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Drying and Rehydration of DLPC/DSPC Symmetric and Asymmetric Supported Lipid Bilayers: a Combined AFM and Fluorescence Microscopy Study Sandra V. Bennun, Roland Faller, and Marjorie L. Longo* Department of Chemical Engineering and Materials Science, UniVersity of California-DaVis, California 95616 ReceiVed May 30, 2008. ReVised Manuscript ReceiVed July 21, 2008 This work characterizes the impact of lipid symmetry/asymmetry on drying/rehydration reorganization in phaseseparated dilauroylphosphatidylcholine (DLPC)/distearoylphosphatidylcholine (DSPC) supported lipid bilayers (SLBs) at the submicron and micron-scale. In addition the prevention of major drying/rehydration reorganization by the use of trehalose is demonstrated. Even though it was found using fluorescence microscopy that micrometer scale structure is preserved in the presence and absence of trehalose upon drying/rehydration, AFM and FRAP experiments successfully revealed major changes in the phase-separated structure such as defects, obstructions, lipid condensation, collapse structures, and complex incomplete DLPC-DSPC mixing/exchange in the absence of trehalose. In the presence of trehalose the membrane preserves its structure at the nanometer scale and mobility. We found that SLBs with asymmetric domain configurations underwent major rearrangements during drying and rehydration, whereas the symmetric domain configuration mainly rearranged during rehydration, that we hypothesize is related to lower transmembrane cohesiveness or lack of anchoring to the substrate in the case of the asymmetric domains.
1. Introduction Life without water or anhydrobiosis is a biochemical and biophysical adaptation response to adverse desiccation conditions entered by certain organisms that are able to regain function from a death-like state upon rehydration.1,2 Apparently the mechanisms of adaptation involve sugars, proteins as well as antioxidants and other molecules that prevent and reverse the structural and functional damage caused by loss of water. Among these molecules, the sugar trehalose was reported to be present in the majority of desiccation tolerant organisms and to play a fundamental role in anhydrobiotic adaptation.3 Trehalose activity as a cryoprotectant and biopreservation agent in the dry state is well-known; applications range from preservation of biomembranes to cells.4-8 Extensive research, has been devoted to reveal the protective mechanisms of trehalose.3,9-11 The water replacement3 and the vitrification hypothesis9 are the most accepted mechanisms and interestingly were proposed from studies on lipid vesicles.3-5,12,13 The vitrification hypothesis suggests that trehalose forms a protective glass that embeds liposomes and prevents their fusion and damage from bulk water removal.9 The high glass transition of Trehalose (Tg ) 113 °C), allows trehalose * To whom correspondence should be addressed. Phone: (530)754-6348. E-mail:
[email protected]. (1) J. H, Crowe., L. M. C., J. Carpenter, S. Petrelski., F. Hoekstra., P. de Araujo., A. D., Panek., Anhydrobiosis:Cellular adaptation to extreme dehydration. In Handbook of Physiology; Dantzler, W. H. , Ed.; Oxford University Press: New York, 1997; Vol 2, pp 1445-1477, (2) Crowe, J. H.; Oliver, A. E.; Tablin, F. Integr. Comp. Biol. 2002, 42, 497– 503. (3) Crowe, J. H.; Crowe, L. M.; Chapman, D. Science 1984, 223, 701–703. (4) Crowe, L. M.; Crowe, J. H.; Rudolph, A.; Womersley, C.; Appel, L. Arch. Biochem. Biophys. 1985, 242, 240–247. (5) Crowe, L. M.; Crowe, J. H.; Womersley, C.; Rudolph, A.; Uster, P. Biophys. J. 1985, 47, A248–A248. (6) Wolkers, W. F.; Walker, N. J.; Tablin, F.; Crowe, J. H. Cryobiology 2001, 42, 79–87. (7) Crowe, J. H.; Tablin, F.; Wolkers, W. F.; Gousset, K.; Tsvetkova, N. M.; Ricker, J. Chem. Phys. Lipids 2003, 122, 41–52. (8) Kanias, T.; Acker, J. P. Cell PreserV. Technol. 2006, 4, 253–277. (9) Green, J. L.; Angell, C. A. J. Phys. Chem. 1989, 93, 2880–2882. (10) Belton, P. S.; Gil, A. M. Biopolymers 1994, 34, 957–961. (11) Bryant, G.; Wolfe, J. Cryo-Letters 1992, 13, 23–36.
to be in a glassy state at higher temperatures and higher water contents compared to other saccharides.9,14-16 The other hypothesized mechanism, water replacement, suggests that trehalose depresses the main lipid transition temperature (Tm) in the dehydrated state by direct interaction with lipid head groups, most explicitly by hydrogen bonding of the OH groups of trehalose with the phosphate groups of the lipids.3,17,18 This preserves the lipid head separation in the dry state similar to the hydrated state and prevents the increase of van der Waals forces.3,12 Other hypotheses also explain trehalose interactions in the dry state, for instance Bryant and Wolfe11 suggest that volumetric and osmotic sugar effects reduce mechanical stresses preventing close approach of the bilayers, thus hindering a gel phase transformation when being dehydrated. In view of the complexity of the mechanisms that govern preservation in the dry state, a variety of model systems have been addressed.3,7,10,11,19-26 Of these biological cell membranes are of enormous interest, because of their implications in cell (12) Tsvetkova, N. M.; Phillips, B. L.; Crowe, L. M.; Crowe, J. H.; Risbud, S. H. Biophys. J. 1998, 75, 2947–2955. (13) Crowe, J. H.; Oliver, A. E.; Hoekstra, F. A.; Crowe, L. M. Cryobiology 1997, 35, 20–30. (14) Angell, C. A.; Bressel, R. D.; Green, J. L.; Kanno, H.; Oguni, M.; Sare, E. J. J. Food Eng. 1994, 22, 115–142. (15) Ding, S. P.; Fan, J.; Green, J. L.; Lu, Q.; Sanchez, E.; Angell, C. A. J. Therm. Anal. 1996, 47, 1391–1405. (16) Wolkers, W. F.; Oliver, A. E.; Tablin, F.; Crowe, J. H. Carbohydr. Res. 2004, 339, 1077–1085. (17) Sum, A. K.; Faller, R.; de Pablo, J. J. Biophys. J. 2003, 85, 2830–2844. (18) van den Bogaart, G.; Hermans, N.; Krasnikov, V.; de Vries, A. H.; Poolman, B. Biophys. J. 2007, 92, 1598–1605. (19) Adams, D. R.; Toner, M.; Langer, R. Langmuir 2007, 23, 13013–13023. (20) Crowe, J. H.; Crowe, L. M.; Carpenter, J. F.; Rudolph, A. S.; Wistrom, C. A.; Spargo, B. J.; Anchordoguy, T. J. Biochim. Biophys. Acta 1988, 947, 367–384. (21) Crowe, J. H.; Crowe, L. M.; Wolkers, W. F.; Oliver, A. E.; Ma, X. C.; Auh, J. H.; Tang, M. K.; Zhu, S. J.; Norris, J.; Tablin, F. Integr. Comp. Biol. 2005, 45, 810–820. (22) Ricker, J. V.; Tsvetkova, N. M.; Wolkers, W. F.; Leidy, C.; Tablin, F.; Longo, M.; Crowe, J. H. Biophys. J. 2003, 84, 3045–51. (23) Koster, K. L.; Lei, Y. P.; Anderson, M.; Martin, S.; Bryant, G. Biophys. J. 2000, 78, 1932–1946. (24) Chiantia, S.; Kahya, N.; Schwille, P. Langmuir 2005, 21, 6317–6323. (25) Albertorio, F.; Chapa, V. A.; Chen, X.; Diaz, A. J.; Cremer, P. S. J. Am. Chem. Soc. 2007, 129, 10567–10574.
