Liquid-Disordered

6289

2007, 111, 6289-6292 Published on Web 05/17/2007

Using Ionic Strength to Control Liquid-Ordered/Liquid-Disordered Separation Kalani J. Seu, Emily R. Lamberson, and Jennifer S. Hovis* Department of Chemistry, Purdue UniVersity, West Lafayette, Indiana 47907 ReceiVed: January 24, 2007; In Final Form: April 16, 2007

In this Letter, we will show that liquid-ordered/liquid-disordered separation can be controlled with ionic strength. Using this observation, a robust method was developed for creating visible, by fluorescence microscopy, liquid-ordered domains in supported lipid bilayers. The details of the method will be discussed.

Introduction Liquid-ordered/liquid-disordered separation has received considerable attention due to the suggestion that liquid-ordered domains localize important signaling molecules.1,2 For a variety of reasons, it is desirable to use supported lipid bilayers to examine aspects of liquid-ordered domain formation and function. On mica, it is possible to form large liquid-ordered domains by slow cooling the samples.3-6 On glass, the preferred substrate for fluorescence microscopy, it is also possible to form liquidordered domains that are visible (by fluorescence microscopy) by slow cooling; however, the reproducibility is less than optimal, ∼67% on etched slides and ∼50% on baked slides.7 In this paper, a method will be detailed that allows for the formation of visible liquid-ordered domains >95% of the time in bilayers on glass. Additionally, for some of the compositions examined, the bilayers do not lie completely flat but form capshaped structures. This last observation is of particular relevance, as cap-shaped liquid-ordered domains have been observed in giant unilamellar vesicles8,9 but not to our knowledge in supported lipid bilayers. The method presented herein makes use of the observation that liquid-ordered/liquid-disordered separation can be controlled with ionic strength. Results and Discussion When comparing liquid-ordered domain formation in giant unilamellar vesicles (GUVs) and supported lipid bilayers (SLBs), there are two significant differences that need to be considered. First, in GUVs, the domains are free to move and coalesce, creating ever larger domains, while, in SLBs, the domains are typically pinned. Second, GUVs are typically heated and cooled slowly, while SLBs are typically formed by vesicle deposition onto substrates below the mixing temperature. Due to these differences, it has been assumed that domains visible by fluorescence microscopy can be formed in GUVs but not SLBs. However, Longo and co-workers have shown that if SLBs are heated above the mixing temperature and slowly cooled, large liquid-ordered6 and gel-phase3-5 domains can be formed in SLBs. Domain size is largely determined by the nucleation density; from classical nucleation theory, it is known that a low * Corresponding author. E-mail: [email protected]. Phone: (765) 4944115. Fax: (765) 494-0239.

10.1021/jp070624s CCC: $37.00

nucleation rate, and consequently large nuclei, is observed just below the mixing temperature. In previous work, we were able to form visible liquid-ordered domains in SLBs that had been heated and cooled only ∼67% of the time;7 while not explicitly stated, many samples were slow cooled and this did not improve the reproducibility. This work differed from the previous work in two important aspects: the substrates were different (glass versus mica) and the lipid composition was different. As glass is the substrate of choice for fluorescence microscopy, we wanted to see if the reproducibility could be improved. It is known that ions can affect the phase transition temperature of lipids.10 Therefore, to eliminate the heating step, we decided instead to vary the ionic strength of the solution. With the following method (method 1), >95% reproducibility was achieved; examples are shown in Figure 1. Step 1: Bilayers were formed by vesicle fusion on glass slides that had been piranha etched for 5 min; this treatment method resulted in the lowest frictional coupling.7 The vesicles were formed in 18 MΩcm water and immediately prior to fusion mixed 1:1 with a 100 mM NaCl buffer solution (50 mM 4-(2-hydroxyethyl)-1piperazineethanesulfonic acid (HEPES), 0.1 mM ethylenediaminetetraacetic acid (EDTA), pH 7.4). It is possible to do the fusion in 18 MΩ-cm water; however, mixing with a little salt helps to make the process more consistent. After fusion, the excess vesicles were rinsed away; at this point, very faint, small, liquid-ordered domains were frequently observed. Step 2: The bulk solution was then exchanged for one containing 500 mM NaCl (preliminary work indicates that concentrations from 100 mM to 1 M can be used); at this point, the bilayers were always uniform in appearance. Bilayers were incubated in this salt solution for 15 min, although varying the incubation time from 5 to 20 min does not significantly affect the results. Step 3: The bulk solution was exchanged for one containing 0 mM NaCl (water); at this point, liquid-ordered domains were visible >95% of the time. Once formed, the liquid-ordered domains can be made to disappear and appear by increasing and decreasing the ionic strength of the bulk solution. Representative images of bilayers composed of two different compositions with two different probes are shown in Figure 1: (A and B) 1:1 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC)/ 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) with 30 mol % cholesterol; (C and D) 1:1 DOPC/1,2-distearoyl-snglycero-3-phosphocholine (DSPC) with 30 mol % cholesterol. © 2007 American Chemical Society

