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J. Phys. Chem. 1996, 100, 2766-2769
Slow Nonequilibrium Dynamical Rearrangement of the Lateral Structure of a Lipid Membrane Kent Jørgensen,† Alex Klinger,‡ Mark Braiman,‡ and Rodney L. Biltonen*,‡,§ Department of Physical Chemistry, Technical UniVersity of Denmark, DK-2800, Lyngby, Denmark; Department of Biochemistry and the Graduate Program in Biophysics, UniVersity of Virginia Health Sciences Center, CharlottesVille, Virginia 22908; and Department of Pharmacology, UniVersity of Virginia Health Sciences Center, CharlottesVille, Virginia 22908 ReceiVed: October 13, 1995; In Final Form: December 11, 1995X
The lipid bilayer of the biological membrane is a multicomponent molecular mixture capable of exhibiting compositional and structural heterogeneity. Using Fourier transform infrared spectroscopy and Monte Carlo computer simulation techniques, we report here for the first time the existence of a long-lived nonequilibrium heterogeneous lateral membrane structure composed of gel and fluid domains in a binary dipalmitoylphosphatidylcholine-dibehenoylphosphatidylcholine (DC16 PC-DC22 PC) lipid membrane. The nonequilibrium dynamic ordering process of coexisting phases following a thermal quench from the fluid state into the gelfluid phase coexistence region is characterized by a relaxation time on the order of hours. This slow process leads to a long-lived compartmentalized percolative lateral membrane structure with a dynamic network of interfacial regions having properties different from the coexisting gel and fluid bulk phases.
Introduction Lipid membrane heterogeneity and domain formation are considered to be of relevance for the functioning of biological membranes.1 In recent years it has become clear that the cooperative behavior of the lipid membrane which determines the global phase structure also leads to local nanometer structures and a dynamic heterogeneous membrane arising from equilibrium structural and compositional fluctuations of the many-particle system.2,3 By means of combined experimental and theoretical investigations of nonequilibrium ordering phenomena of coexisting gel and fluid phases, we have been able to show that nonequilibrium effects can give rise to a highly heterogeneous membrane structure resulting in domain formation on various length and time scales. FTIR spectroscopy reveals that upon quenching from the high-temperature fluid phase into the gel-fluid coexistence region the shorter-chain DC16 PC lipid initially becomes more ordered than in equilibrium but then relaxes slowly back to a more disordered state. These experimental results are corroborated by Monte Carlo simulations which show that gel clusters of the longer-chain DC22 PC lipid are formed rapidly, resulting in a significant amount of conformational order of the shorter DC16 PC at the domain interfaces. Subsequently, these clusters coalesce and phase separate resulting in less conformational order of the shorter DC16 PC. Such effects might play a significant role for the lateral organization of the lipids in biomembranes. Results and Discussion The equilibrium phase behavior of the DC16 PC-DC22 PC mixture was established using differential scanning calorimetry which demonstrated a high degree of nonideal mixing behavior giving rise to a broad gel-fluid coexistence region. The excess heat capacity function of a multilamellar equimolar DC16 PC†
Technical University of Denmark. Department of Biochemistry and Graduate Program in Biophysics, University of Virginia Health Science Center. § Department of Pharmacology, University of Virginia Health Science Center. X Abstract published in AdVance ACS Abstracts, January 15, 1996. ‡
0022-3654/96/20100-2766$12.00/0
Figure 1. Differential scanning calorimetry (DSC) excess heat capacity curve of a multilamellar dispersion of DC16 PC:DC22 PC vesicles (1:1 mole ratio) obtained at a scan reate of 10 °C/h using a high-sensitivity differential scanning calorimeter based on the heat leak principle.16 The lipid concentration was 40 mM, and the sample volume was 0.75 mL. DC16 PC and DC22 PC were purchased from Avanti Polar Lipids (Birmingham, AL). To form multilamellar vesicles, appropriate amounts of the lipid components were dissolved in chloroform and dried under high vacum overnight. The dried lipids were then resuspended in 50 mM KCl and 1 mM NaN3 above the phase transition temperature of the highest melting lipid, DC22 PC. The melting temperature of DC16 PC is 41.5 °C,21 and that of DC22 PC is 74.8 °C.22 The inital and final temperatures used in the thermal quench experiments are marked by the arrows in the figure.