10.1021/la8016694 CCC: $40.75 2008 American Chemical Society Published on Web 08/16/2008
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preservation. However, cells present heterogeneous membrane compositions and a still unresolved microstructure associated with the formation of microdomains, so-called rafts.27-29 This makes the characterization of cell membrane transformations resulting from desiccation very challenging. Therefore there is a necessity of addressing drying effects in well characterized systems. Chiantia et al.,24 performed AFM to characterize dehydration stresses on supported lipid bilayers (SLBs) of sphingomyelin/dioleoylphosphatidylcholine/cholesterol. Hydrated SLBs presented large area fraction of small liquid ordered phase domains. Without any stabilizing substance, major defects and structural damage were observed, such as extensive uncovered areas of mica (delamination), the presence of bulky and large structures, and irregular phase separation with no recognizable structure other than suggested large and irregular domains formed by aggregation of the smaller domains. The capability of membrane structure preservation by stabilizing agents was also evaluated. DMSO/dextran/ and glucose bilayers showed similar but less extensive damage of the bilayer structure. While only sucrose and trehalose provided good protection. In another study, Albertorio et al.,25 employed fluorescence techniques to study SLBs of 1-palmitoyl-2-oleoyl-phosphatidylcholine on glass. Dried and rehydrated lipid bilayers showed ∼100% delamination assessed by correspondence to nonfluorescence areas. On the other hand, R,R-trehalose and R,R-galacto-trehalose preserved the bilayer structure and two-dimensional lateral fluidity. This was attributed to the R,R-(1f1) glycosidic linkage and the clam shell structure that has the right size to span between two adjacent lipids head groups. Maltose and GM1 prevented delamination of the bilayers however lipid bilayers lost lateral mobility. Interestingly, more than 60% delamination, took place when the bilayer was preserved with other sugars (R,β-trehalose, sucrose, lactose, glucose) or contained other glycolipids (lactosyl PE, glucosyl-cerebroside). Here we have worked on supported lipid bilayers (SLBs) of saturated phosphatidylcholines, dilauroylphosphatidylcholine (DLPC) and distearoylphosphatidylcholine (DSPC), since we can control domain size and leaflet distribution.30,31 The DLPC/ DSPC phase diagram32,33 presents phase coexistence between a liquid crystalline phase and a gel phase, with a main phase transition temperature for pure DLPC of 268 K and for pure DSPC of 328 K.32 At room temperature in the hydrated state gel phase DSPC-rich domains are surrounded by a bulk liquid crystalline DLPC-rich phase. The large length difference of six carbons between DSPC and DLPC acyl tails is the cause of a large hydrophobic mismatch free energy penalty and may be the reason for a variety of observed DSPC leaflet distributions in DLPC SLBs.34,35 The distributions of gel phase DSPC lipids on the bilayer leaflets range from a symmetric distribution where DSPC domains in one leaflet are in registry with DSPC domains in the opposing leaflet, an asymmetric distribution where DSPC (26) Gousset, K.; Wolkers, W. F.; Tsvetkova, N. M.; Oliver, A. E.; Field, C. L.; Walker, N. J.; Crowe, J. H.; Tablin, F. J. Cell. Physiol. 2002, 190, 117–28. (27) Simons, K.; Ikonen, E. Science 2000, 290, 1721–6. (28) Hancock, J. F. Nat. ReV. Mol. Cell Biol. 2006, 7, 456–462. (29) Simons, K.; Ikonen, E. Nature 1997, 387, 569–72. (30) Lin, W. C.; Blanchette, C. D.; Ratto, T. V.; Longo, M. L. Biophys. J. 2006, 90, 228–37. (31) Blanchette, C. D.; Lin, W. C.; Ratto, T. V.; Longo, M. L. Biophys. J. 2006, 90, 4466–4478. (32) Mabrey, S.; Sturtevant, J. M. Proc. Natl. Acad. Sci. U.S.A. 1976, 73, 3862–6. (33) Bennun, S. V.; Longo, M.; Faller, R. J. Phys. Chem. B 2007, 111, 9504– 9512. (34) Bennun, S. V.; Longo, M. L.; Faller, R. Langmuir 2007, 23, 12465– 12468. (35) Lin, W. C.; Blanchette, C. D.; Ratto, T. V.; Longo, M. L. Biophys. J. 2006, 90, 228–237.
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domains are in only one leaflet and, and a mixed symmetry distribution where symmetric and asymmetric distributions coexist in the same domains.34,35 In SLBs at room temperature DSPC was found to be slightly soluble in DLPC (19:1 or higher, DLPC/ DSPC mol ratio)36 while DLPC presented a higher solubility in DSPC domains in SLB (1: 9 or greater, DLPC/DSPC mol ratio)36 or 15% solubility of DLPC in DSPC at 273 K in lipid vesicles.32 The advantage of DLPC/DSPC supported lipid bilayers as models for biological cell membranes is that they facilitate specific surface studies on well characterized lipid domain distributions. They have a reduced number of degrees of freedom with respect to biological cell membranes allowing a clear identification of the inherent mechanisms of drying/rehydration on lipid domains as models of cell membrane microdomains. Here we identify those mechanisms in a DLPC/DSPC system by correlating atomic force microscopy (AFM), fluorescence microscopy (FM), and fluorescence recovery after photobleaching (FRAP) data with the aim of a deeper understanding of desiccation mechanisms at ambient conditions.2,21,37,38 Plasma membranes have an asymmetric lipid distribution39 and therefore, we compare drying/ rehydration-induced lipid redistribution in asymmetric and symmetric DLPC/DSPC SLBs. As a comparison, the relative lack of redistribution in the presence of trehalose is demonstrated.