6290 J. Phys. Chem. B, Vol. 111, No. 23, 2007

Figure 1. Epifluorescence images of bilayers treated with method 1: (A and B) 1:1 DOPC/DPPC with 30 mol % cholesterol; (C and D) 1:1 DOPC/DSPC with 30 mol % cholesterol. In parts A and C, the bilayers contained 0.5 mol % Texas Red DHPE, while, in parts B and D, the bilayers contained 0.5 and 1 mol % tail-labeled NBD-PC, respectively. Images were acquired using a 40×, 0.75 NA, air objective and are 40 µm × 40 µm.

In parts A and C, the bilayers contained 0.5 mol % Texas Red 1,2-dihexanoyl-sn-glycero-3-phosphoethanolamine (DHPE), while, in parts B and D, the bilayers contained 0.5 and 1 mol % taillabeled 1-palmitoyl-2-[6-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl]-sn-glycero-3-phosphocholine (NBD-PC), respectively. The miscibility transition temperatures of these mixtures are ∼29 °C (1:1 DOPC/DPPC with 30 mol % cholesterol) and ∼35 °C (1:1 DOPC/DSPC with 30 mol % cholesterol).11,12 The salt exchange was done at room temperature; consequently, the domains form at ∼6 and ∼12 °C below the miscibility temperature. Critical nucleation theory predicts that the bilayers containing DSPC would have more nucleation sites than those containing DPPC; this will be discussed further at the end. Rings were rarely observed in the 1:1 DOPC/DPPC with 30 mol % cholesterol and 0.5 mol % Texas Red DHPE mixtures and were almost always observed in the other mixtures. In Figure 1A, circular liquid-ordered domains are clearly visible; it has previously been shown that this probe depletes out of the liquid-ordered domains;11 in this case, the depletion is ∼50%. It is possible that the domains are only in the top leaflet. To determine if this is the case, several samples were examined before and after the solution was exchanged for one containing 50 mM NaI. Iodine quenches the Texas Red emission and can be used to determine the distribution of the Texas Red lipids between the leaflets.13 The intensity decreased 30 ( 6%, indicating that ∼70% of the Texas Red lipids are in the lower leaflet, and thus, the domains must be in both leaflets.14 To confirm that the regions are liquid-ordered domains and not gelphase domains, the regions were photobleached and rapid recovery was observed. On rare occasions, these domains have been observed to move and coalesce; see the Supporting Information for an example. Both circular dark domains and bright rings are visible in Figure 1B-D. To determine if the rings appear because the bilayer has curved away from the surface, a 100×, 1.45 NA, oil-immersion objective was used to image a bilayer composed

Letters

Figure 2. (A-F) Epifluorescence images of a 1:1 DOPC/DSPC with 30 mol % cholesterol and 1 mol % NBD-PC bilayer. A 100×, 1.45 NA, oil-immersion objective was used to acquire the images; the focus was gradually adjusted away from the glass slide (A-F). Images are 20 µm × 20 µm. (G and H) Line scans at all focus levels (A-F) of the lines indicated in part A. The top left line is shown in part G, and the bottom right line is shown in part H. A schematic illustration of the bilayer with the corresponding focal planes is shown in part I.

