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Departments of Chemistry and Physics, University of Washington, Seattle, Washington 98195-1700 USA
Correspondence: Address reprint requests to Sarah L. Keller, University of Washington, Seattle, WA 98195-1700. Tel.: 206-543-9613; Fax: 206-685-8665; E-mail: slkeller{at}chem.washington.edu.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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; Brown and London, 1998; McConnell and Vrljic, 2003; Silvius, 2003). In cell membranes, lipid rafts are currently thought to be localized regions that are on the order of 100 nm in diameter in which certain proteins and lipids are concentrated (Simons and Ikonen, 1997). Both the raft domains and the surrounding lipid matrix are liquid (Pralle et al., 2000). Lipid rafts have been associated with important biological processes such as endocytosis, adhesion, signaling, protein transport, apoptosis, and cytoskeleton organization (Deans et al., 2002; Edidin, 2003; Pike, 2003; Tsui-Pierchala et al., 2002). Since rafts in cell membranes have not been directly observed by standard microscopy, most current assays employ either indirect methods (e.g., detergent resistance), or cross-linking of rafts into larger aggregates (e.g., colocalization) (Thomas et al., 1994).
Lipid-driven lateral separation of immiscible liquid phases is likely a factor in the formation of rafts in cell membranes. Our laboratory and other groups have recently used fluorescence microscopy to directly observe coexisting liquid phases in GUV membranes containing ternary mixtures of saturated lipids, unsaturated lipids, and cholesterol (Dietrich et al., 2001; Samsonov et al., 2001; Veatch and Keller, 2002). This model system has the advantage that large, micron-scale domains can be directly observed by fluorescence microscopy, that miscibility transition temperatures can be measured, and that liquid phases can be studied from a more controlled physical perspective than in cells. Since vesicles can be made without proteins, the lipid contribution to membrane phase separation is isolated. Liquid immiscibility in vesicles has also been probed at smaller length-scales using methods such as FRET and NMR (Vist and Davis, 1990; Wang and Silvius, 2003; Xiaolian and London, 2000).
In earlier work, we varied cholesterol composition against a fixed 1:1 ratio of saturated to unsaturated phospholipids. We observed coexisting liquid phases and mapped the miscibility phase boundary (Veatch and Keller, 2002). Our work left us with clear questions. What are the lipid compositions of the two liquid phases? Over what range in phospholipid compositions are liquid domains observed? To address these questions, we have explored the ternary phase diagram of the saturated lipid DPPC (di(16:0)PC), the unsaturated lipid DOPC (di(18:1)PC), and cholesterol.
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MATERIALS AND METHODS |
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Vesicles
The methods are as described in Veatch and Keller (2002) and are repeated here. GUVs of diameters 10–50 µm were prepared in >18 M
/cm water as described by (Angelova et al., 1992) with modifications to increase yield and compositional uniformity. Specifically, 0.25 mg of lipids were dissolved in chloroform to 
5 mg/ml and spread evenly onto the conducting side of an ITO-coated glass slide. Lipid-coated slides were placed under vacuum for at least 30 min to remove remaining solvent. A capacitor was made from a lipid-coated and uncoated slide coupled with a 0.3-mm Teflon spacer, filled with water, and sealed with vacuum grease. Vesicles were grown for at least 1 h at 60 ± 3°C, then stored warm for at most 2 h before observation. Lipid samples were checked for oxidation by UV/VIS absorption and thin layer chromatography, and no oxidized lipids were detected to within 0.5%. The growth temperature was chosen to be the maximal temperature at which a high yield of GUVs was consistently obtained. Low growth temperatures can result in altered lipid composition. This was observed as a change in transition temperature throughout the sample and was most noticeable near steep regions of the miscibility transition boundary.
Although deposited lipids were uniformly mixed, each vesicle varied slightly in composition. This could be observed as different miscibility transition temperatures and brightness (mol % dye) between vesicles. This composition error is estimated to be <2 mol %. In all cases, we report lipid compositions as prepared in organic solvent before GUV growth. We have not confirmed that resulting GUVs have identical lipid compositions.