2. Materials and Methods 2.1. Materials. R,R-Trehalose dehydrate from Saccharomyces cereVisiae (98.5%) HPAE (high performance anion exchange chromatography) was purchased from Sigma-Aldrich (St. Louis, MO). 1.2-dilauroy-sn-glycero-3-phosphocholine (DLPC), 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC), and 1-oleoy1-2-[-6[(7nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl]-sn-Glycero-3-phosphatidylcholine (NBD-PC) were purchased in chloroform from Avanti Polar lipids (Alabaster, AL) and used without further purification. Water used in these experiments was purified in a Barnstead Nanopure System (Barnstead Thermolyne, Dubuque, IA) and presents a resistivity of 17.9 MΩ and pH of 5.5. 2.2. Supported Lipid Bilayer Preparation with and without Trehalose. Giant multilamellar vesicles (GMVs) were prepared by drying a lipid suspension of 64 mol % DLPC, 35 mol % DSPC and 1 mol % of fluorescence probe NBD-PC under a slow stream of nitrogen. The dried lipid film was resuspended with nanopure water or trehalose solution of 0.4 mg/mL for bilayers with trehalose. In both cases the final lipid concentration was 0.25 mg/mL. The lipid suspension with and without trehalose was placed for 10 min in a water bath at 70 °C and agitated twice for 10 s each time. The GMVs suspension was transferred to plastic test tubes to obtain small unilamellar vesicles (SUVs) using a tip sonicator (Branson sonifier, model 250, Branson Ultrasonics, Danbuy, CT) at power level 3 for 30 s, followed by power level 2 for 30 s. The plastic test tubes containing the SUVs were placed in a hot water bath (70 °C) for 5 min, after that 190 µL of the heated suspension of SUVs (70 °C) was deposited onto a heated mica surface (70 °C) glued to a metal puck, this technique results in asymmetric DSPC domains. For formation of symmetric domains the heated suspension of SUVs (70 °C) was deposited onto a room temperature mica surface (25 °C), the SLB was submerged into hot water at 70 °C. For both techniques the mica disk was then slowly cooled to 25 °C at a cooling rate of 5 °C/h in a temperature controlled oven (programmable digital chilling incubator model IN35, Torrey Pines Scientific, Inc.). After cooling, excess vesicles were removed by leaving the bilayers under water or trehalose solution for 5 h and then oscillating the bilayers under water by hand. The trehalose concentration used in this work for (36) Kraft, M. L.; Weber, P. K.; Longo, M. L.; Hutcheon, I. D.; Boxer, S. G. Science 2006, 313, 1948–1951. (37) Goyal, K.; Walton, L. J.; Browne, J. A.; Burnell, A. M.; Tunnacliffe, A. Integr. Comp. Biol. 2005, 45, 702–709. (38) Potts, M.; Slaughter, S. M.; Hunneke, F. U.; Garst, J. F.; Helm, R. F. Integr. Comp. Biol. 2005, 45, 800–809. (39) Devaux, P. F.; Morris, R. Traffic 2004, 5, 241–246.
Drying DLPC/DSPC Supported Lipid Bilayers preparation of supported lipid bilayers (SLBs), seems to have a negligible effect on the domain microstructure, apart from generating more adhered vesicles than SLBs prepared in absence of trehalose. The amount of vesicles increases with trehalose concentration to a point that vesicles cover the bilayer and make indistinguishable domains from the surrounding lipid phase. 2.3. Drying and Rehydration of Supported Lipid Bilayers. Drying was performed after FM, FRAP, and AFM imaging of hydrated supported lipid bilayers (SLBs) were used to characterize the domain microstructure. The water/trehalose solution drop on the top of the bilayer was minimized at the smallest possible size by carefully pipeting small amounts of water without disrupting the bilayer. The samples with and without trehalose solution were dried under a 22 mmHg gauge vacuum pressure for 48 h. Our goal was a minimization of non inherent drying effects on the bilayer; such as the ones caused by convective flux of gas streams that are not part of drying it self. After characterization of dried bilayers by FM and AFM imaging, the samples were rehydrated by adding nanopure water to non trehalose bilayers and 0.4 mg/mL trehalose solution to the bilayers prepared with trehalose. The rehydrated supported lipid bilayers (SLBs) were allowed to incubate for at least 30 min, before further FM, FRAP and AFM imaging. 2.4. Atomic Force Microscopy (AFM) Imaging. Contact mode topographic images of supported lipid bilayers were obtained with a Veeco Dimension 3100 Scanning Probe Microscope with Hybrid XYZ scanner (Santa Barbara, CA). Sharpened, coated silicon AFM cantilevers (Model MSCT-AUHW, Veeco, Santa Barbara, CA) with nominal spring constants between 0.01 and 0.05 N/m used for all scans. To minimize the force applied to the surface, the scanning set point was decreased until the tip left the surface and subsequently slightly increased until just regaining contact. To maintain the force at the lowest possible value, adjustment of the set point was required a few times during scanning. The scan rates ranged from 0.1 to 1 Hz, with set points larger than 0.2 V. Nanoscope V6 commercial AFM software package (Veeco, Santa Barbara, CA) was used to analyze AFM images heights and roughness, while domain surface coverage was analyzed with Image Tool Software version 3.0 (University of Texas Health center, San Antonio, TX). For imaging, samples were placed in Petri dishes containing either purified water or 0.4 mg/mL trehalose solution. Dried supported bilayer imaging was performed in empty Petri dishes. To obtain AFM scans of the rehydrated bilayer corresponding to the same place or region of the dried bilayer, water or 0.4 mg/mL trehalose solution was added to the Petri dish containing the dried supported lipid bilayer, always keeping the cantilever and bilayer in the same position and allowing a few minutes to stabilize the horizontal and vertical deflection fluctuation. 2.5. Fluorescence Microscopy (FM) and Fluorescent Recovery after Photobleaching (FRAP). Fluorescent microscopy and fluorescent recovery after photobleaching experiments were carried out on a Nikon Eclipse 400 fluorescence microscope (Nikon, Melville NY) equipped with a fluorescent filter cube (EF-4 FITC HYQ, Nikon) that matches the excitation emission spectrum of NBD-PC and is coupled with a 100-W xenon lamp as the fluorescence source. A 60× water immersion lens was used for fully hydrated and rehydrated state bilayers, while a 50× nonimmersion lens was used for dried bilayers. Images were captured with either a high resolution OrcaER Digital camera (Hamamatsu, Japan) or a Cascade II Digital camera (Photometrics, Tucson AZ). All FRAP measurements were done using a bleached spot of 30 µm, with the spot area free of gel phase domains to ensure a Gaussian fluorescent intensity profile. After bleaching, the lamp output was attenuated with neutral density filters. The FRAP curves were obtained by following the recovery of the bleached spot over time by fluorescence imaging for hydrated and rehydrated bilayers. The temporal changes of fluorescence intensity of the bleached spot was determined and normalized with respect to the maximum intensity at each time frame using Simple PCI Imaging Software version 6 (Compix Inc., Cranberry Township, PA). Recovery curves were fit to the fluorescence recovery in fractional form (eq 1) assuming FK(∞) ) F- as considered in ref
Langmuir, Vol. 24, No. 18, 2008 10373 40 with the Axelrod’s fluorescence recovery solution41 (eq 2) for Gaussian intensity profile beam assuming only lateral diffusion as a contributor to the recovery.