of 1:1 DOPC/DSPC with 30 mol % cholesterol and 1 mol % NBD-PC. With this objective, it is possible to focus through three-dimensional structures, Figure 2. Two of the dark regions in Figure 2A are marked with red lines; the line scans across these regions for each focus level are shown in parts G (top left line) and H (bottom right line) of Figure 2. As the focus is changed, the region indicated by the top line disappears from view (background levels are subtracted from the line scans). We conclude that this region, and the others like it, is a liquid-ordered domain lying in the flat part of the bilayer. The lower region changes dramatically as the focus is adjusted; it is clearly a cap-shaped structure; a schematic illustration is shown in Figure 2I. Because this is an epifluorescence experiment, and not a confocal experiment, there is background noise from all heights. Consequently, the counts do not drop to zero at the center in Figure 2A, when the focus is under the cap, or in Figure 2B and C, when the focus is in the middle of the caps. In a previous paper, we observed very similar structures (in a different lipid composition) with the same objective (100×, 1.45 NA).15 Examining those samples with confocal microscopy confirmed that the structures were caps.15 In giant unilamellar vesicles, liquid-ordered domains have been observed to form cap-shaped structures.8,9 We suggest that the caps in Figures 1 and 2 are liquid-ordered domains. The NBDPC does not completely deplete from the liquid-ordered domains; consequently, the caps will remain bright even if they are liquid-ordered domains. In the line scan, Figure 2H, it can be seen that the intensity of the ring never exceeds the intensity of the flat region, when it is in focus, Figure 2A. The rings are collecting fluorophores from many height levels; if the fluorophore doping was the same as the flat region, the rings would be expected to be significantly more intense. In Figure 2B and

Letters C, it appears as though some of the rings are touching one another. They were not observed to merge with one another; however, the samples were only examined for at most ∼1 h which may not have been long enough to observe merging. In recent work,9 it was shown that in giant unilamellar vesicles the cap-shaped liquid-ordered domains merge very slowly; this is postulated to be due to repulsive interactions between the domains. The method outlined above (method 1) requires three steps. To simplify it, a two-step method was explored (method 2). Step 1: Vesicles were reconstituted, fused, and incubated in a high NaCl solution (concentrations from 100-500 mM were investigated). Step 2: Excess vesicles were rinsed away, and the bulk solution was exchanged for one containing 0 mM NaCl (water). After this step, domains were rarely observed. Possible reasons why this simpler method does not work as well are detailed below. On the proximal side of the bilayer, there is a high ionic strength solution; this may prevent liquid-ordered domains from forming in the lower leaflet. In method 1, the bilayers were formed in a 50 mM NaCl solution; however, there is evidence that the concentration of ions trapped between the bilayer and the support depends not on the ionic concentration in solution but on the charge density of the support.16 If so, then the bilayers formed with either method should have the same ionic strength trapped between the bilayer and the support. Many studies have suggested that there is strong coupling between the leaflets; if domains can form in the top leaflet, they might be able to drive formation in the lower leaflet regardless of ionic strength. A second and more probable reason that method 1 yields better results than method 2 is simply that better bilayers are formed when vesicles are osmotically stressed prior to fusion.17,18 No supported bilayer is completely defect-free, but a fairly uniform, continuous bilayer is essential for the mixing of lipids that leads to the formation of visible domains. We have observed that bilayers formed using method 2 will often show domains after the bulk solution has been cycled between 500 mM NaCl and 0 mM NaCl two to three times; see the Supporting Information. It is possible that initially the bilayer contains many defect sites and adsorbed vesicles. The ionic cycling may help the vesicles in the defect sites to fuse, filling in the defect sites and creating a nearly defect-free single bilayer. Due to entropic effects when bilayers are formed in 500 mM NaCl, there are frequently many extra vesicles sitting on top of the bilayer that are difficult to rinse away. Care needs to be taken with these samples; if the vesicles are not rinsed away, a wide variety of three-dimensional structures form when the ionic strength is dropped, likely due to the extra lipid material. In summary, we have observed that liquid-ordered/liquiddisordered separation can be controlled by ionic strength. Using this observation, we developed a robust method for creating visible liquid-ordered domains in supported lipid bilayers. There are at present two remaining questions, with the first one being the following: Why does salt disrupt the liquid-ordered domains? Liquid-ordered domain formation is controlled by a number of interactions that are sensitive to ionic strength: dipole-dipole, van der Waals, and hydration forces, among others. Adding salt may have the effect of decreasing the miscibility temperature, causing the lipids to mix; when the salt is removed, the miscibility temperature returns to one above the sample temperature and domains form. If so, it should be possible to control the nucleation density by incubating at different temperatures; the larger the temperature difference, the more nucleation sites. To examine if this is the case, a sample

J. Phys. Chem. B, Vol. 111, No. 23, 2007 6291

Figure 3. Epifluorescence images of a 1:1 DOPC/DPPC with 30 mol % cholesterol and 1 mol % Texas Red DHPE bilayer prepared using method 1. A 500 mM NaCl solution was exchanged for water at (A) 23 °C and (B) 10 °C; the images are of the same location on the bilayer. Images are 40 µm × 40 µm and were acquired at room temperature.