To view the sample, vesicle solution was placed between two coverslips and sealed with vacuum grease to prevent evaporation. Heat sink grease was used to thermally couple the coverslip sandwich to a temperature stage. An Alpha-Omega temperature controller was used with a home-built temperature stage consisting of a thermoelectric heater/cooler and a thermistor temperature probe (0.2°C accuracy, Sensor Scientific, Fairfield, NJ). The thermistor was thermally glued to an identical coverslip attached with heat sink grease beside the sample. The same thermistor was used for all experiments, and manufacturer temperature curves were used without further calibration. The accessible temperature range for the sample was between 10°C and 50°C. Epifluorescence microscopy was accomplished with a Nikon Microscope (Melville, NY) with an air objective that did not touch the sample. The depth of field was on the order of 10 µm so that either the top or bottom hemisphere of a vesicle was in focus at one time. Frames were captured with a high sensitivity Photometrics FX CCD camera (Roper Scientific, Tucson, AZ). Transitions were recorded as the temperature at which recognizable domains appeared and then disappeared as temperature was decreased and then increased. Experimental errors in transition temperatures had a systematic contribution of ±1°C from the response time of the thermistor and a standard deviation from averaging data from at least 10 vesicles. Since miscibility transitions were measured with both increasing and decreasing temperature, error bars represent upper and lower bounds on the true equilibrium miscibility transition temperature even if equilibrium conditions are not met.
Under the microscope, prolonged exposure to light led to photo-oxidation of the few vesicles in the field of view. The effects of photo-oxidation were most noticeable near the miscibility transition and included an increase in the transition temperature and the formation of new, smaller domains in vesicles that were already phase separated. Vesicles that were outside the field of view in the same sample were not affected. To avoid photo-oxidation, transition temperatures were measured by either scanning through temperature at a moderate rate (0.2°C/s) or stepping at a slow rate (<0.5°C/min) while minimizing light exposure. Both methods produced identical transition temperatures within the reported error bounds.
Potential pitfalls
Although we observe well-defined phase boundaries at some lipid compositions (e.g., between regions B and E in Fig. 2 a), we have not established other phase boundaries as precisely and document here all the possible pitfalls for completeness. To start our list, at the boundary between regions D and E, solid and liquid phases are difficult to distinguish when the surface fraction of dark phase is very small (at high DOPC concentrations) and when dark liquid domains are viscous and take long times to become circular (at low cholesterol and/or low temperatures). Similarly, it is difficult to observe solid domains that have elongated into thin lines due to a reduced line tension at increased cholesterol concentration (Korlach et al., 1999). The other side of the diagram, with low DOPC concentration, presents similar difficulties. Small bright phase domains are difficult to distinguish, and domains can take a long time to become circular because viscosity of the dark phase is significant. This is most apparent at low cholesterol and/or low temperatures. In addition, it is likely that a transition to a solid phase exists below the miscibility transition. This transition is challenging to distinguish since our fluorescent probe partitions away from both solid and the ordered liquid phase. At this time, we know of no fluorescent probe in the visible range that preferentially partitions into the ordered liquid phase. Solid transitions were not a focus of this study, but may lead to mischaracterization of liquid coexistence as solid-liquid coexistence when the liquid coexistence temperature range is narrow. Last, as the two-component DPPC-Chol line is approached, the number of DOPC molecules in the bright phase approaches the number of fluorescent probe molecules, which may skew miscibility transition temperatures.
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RESULTS |
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; Veatch and Keller, 2002). The relationship between the main chain transition temperature and liquid immiscibility is remarkably linear for saturated PC lipids in mixtures with DOPC and Chol (Fig. 1 a). In addition, transition temperatures in membranes containing pure SM lipids in mixtures with DOPC and Chol fall close to this line. ). This may be why SM lipids are commonly associated with lipid rafts, given that asymmetric lipids such as POPC are present in large quantities in most biological membranes.