fK(t) ) [FK(t) - FK(0)] ⁄ [F- - FK(0)]
(1)
∞
FK(t) ) (qP0C0 ⁄ A
∑ [(-K)n ⁄ n!][1 + n(1 + 8Dt ⁄ w2)]-1)
n)0
(2) FK(0) ) (qP0C0 ⁄ A)K-1(1 - e-k)
(3)
F- ) (qP0C0 ⁄ A)
(4)
MATHEMATICA 5.0 (Wolfram Research, Inc., Champaign, IL (2003)) was used to obtain the nonlinear correlation parameters for K and D with K defined as the amount of bleaching, D the lateral diffusion coefficient, w the Gaussian beam radius, q the product of quantum efficiencies of light absorption, emission, and detection, A the attenuation factor of the beam during observation of recovery, P0 the total light power and C0 the initial fluorophore concentration.
3. Results DLPC/DSPC supported lipid bilayers were prepared containing large (>5 µm) asymmetric DSPC-rich domains, where DSPC domains appeared in only the leaflet distal to the substrate,30 or large symmetric DSPC-rich domains, where both leaflets were in registry. Briefly, differences in the thermal history during SLB formation allowed us to direct the symmetry of the SLB, specifically by including an additional heating step to form symmetric SLBs as we did in previous work.30 These large welldelineated domains were imaged in the hydrated state by both FM and AFM, accessing micron and submicron-scale structural details, and then exposed to drying and rehydration in the presence and absence of trehalose accompanied by further characterization by FM and AFM in the dried and rehydrated states. We also present lateral diffusion FRAP analysis on hydrated and rehydrated (after drying) SLBs containing asymmetric domains. The AFM and FM imaging was performed on at least 20 different preparations corresponding to at least 60 samples. In the discussion, we relate the major structural rearrangements that we observe to driving forces that initiate changes such as area contraction/expansion and air-water surface tension induced stress and processes that are then brought about in the membrane such as lipid flip-flop, lipid exchange, and membrane collapse. We relate our observations to the literature and discuss their relevance and implications to anhydrobiotic preservation as well as their possible biological significance. 3.1. Micrometer Scale Morphology and Lateral Diffusion in Large Asymmetric Domains: Hydrated, Dried, and Rehydrated States. Fluorescence imaging shows micrometerscale resolution of hydrated DLPC/DSPC asymmetric bilayers in the absence of trehalose (Figure 1 A) and presence of trehalose (Figure 1 B). The DSPC-rich domain shapes are rounded indicative of slow domain growth.42 No differences were noted with respect to domain shapes in lipid membranes hydrated without trehalose or with trehalose. The SLBs were then dried and rehydrated. Surprisingly, in the absence of trehalose upon rehydration, DSPC-rich domains were still observable and round, yet the majority of the samples displayed changes. Uniformly, in the absence of trehalose upon rehydration there was a lightening of the domains in comparison to the hydrated (40) Soumpasis, D. M. Biophys. J. 1983, 41, 95–97. (41) Axelrod, D.; Koppel, D. E.; Schlessinger, J.; Elson, E.; Webb, W. W. Biophys. J. 1976, 16, 1055–1069. (42) Blanchette, C. D.; Orme, C. A.; Ratto, T. V.; Longo, M. L. Langmuir 2008, 24, 1219–1224.
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Figure 1. FM images showing DLPC/DSPC asymmetric supported lipid bilayers hydrated without (A) and with (B) trehalose, then dried and rehydrated without (C-F) and with trehalose (G). Arrows point to light mottling within domains.
Figure 2. FM images showing DLPC/DSPC asymmetric SLBs. (Left) Dried DLPC/DSPC bilayer in absence of trehalose. (Right) DLPC/DSPC dried bilayer in presence of trehalose.
samples; however the details of this lightening differed from sample to sample and region to region. This included, a general loss of contrast between the domains and surrounding (Figure 1C), lighter mottled regions on the domains (Figure 1D see arrows), domains that appear to have dissolved almost completely (Figure 1E), and domains with dark rims, light mottling throughout, and attached bright aggregates, surrounded by regions containing dark mottling (dark spots) (Figure 1F). Interestingly, large-scale delamination of the mica substrate, as observed in another study using POPC SLBs on glass substrates,25 was never observed by us. The NBD-PC probe partitions with the DLPC component,43 therefore our results may be indicative of a change in DLPC distribution, with DLPC somehow moving into the domains. This would also imply that DSPC may be moving out of the domains, and there is evidence of this in some samples in the form of dark mottling, see Figure 1F, between domains. In order to determine if lipid redistribution, was occurring we observed this system using AFM, described in the section below. In contrast, in the presence of trehalose, upon rehydration we observed a similar morphology and contrast between domains and surroundings in comparison to hydrated bilayers (Figure 1G). We imaged by FM, in the dry state, DLPC/DSPC SLBs without trehalose and others with trehalose. Without trehalose, similarities of the dried and rehydrated samples were noted, indicating that some aspect of the rearrangements occurred during the drying process. As shown in Figure 2, left, we observed that the domains and surroundings were heterogeneous in fluorescence intensity with mottled appearance and darker rims around many of the domains. In contrast, dry SLBs in the presence of trehalose, shown in Figure 2, right, very closely resembled the hydrated (43) Mesquita, R. M. R. S.; Melo, E.; Thompson, T. E.; Vaz, W. L. C. Biophys. J. 2000, 78, 3019–3025.
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Figure 3. Representative FRAP experiments performed on DLPC/DSPC asymmetric SLBs containing 1% of NBD-PC Fluorescent Probe. Experiments were done before drying (dashed line) and after rehydration in presence of trehalose (solid line). Both curves show similar fast recovery trends. Insets of FM images of 133 µm × 133 µm for both (A, before drying and B, after rehydration) show on the left the dark bleaching spot and on the right the extent of recovery after 2 min.