DC22 PC lipid mixture shown in Figure 1 is indicative of the phase behavior of the binary lipid mixture. The transition centered at 32 °C is associated with the formation of the ripple phase, the most stable phase at temperatures just below the chain melting transition. The distinct separation between the sharp chain melting transition of the predominantly DC16 PC-rich gel phase centered at 41 °C and the melting of the predominantly DC22 PC rich gel phase at 68 °C indicates highly nonideal mixing of the two components in both the gel state and the gel-fluid coexistence region. The asymmetry on the low © 1996 American Chemical Society
Letters
Figure 2. (A, top) Monte Carlo simulations of the lateral structure of an equimolar DC16 PC-DC22 PC binary lipid mixture after a simulated temperature quench. Times indicated are in Monte Carlo steps (mcs). The initial snapshot (time e 0) shows an equilibrium configuration in the fluid phase (80 °C) prior to the temperature quench; small clusters of like lipids are noted as a results of the poor mixing properties of the lipids. Snapshots at time ) 400 and 8000 mcs are nonequilibrium membrane configurations in the gel-fluid phase coexistence region (48 °C). The simulations were carried out using a molecular interaction model, originally developed to describe chain melting of one-component phospholipid bilayers5 and now extended to binary systems by including a mismatch term taking into account the difference in acyl chain lengths.3 A detailed account of the procedure to calculate both equilibrium and nonequilibrium time-dependent properties is given in ref 4. The simulations reported here were performed on a lattice with 5000 lipid molecules. The properties of the system were calculated using both single-chain conformational kinetics and nearest-neighbor exchange kinetics for lateral diffusion. The symbols for the conformational states of the acyl chains are gel-DC16 PC (+ ), fluid-DC16 PC (blank), gel--C22 PC (b), fluid-DC22 PC (‚). (B, bottom) Calculated order parameters for the individual lipid species DC16 PC and DC22 PC in an equimolar mixture after a temperature quench from 80 to 48 °C. The acyl chain order parameter for each lipid species is given by S ) 1/2〈3 cos2 θ - 1〉 where θ is the angle between the bilayer normal and the normal to the plane spanned by a methylene group.23
temperature side of the melting curve of the DC22 PC is the result of partial mixing of the two lipids in the fluid phase. Monte Carlo simulations of the binary DC16 PC-DC22 PC mixture were performed using a modified molecular interaction model3,4 originally designed to describe the chain melting transition of one-component phospholipid bilayers.5 The simulation results following an instantaneous quench of the twocomponent system from a temperature corresponding to the fluid phase (80 °C) to a temperature in the gel-fluid phase coexistence region (48 °C) are shown in Figure 2. The snapshots in Figure 2A show equilibrium and nonequilibrium membrane configurations as obtained from simulations of the molecular interaction model. The equilibrium membrane configuration in the fluid state (80 °C, time e 0) displays clustering of like lipids due to the poor mixing properties of the DC16 PC and DC22 PC lipids induced by the acyl chain mismatch. Upon the
J. Phys. Chem., Vol. 100, No. 8, 1996 2767 instantaneous quench into the gel-fluid coexistence region (48 °C) formation of gel clusters of the longer DC22 PC lipid is observed (time ) 400). These clusters slowly grow and coalesce (time ) 8000) until macroscopic phase separation is eventually observed at equilibrium. The results of these simulations are represented quantitatively in Figure 2B in terms of an average chain order parameter for each lipid as a function of time (number of Monte Carlo steps). The average chain order parameter of DC22 PC achieves its equilibrium value rapidly, whereas that of DC16 PC overshoots the equilibrium value and then slowly decays toward the final equilibrium value. Detailed analysis of the simulations indicate that the overshooting effect and the degree of excess conformational