Liquid immiscibility phase boundary for DPPC/DOPC/Chol ternary mixtures
In the following study, we outline the miscibility phase boundary for ternary mixtures of DPPC, DOPC, and Chol. We have chosen the DPPC/DOPC/Chol system for the following reasons:
; Lentz et al., 1976, 1980; McMullen and McElhaney, 1995; Shimshick and McConnell, 1973; Vist and Davis, 1990). ). Ongoing work using FRET and fluorescence microscopy to study membranes containing SM, DOPC, and cholesterol is producing similar phase boundaries (Kahya et al., 2003; Smith et al., 2003). In POPC/PSM/Chol mixtures, coexisting liquid domains are observed by FRET at high POPC concentrations presumably due to interactions between POPC and cholesterol (de Almeida et al, 2003).
Phase morphology
The phases we observe in the DPPC/DOPC/Chol system are roughly outlined in Fig. 2 a. This map records the first nonuniform phase to be observed at different compositions as temperature is lowered from a high temperature region of one uniform phase. Hence, Fig. 2 a contains information about more than one temperature and is not a phase diagram. To make this clear, Fig. 2 a is analogous to viewing a hilly landscape from above, rather than taking a cut through that landscape at one elevation.
Fig. 2 a contains five distinct regions. We will describe regions A through D before coming to our region of interest, the liquid-liquid coexistence observed in region E. At a high mole fraction, cholesterol is expelled from vesicles such that high cholesterol compositions are inaccessible in region A (Bacha and Wachtel, 2003; Huang et al., 1999). Nevertheless, the majority of the ternary composition space is accessible. Excluding region A and starting at a high temperature above the chain melting temperature of DPPC, all vesicles are observed to be in one uniform liquid phase. As temperature is lowered, the mixtures face four possible outcomes. First, they can remain in one uniform liquid phase as in region B. This occurs when vesicles contain a large fraction of unsaturated lipid (DOPC), greater than 50% Chol, or contain binary mixtures of DPPC and Chol as shown in Fig. 2 a, micrographs 2 and 3.
In the three remaining regions (C, D, and E), phases other than one uniform liquid are observed below the chain melting temperature of DPPC. In region C, in the lower right corner of the diagram at large fractions of saturated lipid, lipids are in a solid phase. Traveling left along the DOPC-DPPC axis into region D, there is solid-liquid coexistence. Phase diagrams of similar binary mixtures of high and low temperature phospholipids have been studied both experimentally and theoretically (e.g., Bagatolli and Gratton, 2000, and references therein). Boundaries of the solid-liquid phases can be described in terms of freezing point depression (Ipsen and Mouritsen, 1988). For small increases in cholesterol within region D, the freezing point is further depressed. The coexistence of solid and liquid phases in vesicles and the disruption of solid by higher cholesterol compositions have been documented previously (Feigenson and Buboltz, 2001; Korlach et al., 1999).
The last possible outcome as temperature is lowered is that the vesicle enters a region of coexisting liquid phases (region E). This region is the primary focus of this study. The liquid-liquid coexistence region E is distinguished from the solid-liquid coexistence region D by its domain morphology. Solid domains in region D strongly exclude the fluorescent dye, are generally noncircular, and rotate as rigid bodies in a background of liquid phase (Fig. 2 a, micrograph 1). In contrast, we observe that the liquid phases in region E exhibit circular domains that merge by continuously deforming when domains collide (Fig. 3 a).
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We know that as temperature is decreased through the miscibility transition, lipid acyl chain order should increase. Indeed, we frequently observe that vesicle area decreases near the miscibility phase transition. Our conclusion that the dark phase is more ordered than the bright phase is supported by two observations. First, as temperature is lowered, the surface fraction of dark phase increases. This increase is monotonic, but not a simple function of temperature, and will be discussed in a subsequent manuscript. Second, bright domains on a dark background diffuse more slowly than dark domains on a bright background. This difference becomes more pronounced at low temperatures.
We observe strong partitioning of TR-DPPE into the less ordered liquid phase over the entire temperature and composition range explored in this manuscript. Estimates from fluorescent micrographs produce partition coefficients KP < 0.4. This is consistent with results using a similar dye, rhodamine-DMPE, in DMPC/Chol membranes. In that system, the probe also partitioned away from the more ordered state with a coefficient of Kp = 0.30 at 30°C (Loura et al., 2001).