Figure 4. Quantitative traces of fluorescent intensity across the bleach spot in x and y directions at time 0 min (left) corresponding to the bleach spot after exposure to the bleaching light and 25 min later (right). The intensity scales on the left of each image indicate an overall loss of fluorescence intensity from time 0 to 25 min.
bilayers except for evidence of the drying pattern of trehalose where thick layers of trehalose have deposited and there is a dark network-type appearance. Note that the network pattern is superimposed over both the domains and the interdomain regions as would be expected for a dried coating. Mobility of the lipids surrounding the domains was assessed using spot fluorescence recovery after photobleaching (FRAP). Hydrated bilayers in the presence or absence of trehalose recovered quickly (Figure 3A). Fitting of the FRAP data (example see Figure 3) resulted in lipid diffusion coefficients (D) of 5.2 ( 1.2 µm2/s for hydrated SLBs with and without trehalose. The large domains, on the order of micrometers, apparently did not affect to large extent D values compared with pure DLPC bilayers.44 Rehydrated bilayers in the presence of trehalose, also recovered quickly (Figure 3B) displaying not statistically different D values (4.6 ( 1.3) µm2/s in comparison to hydrated bilayers. Hydrated bilayers with and without trehalose presented a recovery of 97 ( 1% at longer times. While for bilayers rehydrated with trehalose, a recovery of 95 ( 1% was obtained in agreement with a 97% recovery reported for POPC bilayers rehydrated with trehalose.25 Qualitatively, for the rehydrated samples containing trehalose, there was rapid disappearance of a bleach spot between 2 and 7 min. In the absence of trehalose quantification was confounded by the loss of fluorescence intensity in the regions surrounding the domains that accompanied an overall lost of contrast between domains and surroundings. However, when we were successful in performing spot photobleaching, recovery times were extremely slow as demonstrated in Figure 4 that had not completely recovered in 25 min. These (44) Ratto, T. V.; Longo, M. L. Biophys. J. 2002, 83, 3380–3392.
Drying DLPC/DSPC Supported Lipid Bilayers
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Figure 5. Atomic force microscopy images, sections, and DSPC domain height histograms from 100 µm × 100 µm scans of DLPC/DSPC asymmetric SLBs. (A-C) Hydrated with trehalose. (D-F) Rehydrated with trehalose. (G-I) Rehydrated without trehalose. Sections B, E, and H correspond to domains in white dotted circles for respective images A, D, and G. While section I corresponds to enlargement of image G. Arrows in sections and corresponding images indicate high features >3 nm in height. Bar at right corresponds to image G.
images demonstrate that very limited exchange appeared to be occurring with the region surrounding the illuminated viewing area and general loss of fluorescence, from observation illumination, prevented accurate determination of D values in all cases. Evidence that this phenomena may be caused by compartmentalization/obstruction, rather than gelling of the DLPC, is present in the photobleaching spot in Figure 4 as a sharp edge on the upper portion of the bleach spot. Interestingly, the bleached region did not spread across this edge, indicating that some kind of a continuous barrier existed across the sample at that location. 3.2. Nanometer-Scale Morphology in Lipid Membranes with Large Asymmetric Domains: Hydrated, Dried, and Rehydrated States. By AFM imaging, we found that asymmetric domains in DLPC/DSPC SLBs hydrated in the presence and absence of trehalose project ∼1nm from the DLPC fluid-phase
(Figure 5A-C), in agreement with a reported height difference of 1.1 nm between asymmetric (domain in only one leaflet) DSPCrich domains and DLPC-rich fluid phase in SLBs. 35 The bilayers appeared unchanged at least over a period of 6 h. Sometimes small trapped fluid DLPC-rich subdomains >200 nm were identified in the large asymmetric DSPC-rich domains (enlarged area of Figure 5A). Bilayers rehydrated in presence of trehalose (Figure 5D) have a similar appearance to the original fresh hydrated bilayers (Figure 5A) in agreement with the FM results. The height difference between the domain phase and surrounding phase for the rehydrated bilayers was ∼1 nm (Figure 5E and F) corresponding to an asymmetric domain distribution. By AFM, we were able to observe detailed changes in the 2D and 3D distribution of DLPC and DSPC of rehydrated SLBs in the absence of trehalose, as illustrated in Figure 5G. In agreement
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Figure 6. AFM height images and sections in DLPC/DSPC asymmetric SLBs upon rehydration without trehalose. (A-B) High density of DLPC subdomains; (C-D) highly fragmented domains. Dotted circles represent high features of height >3nm. Bar at right corresponds to image C.
with our FM observations, the details of these changes varied from sample to sample, yet always a large area fraction within the domains displayed a height corresponding to a DLPC bilayer. This included domains that remained relatively intact yet contained a large area fraction of DLPC subdomains (>25%) as shown in Figure 6A-B in comparison to typical area fractions of DLPC subdomains in hydrated SLBs of 0-7% as determined by AFM. Note that sections around the rim of the domain shown in this image remain relatively subdomain-free. Other domains appear to have dissolved and become a collection of smaller 2.5 nm per area range from 8 to 23 features per 100 µm while for rehydrated bilayers of DLPC/DSPC without trehalose the density of features per area is 238-912 features per 100 µm. The presence of deep fissures generally in the radial direction across the domains (radial fissures) as seen in figure 5G, indicates that lipid is being shed to the surrounding fluid phase, most likely as a result of the formation of the high features. Figure 5G shows a combination of these features (subdomains, radial fissures, domain fragmentation, and high features). We imaged by AFM, in the dry and rehydrated state, the same area of a DLPC/DSPC SLB without trehalose as illustrated in Figure 7. For the trehalose sample, a thick trehalose layer (45) Takamoto, D. Y.; Lipp, M. M.; von Nahmen, A.; Lee, K. Y. C.; Waring, A. J.; Zasadzinski, J. A. Biophys. J. 2001, 81, 153–169.