was prepared as in Figure 1A; 500 mM NaCl was exchanged for water, first at 23 °C, Figure 3A, and then at 10 °C, Figure 3B. In Figure 3B, there are clearly many more nucleation sites. There is also a larger surface area coverage; it is known that the larger the temperature drop from the miscibility temperature, the more lipids are involved in domain formation.12 The samples incubated at 10 °C were imaged rapidly but at 23 °C; however, given how slow domains move on supported bilayers, see the Supporting Information, this image can be considered representative of how the surface appeared at 10 °C. Many of the domains in Figure 3A appear at the same location as those in Figure 3B, suggesting that there is some heterogeneous nucleation. As mentioned previously, there should be more nucleation sites in the DSPC containing bilayers, Figure 1C and D, than in the DPPC bilayers, Figure 1A and B. Examining several samples, we see that this in general true; however, quantification is frustrated by the large polydispersity from one sample to the next (a larger sample size is needed). The polydispersity can be seen in Figures 1A and 3A which were formed by exactly the same method with the same composition. All of this indicates that on glass both heterogeneous and homogeneous nucleation occur. The second remaining question is the following: Why is salt incubation more robust than slow cooling in this system? One possibility is that upon heating the DPPC lipids leave the surface. With either slow cooling7 or ionic incubation (data not shown), domains are more frequently observed in bilayers on etched rather than baked glass slides, suggesting that the baked slides have more defects than the etched slides, resulting in many small domains below the diffraction limit. Acknowledgment. Jennifer S. Hovis is a recipient of a Career Award in the Biomedical Sciences from the Burroughs Wellcome Fund. We thank one of the reviewers for suggestions that led to the experiment in Figure 3 and Mandy Green for help with the quenching experiments. Supporting Information Available: Experimental details and images of domain movement and formation. This material is available free of charge via the Internet at http://pubs.acs.org. References and Notes (1) Brown, D. A.; London, E. J. Membr. Biol. 1998, 164, 103. (2) Simons, K.; Ikonen, E. Nature 1997, 387, 569. (3) Blanchette, C. D.; Lin, W. C.; Ratto, T. V.; Longo, M. L. Biophys. J. 2006, 90, 4466. (4) Lin, W.-C.; Blanchette, C. D.; Ratto, T. V.; Longo, M. L. Biophys. J. 2006, 90, 228.

6292 J. Phys. Chem. B, Vol. 111, No. 23, 2007 (5) McKiernan, A. E.; Ratto, T. V.; Longo, M. L. Biophys. J. 2000, 79, 2605. (6) Lin, W. C.; Blanchette, C. D.; Longo, M. L. Biophys. J. 2007, 92, 2831. (7) Seu, K. J.; Pandey, A. P.; Haque, F.; Proctor, E. A.; Ribbe, A. E.; Hovis, J. S. Biophys. J. 2007, 92, 2445. (8) Baumgart, T.; Hess, S. T.; Webb, W. W. Nature 2003, 425, 821. (9) Yanagisawa, M.; Imai, M.; Masui, T.; Komura, S.; Ohta, T. Biophys. J. 2007, 92, 115. (10) Koynova, R.; Brankov, J.; Tenchov, B. Eur. Biophys. J. 1997, 25, 261.

Letters (11) Veatch, S. L.; Keller, S. L. Phys. ReV. Lett. 2002, 89, 268101. (12) Veatch, S. L.; Polozov, I. V.; Gawrisch, K.; Keller, S. L. Biophys. J. 2004, 86, 2910-2922. (13) Zhang, L. F.; Granick, S. J. Chem. Phys. 2005, 123. (14) We have observed the same distribution of Texas Red lipids in DOPC bilayers on etched supports. (15) Cambrea, L. R.; Hovis, J. S. Biophys. J. 2007, 92, 3587-3594. (16) White, R. J.; Zhang, B.; Daniel, S.; Tang, J. M.; Ervin, E. N.; Cremer, P. S.; White, H. S. Langmuir 2006, 22, 10777. (17) Reimhult, E.; Hook, F.; Kasemo, B. Langmuir 2003, 19, 1681. (18) Richter, R. P.; Be´rat, R.; Brisson, A. R. Langmuir 2006, 22, 3497.