order associated with the shorter DC16 PC lipid is related to partial ordering at the cluster interfaces (see snapshots of the membrane configurations (time ) 400, 8000) in Figure 2A). FTIR spectroscopy was used to observe the time courses of asymmetric and symmetric methylene stretching vibration bands that are representative of conformational order of the acyl chains in the membrane system.6 Perdeuterated DC16 PC was used in the equimolar lipid mixture in order to separate the acyl chain vibrations of DC16 PC from those of DC22 PC. The deuterated methylene (CD2 ) stretching vibrations of the perdeuterated DC16 PC appear at around 2100 cm-1, while the methylene (CH2 ) stretching vibrations occur at around 2800 cm-1 (Figure 3A).7 The difference spectrum calculated by subtracting the spectrum measured at 80 °C from the spectrum measured at 50 °C is shown in Figure 3B. The largest changes are seen in the bands due to the CH2 stretching vibrations at around 2900 cm-1. Much smaller are the difference bands due to the CD2 stretches at around 2150 cm-1. These analogous difference bands differ substantially in size because the extinction coefficient of a CD stretching vibration is about 0.7 times as strong as that of a CH stretching vibration; because there are only two-thirds as many CD2 groups in deuterated DC16 PC as CH2 groups in DC22 PC; and because the undeuterated DC22 PC changes state almost completely from fluid to gel conformation upon dropping the temperature from 80 to 50 °C, while the perdeuterated DC16 PC exhibits only a modest change in conformational state. The two sets of difference bands due to the acyl chain CH or CD stretching vibrations, now normalized to the maximum peak difference, are shown in Figure 3C,D. The time course for the asymmetric CH2 and CD2 stretch difference signals subsequent to a rapid temperature quench from 80 to 50 °C are shown in Figure 4. In Figure 4A, the uppermost curve represents the time course of the magnitude of the asymmetric CH2 stretch difference peak. The lowermost curve represents the time course for the analogous CD2 stretch difference peak drawn on the same scale. The middle curve represents a magnification (×22) of the lowermost curve of the time course for the CD2 stretch difference peak. Both the CH2 and the CD2 time courses have a similar initial fast rise. While the CH2 time course subsequently remains flat, the CD2 curve displays a slow decay. The details of the latter portion of the time courses are shown on an expanded scale in Figure 4B. Here the two curves are shown with the same vertical magnification, but the CD2 time course is shifted up 0.27 absorbance units so that the traces overlap. From this presentation, it is clear that the CH2 curve rises monotonically, while the CD2 curve is biphasic, overshooting and then slowly returning toward its equilibrium value. Analysis of the time course provides an estimate of a half-time of 50 min for the slow decay of the conformational state of the shorter DC16 PC. A remarkable similarity is observed by comparison of the Monte Carlo computer simulation results of the individual acyl chain
2768 J. Phys. Chem., Vol. 100, No. 8, 1996
Figure 3. FTIR difference spectra of a binary mixture of perdeuterated DC16 PC and undeuterated DC22 PC (1:1 mole ratio) multilamellar vesicles. Perdeuterated DC16 PC was obtained from Avanti Polar Lipids (Birmingham, AL). Vesicles were prepared as described in Figure 1. FTIR measurements were obtained at constant temperature on a sample taken from a spun pellet of the lipid vesicle suspension. The samples were squeezed between two CaF2 windows with a 6 µm Mylar spacer. The window sandwich was placed in a temperature-controlled flow cell attached to a water bath to maintain constant temperature. (A) FTIR spectra of the 1:1 mixture at 80 and 50 °C. (B) The difference spectrum of the sample obtained by subtracting the 80 °C spectrum from the spectrum obtained at 50 °C. (C) The difference spectrum of the CH2 region normalized to the maximum peak difference. (D) The difference spectrum of the CD2 region normalized to the maximum difference peak.