prevented any useful information being obtained by AFM imaging. The most noticeable feature of the dried sample without trehalose is that features of similar height in comparison to the larger rounded domains appear throughout the regions between the domains and they have a crystalline-like facetted appearance (see bottom arrow in Figure 7A). In addition, the domains themselves have a high density of “low spots” of width less than 500 nm and high features (see top arrow in Figure 7A). By section analysis, it appears that both defects (depth ∼5 nm) and DLPC subdomains exist within the dried domains. Interestingly, the rims of the domains appear more crystalline and show an absence of ”low spots”. In the rehydrated state, the domain area shrinks by 20-46% as illustrated by the domain circled in both Figure 7A (dried) and C (rehydrated). In the circled domain, a radial fissure of depth between 1.5 and ∼5 nm has appeared across the center. There is evidence in the other domains that these are forming across crystalline facets present in the dried domains. High features are present in the rehydrated samples, often in the same location as they were observed in the dried samples, but generally wider and higher (top arrows Figure 7A and C). It is difficult to say exactly how much of the features that appeared between the domains in the dried sample still exist between domains in the rehydrated sample. This is because domains that are highly fragmented tend to be easily compressed by the AFM tip. By using a lower force the features appeared wider and more numerous (image not shown). However, it is clear that some of these features still exist (for example, bottom arrows in Figure 7A and C) and that high features are often associated with them. A smaller scan area image of the rehydrated sample reveals elongated defects of ∼200 nm in width in the DLPC-rich region (left blue cross Figure 7E and corresponding section in Figure 7F). We have quantified the presence of DLPC subdomains in the DSPC domains and vise versa, by comparisons of roughness (Ra). We use Ra ) 1/N∑Nj ) 1 |Zj|, defined as the arithmetic average of the absolute values of the surface height deviations measured
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Figure 7. AFM images and sections in DLPC/DSPC asymmetric SLBs without trehalose after drying (A-B) and rehydration (C-F). Bottom arrow in A shows faceted feature of similar height compared to large domain (e.g., circled domain) that partially still exists in C. Top arrow in A shows a high feature that is associated with a larger and higher feature in C. Left blue cross in E intersects an elongated defects of width ∼200 nm corresponding to the left dashed line in F.
from the mean plane (Nanoscope Software manual version 6.13, Veeco Instruments Inc., Santa Barbara, CA). Roughness of (1.05 ( 0.22) nm was obtained within the domains rehydrated without trehalose. While the surrounding phase yields a roughness of (0.29 ( 0.03) nm. Larger height features (>2.5 nm) were avoided for roughness calculations. These values are larger compared to the roughness for hydrated bilayers, that yielded a roughness of (0.134 ( 0.009) nm within the domains and a roughness of (0.142 ( 0.006) nm for the surrounding region and larger compared to trehalose rehydrated bilayers with roughness of (0.217 ( 0.007) nm for both the corresponding domain and surrounding phase. For the SLBs dried without trehalose, Ra ) (1.08 ( 0.16) nm was obtained within the domains while the surrounding phase yielded Ra ) (0.89 ( 0.25) nm; in this measurement again larger height features >2.5 nm were avoided. 3.3. Nanometer-Scale Morphology in Lipid Membranes with Large Symmetric Domains: Hydrated, Dried, and Rehydrated States. In the hydrated state in presence and absence of trehalose the difference in height between the DSPC-rich domains as shown in Figure 8A and the surrounding fluid phase is ∼2 nm (Figure 8 B) in agreement with reported values of 1.8 nm for small symmetric DSPC domains in SLBs. 35 We also observe high features of height usually ∼7 nm, a region of double bilayer probably formed during the area expansion that accompanies the heating step required to form symmetric domains. After drying, rehydrated bilayers with trehalose (Figure 8 C) are similar in appearance to hydrated bilayers as shown in Figure 8A with the difference in height between the DSPC-rich domains and the surrounding fluid phase also approximately 2 nm (Figure 8 D).
After drying followed by rehydration, in the absence of trehalose we observed major changes in the 2D and 3D structure of the DLPC/DSPC symmetric SLBs as demonstrated in Figure 8E, with similarities to the asymmetric case. However the lipid symmetry and probably the presence of a solid support also appeared to be playing an important role here. Notably, one of the DSPC-rich domain leaflets in all of the domains did not undergo any significant replacement with DLPC. However, the opposing DSPC-rich leaflet underwent various degrees of replacement/exchange with DLPC, varying from near 0%, evidenced by symmetric domains that appeared nearly intact with domain heights throughout of ∼2 nm (Figure 8 E bottom arrow), to ∼50% (Figure 8E top arrow). Regions in which replacement/exchange had occurred had heights ∼ 1 nm compared to much of the surrounding region as shown in the section of Figure 9A (Figure 9B). It was quite evident from domains with more than 25% exchange in one of the leaflets that the remaining DSPC in that leaflet was developing a fractal shape as illustrated in the enlargement of Figure 9A, characteristic of a rapid surface exchange process that is diffusion limited.42 In addition, often regions in the domains did not show evidence of exchange with DLPC similar to our observation at the domain rims of asymmetric domains, perhaps suggesting regions of higher long-range crystalline order. Features of ∼1 and 2 nm height appeared throughout the regions between the rounded domains sometimes with a fractal-like appearance as if they had formed rapidly as illustrated in the enlargement of Figure 9A. High features were more commonly seen than in the hydrated samples, as well as round 100 nm-scale pinhole defects in the DLPC-rich regions (Figure 9 small inset, arrow points to three aligned pinholes).
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Figure 8. AFM images and sections in DLPC/DSPC symmetric SLBs. (A-B) Hydrated bilayers. (C-D) Rehydrated bilayers with trehalose. (E-F) Rehydrated bilayers in absence of trehalose. Left arrow in E points to a domain that has undergone significant lipid exchange in one of the leaflets and right arrow points to a domain that is relatively intact. Section images B, D, and F correspond to domains in white dotted circles for respective figures A, C, and E.
Figure 9. AFM image (A) and section (B) of rehydrated symmetric DLPC/DSPC SLB without trehalose showing that DSPC has been replaced by DLPC in large areas while in other domains much less exchange has occurred. Top left corner of enlargement shows a large surface fissure surrounding the domain and extending 5 nm in depth according to the section analysis. Small inset shows 100 nm-scale pinhole defects in the DLPC-rich region with arrow pointing to three such defects that are in a line going away from the arrow point. Dotted circles show features higher than 5 nm.
Occasionally, large surface cracks or fissures (3-4 nm in depth) could be observed surrounding domains and extending to the mica substrate as can be seen in the enlargement of Figure 9A. We imaged by AFM, in the dry and rehydrated state the same area of a DLPC/DSPC SLB without trehalose as illustrated in Figure 10. The desiccation changed the height between the domain phase and surrounding phase, which decreased from 2 nm in the hydrated state to approximately 1.2-1.5 nm in the dry state, suggesting probably that DLPC has transitioned from fluid phase to gel phase. The height change, considering both leaflets, is in agreement with reported increases from 0.8 Å per acyl chain carbon atom in the fluid phase to 1.1 Å per acyl chain carbon
atom in the gel phase.46,47 For these bilayers (Figure 10A-B), there was much less evidence of changes in 2D than had been previously observed for dried asymmetric SLBs (Figure 7A). For hydrated bilayers, domain roughness was (0.203 ( 0.008) nm and for the surrounding phase it was (0.186 ( 0.021) nm. For dried bilayers the roughness of the domain region was (0.142 ( 0.006) nm and for the surrounding phase (0.205 ( 0.007) nm. Upon rehydration, as seen in Figure 10C, the sample displayed the same types of changes that are described above. Sections (Figure 10B, D) show a domain that remains primarily as a (46) Lewis, B. A.; Engelman, D. M. J. Mol. Biol. 1983, 166, 211–217. (47) Dumas, F.; Lebrun, M. C.; Tocanne, J. F. FEBS Lett. 1999, 458, 271–277.