Domain kinetics and shapes
Immediately after temperature is lowered into a region of coexisting liquid phases, many small domains are observed. If membranes are given enough time under most experimental conditions, vesicles completely phase separate into a single dark region and a single bright region (Fig. 3 a). Domains grow by colliding and coalescing with other domains and not through Ostwald ripening. Merged domains quickly return to a circular shape (Fig. 3 a). This indicates that line tension is important and explains why we observe domain sizes comparable to vesicle dimensions (10 µm). Since the domains are large, bright and dark domains are truly separate phases. When viscosity in the membrane is high, the ripening of domains occurs slowly. We frequently observe slow ripening when the dark phase occupies most of the membrane and the vesicle is at a low temperature. Under these conditions domain size is kinetically trapped at a size smaller than the equilibrium size.
In the special case of vesicles with roughly equal bright and dark surface fractions, domains form via striping as shown in Fig. 3 b when temperature is decreased through the transition. This striping is either a kinetic effect caused by spinodal decomposition or an equilibrium effect due to a competition between line tension and long-range repulsive forces in the membrane. Both effects occur near critical points. Spinodal decomposition occurs when a miscible system is quenched below the spinodal curve, leading to fundamental instabilities in the one phase system. Certain wavelength composition fluctuations undergo exponential growth under these conditions, forming domains that are elongated and have a characteristic length-scale (Kahn, 1968). Since line tension is significant, thin elongated domains evolve into thicker stripes and then circles as time progresses (Vlatimirova et al., 1999) (Fig. 3 b).
In similar lipid monolayer systems, striping is observed at equilibrium close to a critical point. In these membranes, an interplay between the repulsive dipole interaction and the attractive interfacial line tension leads to stripes of a stable width (Keller and McConnell, 1999). Since photo-oxidation makes it difficult to view a vesicle at a constant offset between the vesicle temperature and the miscibility transition temperature, we cannot definitively test which mechanism is responsible for elongated domains. Evidence against equilibrium striping in bilayers include 1), a lack of a known long-range repulsive force between domains in bilayer membranes, and 2), with the exception of the unique mixture discussed below, uniform striping of domains is not observed when temperature is raised through the miscibility transition.
For the unique case of vesicles with 1:9 DOPC/DPPC + 25% Chol, striped domains are observed when temperature is raised (Fig. 3 c, left series). As discussed earlier, such vesicles have bright domains against a background of dark, ordered, viscous phase, and we believe that the stripes are due to viscous fingering. In this membrane, circular domains elongate near the miscibility transition, and stripe widths are uniform throughout the vesicle at each time frame. As time progresses, stripes thin until domains are no longer visible in vesicles. Fingers appear to have uniform widths, and fractal-like branching is not observed. Viscous fingering is also observed in vesicles with low line tension near the miscibility transition, but no uniform stripes are present. For example, the series on the right of Fig. 3 c shows that as temperature is raised, multiple thin bright fingers of no uniform width extend into the more viscous dark phase. As time progresses, fingers appear to thin and possibly branch below the resolution of our microscope (1 µm). This dependence of finger width on line tension is consistent with viscous fingering of liquid mixtures in Hele-Shaw cells (Arneodo et al., 1990; Wolf and Woermann, 1998). Further support that these structures are due to viscous fingering is that we do not observe fingering of dark phases into the less viscous bright phase.
Miscibility transition temperature
Fig. 4 shows the miscibility transition temperatures of vesicles of DPPC/DOPC/Chol. It is clear that the miscibility transition temperature is strongly dependent on membrane composition. The highest transition temperatures are found in vesicles rich in saturated lipids and cholesterol whereas vesicles with more unsaturated lipids form two liquid phases only at low temperatures.
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). In contrast, when vesicles contain more DPPC than DOPC, miscibility transition temperatures remain high with small error bars (Fig. 4 b, square symbols), and suddenly terminate near 50% cholesterol, where domains are no longer observed. A similar abrupt phase boundary can be seen when the DPPC/Chol axis is approached (Fig. 4 c). Plotted in three dimensions, the shape of the miscibility transition temperature surface in Fig. 4 a resembles a cliff that terminates near the DPPC/Cholesterol binary mixture.
