen, London, 1968). 23 A. Helenius and K. Simons, Biochim. Biophys. Acta, 1975, 415, 29. London, 1983), pp. 199-256. pp. 270-332. Kaplan (Academic Press, New York, 1982), pp. 98-111. in press). New York, 1986). Received 23rd December, 1985

ISSN:0301-7249    

DOI:10.1039/DC9868100117

出版商:RSC    

年代:1986

数据来源: RSC