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Figure 10. AFM images and sections in DLPC/DSPC symmetric SLBs. (A) Dried bilayer. (C) Rehydrated bilayers in absence of trehalose. Enlargements are deflection AFM images. Upper arrows show a domain that remains largely symmetric upon rehydration and lower arrows show a domain that has undergone significant DLPC/DSPC exchange. (B and D) The section analyses for the domains in the enclosed square in figures A and C.
coupled DSPC-rich domain of height 2 nm (upper arrow), while the neighboring domain (lower arrow) has undergone quite substantial exchange and displays two heights (1 nm, 2 nm). Overall, the movement of DLPC into the leaflets of some of the domains and accompanying outward movement of DSPC, and other changes appear to occur primarily as a result of rehydration in the symmetric DLPC/DSPC SLB system. Fluorescent microscopy results after rehydration displayed domains consistent with partial lipid exchange in one leaflet of the domains, i.e., two fluorescence intensities in the domains. In addition, the regions around the domains contained more pronounced dark mottling than we observed for symmetric samples, that could be a result of more DSPC in the regions between the domains, consistent with our AFM results. The lack of contrast resulted in fluorescence images of poor quality, therefore, we reported only the more informative AFM results above.
4. Discussion In this study, we have achieved a detailed view of the 2D lipid exchange and 3D collapse that occurred as a result of drying and rehydration of a mixed SLB containing DLPC/DSPC by FM, FRAP, and AFM. Interestingly, lipid exchange and collapse occurred during both drying and rehydration for an asymmetric bilayer, but mostly during rehydration for a symmetric SLB. This work suggests that during drying and dehydration a number of interconnected and simultaneous processes occur that we will discuss here and put into context with related studies. 4.1. Domain Contraction and DLPC/DSPC Exchange during Drying. For asymmetric domains, our images suggest that a major rearrangement occurred during the drying. Specifically, local contraction in the area/molecule of DSPC in the domains took place as it entered the dried state creating highly condensed regions. The most expansive of these regions were
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often at the rims of the domain, suggesting either that those regions condensed first or that packing difference existed between the lipids near the rims and the domain center prior to drying. Alternatively, this ring-like pattern of ordered lipid may reflect a typical drying pattern where water flowing to a droplet perimeter transports material to the edge.48 This suggests that it may be possible for single water droplets to associate with individual domains during drying. From X-ray diffraction work,49 we can expect a contraction of at least 10% during this drying process. Free area within the domain was created from this shrinkage producing defects and also being filled by DLPC bilayer, evidenced by creation of subdomains. The subdomain DLPC may originate from the opposing leaflet through lipid flip-flop and from the surrounding DLPC region through connectivity (percolation) of the created free area to the surrounding DLPCrich region. Through these processes, the connectivity of DSPC in the domains was disrupted and more DSPC/DLPC interface in the asymmetric domains was created. In the dry state, the domains fingered out into the surrounding DLPC-rich region that is also undergoing condensation. Although the dry DLPC/ DSPC SLB was certainly better mixed than in the hydrated state, the observed heterogeneity would not indicate that the two components are fully mixed. Similarly, FTIR spectroscopy studies indicate that drying induces mixing of these same two components in small unilamellar vesicles.22 However, the components are reported to become fully mixed as evidenced by DSPCd-70 and DLPC presenting similar transition temperatures and the lack of two phosphate asymmetric stretch bands. Differences in the size and distance between domains in our study (microscopic) compared to the other study may be responsible since the length scales for diffusive mixing would be orders of magnitude larger here. As drying completed, local enriched regions of dried crystalline DLPC and DSPC developed in both the domain region and surrounding region. In contrast, symmetric bilayers showed less evidence of rearrangements induced by drying. It is possible that the drying process is gentle enough for the proximal domain leaflet to “anchor” the entire domain through DSPC-substrate interactions. In the asymmetric system, we expect that these DSPC-substrate interactions were not present because DSPC domains form in the distal leaflet.30 Additionally, the general cohesiveness of opposing domain leaflets in registry may prevent significant formation of free area large enough for DLPC to penetrate into the domains. Lipid flip-flop was also not a possible mechanism to fill, with DLPC, any free area that might have formed in symmetric domains. Therefore, significantly less created DLPC/ DSPC interface appears to have prevented drying induced rearrangements/mixing in symmetric SLBs. It is difficult to make a direct comparison of the symmetric domain results with the previous FTIR results22 because the domains in SUVs are asymmetrically distributed at room temperature (DSPC-enriched on the interior)30 and were not interacting significantly with a substrate. 4.2. Initiation of Collapse Structures during Drying. The loss of water generates surface-tension and area contraction induced stresses in the bilayer usually accompanied by the observation of cracking and fissure.11,25 Interestingly, in the case of drying asymmetric DLPC/DSPC bilayers, these stresses seem to have resulted in the appearance of high features, associated with the appearance of DLPC/DSPC interfaces at subdomains and defects, that grew after rehydration. This is in agreement (48) Smalyukh, I. I.; Zribi, O. V.; Butler, J. C.; Lavrentovich, O. D.; Wong1, G. C. L. Phys. ReV. Lett. 2006, 96, 177801. (49) Koenig, B. W.; Strey, H. H.; Gawrisch, K. Biophys. J. 1997, 73, 1954– 1966.