Cholesterol depletion
Although most of this work has focused on using temperature changes to initiate liquid-liquid immiscibility in membranes, the same aim can be achieved by altering the lipid composition of vesicles at a constant temperature. Fig. 5 shows a vesicle that begins with high cholesterol composition in a region of one uniform liquid phase. When ß-cyclodextrin is added to the sample, the cholesterol content in the membrane is decreased and the vesicle enters a region of two liquid phases. As expected, bright circular liquid domains move freely on the dark background and then collide and coalesce. At long times, bright domains no longer diffuse even though they remain roughly circular. If, at this point, temperature is raised through the miscibility transition and then lowered back down, noncircular solid domains appear. This indicates that the previous immobile domains may have been kinetically trapped in that configuration (data not shown). Similar micron-scale liquid domains induced by ß-cyclodextrin have been observed in living cells (Hao et al., 2001). Our results in model vesicles may provide a framework for interpreting results in biological membranes.
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) and is further evidence that line tension is important in this system. Although we do not observe domain budding into the interior of the vesicle, we believe it is possible under different osmotic conditions. We have not further investigated vesicle bulging since we have learned that careful work on this topic is currently being conducted elsewhere (Baumgart et al., 2003).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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We observe two liquid phases, one bright and one dark, in region E of the ternary diagram in Fig. 2. Since we do not know the full composition of each phase, we cannot draw quantitative tie-lines. However, we do know the surface fraction of bright and dark phases in our vesicles. Translating this into a composition of each phase requires knowledge of the molecular area of each species in each phase. This is complicated by the condensing effect of cholesterol and lipids (Phillips, 1972). Nevertheless, from our qualitative results alone, we conclude that the bright phase is enriched in unsaturated lipid (DOPC) and the dark phase is rich in cholesterol and saturated phospholipids (DPPC).
We come to this conclusion by examining vesicles at a constant temperature in Fig. 2 b. First, comparing vesicles with equal fractions of cholesterol (20 mol %), the vesicle in micrograph 6 contains more DPPC and more dark phase than the vesicle in micrograph 4. Next, comparing vesicles with equal DOPC/DPPC ratios (2:1) the vesicle in micrograph 7 contains more cholesterol and more dark phase than the vesicle in micrograph 6. Given that our dark phase is rich in saturated lipids and cholesterol, it is likely equivalent to the liquid-ordered state described previously and may resemble detergent resistant phases in biological membranes (London and Brown, 2000).
We use similar logic to estimate the location of tie-lines in Fig. 2 b. Micrograph 4 shows a vesicle close to the miscibility transition boundary with a small surface fraction of dark phase. Assuming that there are only two phases present in the liquid-liquid coexistence region at 30°C, the lever rule implies that point 4 must lie near one end of a tie-line. Similarly, vesicles containing more saturated lipids and/or cholesterol contain large surface fractions of dark phase (points 7 and 8) and therefore reside near the opposite end of a tie-line in this picture. This implies that the tie-lines run roughly left to right in the shaded region shown in Fig. 2 b with a tilt toward more cholesterol in the DPPC-rich phase.
This tie-line orientation is also supported by our observation of points 5 and 6 in Fig. 2 b. Roughly equal surface fractions of bright and dark phase are present in these vesicles near the phase boundary, implying that these vesicles are near a critical composition. Further evidence in these vesicles is that fluctuations in domain boundaries near the miscibility transition are consistent with reduced line tension (as in Fig. 3 c, right series), and domains form by a process that resembles spinodal decomposition (as in Fig. 3 b). Concluding that points 5 and 6 are near critical points and assuming that only two phases are present in this region, then the tie-lines run roughly left to right with a slope toward more cholesterol in the DPPC-rich phase.
Tie-lines with a shallow slope suggest that cholesterol is more equally distributed between the phases than DPPC is. In fact, our recent results using 2H-NMR to quantify phase composition has shown that DPPC is highly asymmetrically distributed between phases (80% of DPPCd62 is found in the ordered phase of membranes containing 1:1 DPPCd62/DOPC + 30% Chol at 15°C). Cholesterol is more symmetrically distributed (2/3 of cholesterol (3 d1) is found in the more ordered phase in membranes of 1:1 DOPC/DPPC + 30% Chol (3 d1) at 20°C) (Veatch et al., 2003).