order parameters presented in Figure 2B and the FTIR results shown in Figure 4. Both figures demonstrate the same time development of the conformational state of the two lipid species after the quench. The longer DC22 PC lipid rapidly reaches its equilibrium value after the quench whereas the shorter DC16 PC lipid overshoots the equilibrium value and slowly decays toward the final equilibrium value. Similar overshooting behavior was observed using a lipophilic fluorescent probe (pyrene) and either a temperature drop or pressure increase to induce the transition from the fluid phase to the coexistence region. Conclusion Several techniques have been used to monitor the relaxation of single-component lipid bilayers following a rapid temperature or pressure perturbation. A wide range of relaxation times were observed (10-8-10 s), but, generally, the relaxation amplitude and time were maximal near the transition temperature.8 van OsDol et al.,9 using a volume perturbation technique, observed a single relaxation of 2-4 s. Ye and Biltonen10 found in a similar study with a binary lipid mixture that the mean relaxation rate increased by up to 3 orders of magnitude with the presence of only a few percent of the second component. Recently,
Letters
Figure 4. Time-dependent behavior of the size of the asymmetric and symmetric CH2 stretching difference bands after a temperature quench (A). In (B) the scale has been expanded and the difference peak of the CD2 vibration has been displaced (see text for details). The sample was equilibrated in the fluid phase (80 °C) until a stable baseline was obtained. At t ) 0, the temperature-controlled flow cell was quickly switched to the low-temperature bath. A Si diode sensor monitoring the sample cell indicated that the temperature drop was >75% complete within the first 60 s, and >98% complete within the first 150 s. Spectra were measured in a Nicolet 60SXR Fourier transform infrared spectrometer at 2 cm-1 resolution, averaging data over 2 min periods for 40-60 min at the initial temperature and for about 3 h after the temperature jump. Each spectrum contains the average of 256 interferograms. Time courses of difference spectra were calculated by averaging five spectra measured at the starting temperature and then subtracting this average from each of the spectra measured during and after the temperature jump. Time courses of difference peak size were calculated for the peaks in question by subtracting the absorbance difference value at the minimum of the negative band, ∆Amin, from the absorbance difference band at the maximum of the positive band, ∆Amax, at each time point. Note, that when the time-course results are presented on an expanded scale (4B), the CH2 curve appears to have a slower rise time than the CD2 curve. This, however, is not the case. In Figure 4B the portion of the CH2 curve shown is only about 0.2% of the total rise, whereas the portion of the CD2 curve shown accounts for about 6% of the total rise.
Snyder and co-workers11 reported slow lipid demixing in the gel state by monitoring the shape of the methylene scissors band but were unable to observe any demixing in the gel-fluid coexistence region. We have now shown for the first time, using a combined experimental and theoretical approach, that slow demixing of a binary lipid system exhibiting significant acyl chain mismatch can occur in the gel-fluid phase coexistence region. The nonequilibrium ordering process detected appears to be analogous to spinodal decomposition.12,13 Various experimental and theoretical studies have suggested the existence of domain formation in both biological and model membranes.2,3 These domains, which can vary greatly in size, geometry and lifetime, include those which exist as local structures in binary systems in the fluid region,14,15,16 domain formation in the phase coexistence region resulting from poor mixing of the lipid components,17,18 and clustering of lipids in binary systems induced by proteins or peptides19 which interact preferentially with one of the lipid species. Domain formation might be of importance for the activity of membrane processes, e.g., the activity of kinase C19 and the activation of phospholipase A2 in the gel state which appears to be associated with nonequilibrium dynamic domain formation of the reaction products.20 In addition local nonequilibrium phase separation and domain formation might be strongly related to membrane fission and fusion processes.13 The results reported in this letter indicate that long-lived nonequilibrium domains can exist in lipid membranes following thermodynamic, environmental, or compositional perturbations.