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with recent work in monolayers showing that collapse structures tend to nucleate at similar interfaces, due to favorable local curvature.50 Other work involving drying of POPC SLBs supported by silica substrates shows the appearance of similar collapse-like structures (vesicles and folds) by FM.25 In the dry state, it should also be expected that some regions will form lipid trilayers to expose a hydrophobic face to the air although this could not be confirmed due to the extreme heterogeneity in height of the dried samples. There was no significant introduction of collapse structures in the symmetric system through drying, consistent with generally less mixing. 4.3. Mixing,Collapse,DomainContractionduringRehydration. Major rearrangements accompanied rehydration in both the asymmetric and symmetric system. The severity of these rearrangements indicates that rehydration by addition of water may be a more violent process in comparison to slow drying under vacuum. It appears that the formation of DLPC/DSPC interface in the drying process was the major factor for drying/ rehydration-induced damage in the asymmetric SLB. Our work suggests that lipid removal by collapse occurred at the DSPCDLPC interfaces and that the collapse structures nucleated during drying. Because of the creation of DLPC/DSPC interface and stresses created in the drying and rehydration process, we believe that the rehydrated bilayer is dense with submicron-scale pinhole defects and line defects. This is supported by the presence of high features, linear defects, low diffusion coefficients, and imaging of defects in the DLPC-rich phase of the similar symmetric system. However, higher resolution AFM studies will be necessary to confirm this. Close similarities exist with recent work in the adoption of fragmented type domain appearance, although those investigators observed substantially more removal of bilayer from the mica substrate, including as much as 50% delamination in one area.24 Complete delamination has been reported for POPC bilayers supported on silica substrates in which large aggregates are readily visible by fluorescence in the dried state.25 The relatively low delamination that we observe may be related to the absence of salt compared to the previous studies that both used 150 mM NaCl combined with buffers, and differences in substrates, lipids, drying methods, and time between rehydration and imaging. In the case of bilayers containing symmetric DSPC-enriched domains, the changes most closely resemble those observed in dried asymmetric bilayers. This could be expected because the initiation of rehydration must mirror a late state of dehydration. The proximal domain leaflet appears to be “anchored” to the substrate; therefore most of the changes occur in the distal leaflet or at the domain interface. Such anchoring would not be present in the case of a free bilayer, but it is possible that anchoring would be relevant to cell membranes because of the presence of cytoskeletal attachments. Formation and growth of observed collapse structures (high features) at the domain interfaces could and may initiate the reorganization process. Indeed, high features are observed as well as cracks, some surrounding the domains, perhaps eventually halting the DLPC/DSPC exchange process for those domains. 4.4. Comparison to Bilayers Dried/Rehydrated in Presence of Trehalose. Supported bilayers containing asymmetric and symmetric DSPC-rich domains dried in the presence of trehalose displayed minimal traces of damage. A doubling of the roughness (vs 10× without trehalose) indicates that some minor version of the processes described above is occurring, perhaps driven by air-water surface tension driven stresses. Although we do not believe that these experiments answer questions about the (50) Gopal, A.; Lee, K. Y. C. J. Phys. Chem. B 2001, 105, 10348–10354.
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mechanism of the protective effect, e.g., glass formation vs water replacement hypothesis, the work provides snapshots of the 2D and 3D rearrangement processes that are avoided by the presence of trehalose. 4.5. Possible Biological Implications. Maintaining membrane functionality requires both intact structure and fluidity. Our results show evidence that without trehalose, condensation of lipid chains take place and collapse structures form in drying and/or rehydration, both processes that could initiate the observed DLPCDSPC exchange/mixing. These processes, avoided by the presence of trehalose, are most likely related to phase behavior and surfacetension induced bilayer stress respectively. Therefore, both the water replacement hypothesis and vitrification hypothesis are possible explanations for the action of trehalose. Here we show by FRAP experiments that membrane fluidity is severely disrupted by drying and rehydration, which we again can relate back to exchange/mixing causing severely obstructed diffusion possibly through defect formation/collapse at DLPC-DSPC interfaces. This logic again leads us to the conclusion that trehalose could prevent either the collapse or phase transformation initiating mixing by either major mechanistic hypotheses. Recently, it has been shown that DLPC giant unilamellar vesicles (GUVs) resisted cracking and leakage upon desiccation whereas DSPC GUVs underwent significant cracking and leakage (similar to mammalian cells51).19 The distinction in that study appeared to be that the DLPC GUVs did not adhere to a surface where the DSPC GUVs did, implying that the tension-free state of free bilayers could result in rapid closing of defects formed during drying rearrangements. Cell membranes by their cortical tensions and SLBs by their fixed areas, both can not rapidly recover to drying induced changes by decreasing the area of the membrane. Therefore, multicomponent SLBs with controllable lipid symmetry provide a useful model system for studying drying/ rehydration induced phenomena in the absence and presence of trehalose. Using such a system, we identify mixing of lipid species during drying or dehydration as a crucial step in the formation of defects such as pinholes and cracks by increasing nucleation sites at lipid boundaries for formation of aggregates. Previous studies suggest that leakage is driven by defects originating in a fluid phase passing through a gel phase transition upon drying/rehydration.52 The observations here suggest that (51) Steponkus, P. L.; Lynch, D. V.; Uemura, M. Philos. Trans. R. Soc. London, Ser. B 1990, 326, 571–583. (52) Crowe, J. H.; Crowe, L. M.; Oliver, A. E.; Tsvetkova, N.; Wolkers, W.; Tablin, F. Cryobiology 2001, 43, 89–105.
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much larger-scale membrane damage is possible through lipid exchange processes accompanying drying/rehydration. The curvature associated with the interface formed between dissimilar lipids may more effectively nucleate collapse sites in comparison to the interface between gel and fluid phase lipids of the same or similar species. This work also suggests that grain boundary defects may be a minor pathway for leakage in comparison to >100 nm-scale pinholes and larger fissures/cracks that accompany the formation of aggregates at interfaces of dissimilar lipids.
5. Conclusion In this study we followed structural and dynamic changes in DLPC/DSPC supported lipid bilayers under drying/rehydration by characterizing lipid mobility by fluorescence recovery after photobleaching and membrane microstructure by fluorescence microscopy and atomic force microscopy. This work suggests that drying and rehydration induce large-scale rearrangements through mixing/lipid exchange and formation of collapse aggregates at the interfaces between lipids of different phases/ lengths. Furthermore by use of asymmetric vs symmetric DSPCrich domains, we found that mixing in absence of trehalose can begin with the drying process or the rehydration process respectively, indicating that strong interleaflet or substrate-domain interactions can be effective in preventing some drying-induced damage. Different from previous work, we did not see largescale delamination of the substrate and we could follow all stages of the drying/rehydration process with both fluorescence microscopy and atomic force microscopy, giving us enough detail to propose a series of interconnected stages and propose a mechanism for membrane leakage. In the presence of trehalose, the lack of rearrangements point to the necessity of a drying agent for cell membrane preservation during dehydration. Acknowledgment. We thank Dr. Fern Tablin and Dr. John Crowe for collaborative discussions and Dr. Brian Higgins for his Mathematica notes. This investigation was supported by the Graduate Research and Education in Adaptive bio-Technology (GREAT) Training Program of the UC Systemwide Biotechnology Research and Education Program under Grant No. 2005244. We acknowledge funding by the NSF NIRT Program (CBET 0506602) and the NSF MRSEC Program CPIMA (NSF DMR 0213618). LA8016694