Comparison with limiting binary systems
The miscibility phase map in Fig. 2 a agrees well with previously published phase diagrams for binary mixtures along two of the three edges, for DOPC and Chol as well as for DPPC and DOPC. DOPC and Chol have been shown to mix well down to temperatures below which these experiments were conducted (Lentz et al., 1976). Indeed, we observe that vesicles made of primarily DOPC and cholesterol remain in one uniform phase at 10°C. Mixtures of DPPC and DOPC exhibit an extended gel-liquid coexistence region over the temperature range explored in these experiments (Davis and Keough, 1983). This phase behavior is also reproduced here over a wide temperature and composition range.
The relationship between the results presented in Fig. 2 a and previously published phase diagrams for DPPC and Chol membranes is less clear. At first, it seems that these results are in conflict. The lack of observable domains in GUVs with binary lipid compositions is perplexing since many other methods have reported coexisting liquid phases at high temperatures in binary mixtures of DPPC (or DMPC) and Chol. These methods all probe molecular length-scales and include NMR (Vist and Davis, 1990), DSC (McMullen and McElhaney, 1995), ESR (Shimshick and McConnell, 1973), FRET (Loura et al., 2001), fluorescence anisotropy, and freeze fracture (Lentz et al., 1980). By FRET, domain size has been limited to 20 nm at equilibrium in some regions of the liquid-liquid coexistence region of the DPPC/Chol phase diagram (Loura et al., 2001). This is well below the resolution of optical microscopy, and our results therefore are not in conflict.
This distinction between micron and nanometer scale phase behavior in binary mixtures of DPPC is nontrivial and requires a reexamination of the meaning of phase separation in this context. Certainly, lipid organization on the nanometer length-scale can be a result of two thermodynamic phases that may macroscopically phase separate. However, in some cases it may be a result of dynamic intermolecular interactions that exist within a single phase. For example, surfactants above the critical micelle concentration are considered to be in a single phase even though there is an equilibrium between surfactant monomers and nanometer-scale spherical micelles in solution. In other words, a single spherical micelle is not a separate phase (V. A. Parsegian, personal communication). In lipid monolayer systems, there is also evidence of nanometer-scale organization of saturated lipids and cholesterol in lipid membranes that are not phase separated (McConnell and Radhakrishnan, 2003). Whereas nanometer-scale organization in binary mixtures of DPPC and Chol may not always qualify as a true thermodynamic phase separation, it is probably related to the large-scale liquid immiscibility that we observe in ternary mixtures of DPPC, DOPC, and Chol.
Possible three-phase regions
It is also possible that three-phase regions exist for particular ternary compositions of DPPC, DOPC, and Chol. One candidate region has been proposed by others (de Almeida et al., 2003; Silvius et al., 1996; Smith et al., 2003) and is between the solid-liquid (D) and liquid-liquid (E) regions in Fig. 2 a. These two regions could be separated by a uniform liquid phase, a triple point, or an extended region in which three phases are in coexistence, one solid and two liquid. It is also possible that there exists a region of coexistence of three liquid phases in the middle of region E. Since we do not directly observe three-phase coexistence in this experiment, two of these phases would have to be indistinguishable by our fluorescent probe. This is plausible because most ordered phases of DPPC/DOPC/Chol exclude TR-DPPE. To investigate this further by fluorescence microscopy, new probes would have to be discovered that partition well into the liquid phase rich in DPPC and Chol, but not into the solid phase.
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CONCLUSION |
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). We are currently using NMR to investigate the DPPC/DOPC/Chol system, and our results will appear in a subsequent manuscript. |
ACKNOWLEDGEMENTS |
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This work was funded by a National Science Foundation CAREER Award (MCB-0133484), a Research Corporation Innovation Award, the University of Washington Royalty Research Fund, and the Petroleum Research Fund. Sarah Veatch was supported in part by a National Institutes of Health predoctoral training grant in molecular biophysics (5T32-GM08268-14) and a National Science Foundation IGERT fellowship from the University of Washington Center for Nanotechnology.
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FOOTNOTES |
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Submitted on April 28, 2003; accepted for publication July 9, 2003.
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