Letters In a biological membrane which operates at isothermal conditions, nonequilibrium states similar to those reported here following a thermal perturbation can be induced by a sudden change in, for examples, pH or ionic conditions. Nonequilibrium domain formation, which offers a restricted geometrical environment for membrane processes,1 may for that reason be of considerable importance for the functionality of biological membranes in which many processes take place in a nonequilibrium situation controlled by changes in fluxes of matter or energy. Acknowledgment. This work was supported by The Danish Natural Science and Technical Research Councils, The Danish Research Academy, Jenny Vissings Fond, The National Science Foundation, and the National Institutes of Health. References and Notes (1) Bergelson, L. O.; Gawrisch, K.; Ferretti, J. A; Blumenthal, R. Special Issue on Domain Organization in Biological Membranes, Mol. Membr. Biol. 1995, 12, 1. (2) Kinnunen, P. K. J.; Mouritsen, O. G. Special Issue of Functional Dynamics of Lipids in Biomembranes, Chem. Phys. Lipids. 1994, 73, 1. (3) Jørgensen, K.; Sperotto, M. M.; Mouritsen, O. G.; Ipsen, J. H.; Zuckermann, M. J. Biochim. Biophys. Acta. 1993, 1152, 135. (4) Jørgensen, K.; Mouritsen, O. G. Biophys. J. 1995, 69, 942. (5) Pink, D. A.; Green, T. J.; Chapman, D. Biochemistry 1980, 19, 349. (6) Casal, H. L.; Mantsch, H. H. Biochim. Biophys. Acta 1983, 779, 381. Mendelsohn, R.; Mantsch, H. H. In Progress in protein-lipid interactions 2, Elsevier: Amsterdam, 1989.
J. Phys. Chem., Vol. 100, No. 8, 1996 2769 (7) Casal, H. L.; Cameron, D. G.; Boulanger, Y.; Mantsch, H. H.; Smith, I. C. P. Biophys. J. 1981, 35, 1. (8) van Osdol, W.; Biltonen, R. L.; Johnson, M. L. J. Biochem. Biophys. Methods. 1989, 20, 1. Caffrey, M. Annu. ReV. Biochim. Biophys. Chem. 1989, 18, 159. (9) van Osdol, W.; Johnson, M. L.; Ye, Q.; Biltonen, R. L. Biophys. J. 1993, 59, 112. (10) Biltonen, R. L.; Ye, Q. Prog. Colloid. Surf. Sci. 1993, 93, 112. (11) Snyder, R. G.; Strauss, H. L.; Cates, D. A. J. Phys. Chem. 1995, 99, 8432. Mendelsohn, R.; Liang, G. L.; Strauss, H. L.; Snyder, R. G. Biophys. J. 1995, 69, 1987. (12) Jeppesen, C.; Mouritsen, O. G. Phys. ReV. B 1993, 47, 14724. (13) Sackmann, E.; Feder, T. Mol. Membr. Biol. 1995, 12, 21. (14) Knoll, W.; Ibel, K.; Sackmann, E. Biochemistry 1981, 20, 6379. (15) Ruggiero, A.; Hudson, B. Biophys. J. 1989, 55, 1111. (16) Mouritsen, O. G.; Jørgensen, K. Chem. Phys. Lipids 1994, 73, 3. (17) Almeida, P. F. F.; Vaz, W. L. C.; Thompson, T. E. Biochemistry 1992, 31, 7198. (18) Leckband, D. E.; Helm, C. A.; Israelachvili. J. Biochemistry 1993, 32, 1127. (19) Yang. L.; Glaser, M. Biochemistry 1995, 34, 1500. (20) Burack, W. R.; Yuan, Q.; Biltonen. R. L. Biochemistry 1993, 32, 583. Jain, M. K.; Yu, B. Z.; Kozubek, A. Biochim. Biophys. Acta 1989, 980, 23. (21) Suurkuusk, J.; Lentz, B. R.; Barenholz, Y.; Biltonen, R. L.; Thompson, T. E. Biochemistry 1976, 15, 1393. (22) Lewis, R. N. A. H.; Mak, N.; McElhaney, R. N. Biochemistry 1987, 26, 6118. (23) Ipsen, J. H.; Mouritsen, O. G.; Bloom, M. Biophys. J. 1990, 57, 405.
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