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Dynamics of supported lipid bilayer deposition from vesicle suspensions
Journal of Colloid and Interface Science 348 (2010) 608–614
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Journal of Colloid and Interface Science www.elsevier.com/locate/jcis
Dynamics of supported lipid bilayer deposition from vesicle suspensions Seyda Bucak *, Chen Wang 1, Paul E. Laibinis 2, T. Alan Hatton Department of Chemical Engineering, Massachusetts Institute of Technology, Cambridge, MA 02139, USA
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Article history: Received 10 March 2010 Accepted 29 April 2010 Available online 4 May 2010 Keywords: Solid-supported membranes Liposomes Surface coverage Adsorption kinetics
a b s t r a c t The equilibrium and dynamic adsorption behavior of mixed lipid (DMPC and DMPG) vesicles on a hydrophilic silica bead substrate indicate that the adsorption is a two-step process. Vesicles initially adsorb and spread on the silica surface as lamellar bilayer discs, with a rate mediated by the charge–charge interactions between vesicles and the silica beads. Vesicle adsorption occurs more readily on the growing discs than on the bare silica surface, and consequently the adsorption rate increases as the discs grow. Eventually, the growth is arrested because of unfavorable charge interactions between the discs as they cover more of the surface, and the equilibrium condition of partial surface coverage is attained. Ó 2010 Elsevier Inc. All rights reserved.
1. Introduction Over the past two decades, special attention has been paid to vesicles made from both natural and synthetic phospholipids with the realization of their potential applications as drug delivery carriers and as model systems for the study of cell–cell and cell–membrane interactions in the immune system [1–4]. A number of problems were noted, however, when quantitative studies of the properties and behavior of the artificial lipid membrane systems were required. These include the stability of the prepared liposome dispersion over time due to vesicle fusion, the variable number of bilayers and water content in each compartment, and the change of phase structures under different conditions [5]. A well-defined lipid bilayer system is thus necessary to facilitate investigations on e.g., lipid–protein interactions. Lipid membranes supported on solid substrates can be prepared to meet these needs, with potential use also in more practical applications. The number of lipid layers in supported lipid membranes can be controlled by employing a suitable substrate and preparation method. Two types of membranes that are particularly interesting are the so-called supported lipid bilayer (SLB) and supported lipid monolayer (SLM). An SLB is formed on a hydrophilic surface such as silica, quartz, oxidized silicon, and glass consists of two leaflets of phospholipids [6]. An SLM can be assembled on silane-derivatized silica surfaces [6,7] or thiol-coated gold surfaces [8–12], or it can be
* Corresponding author. Present address: Yeditepe University, Department of Chemical Engineering, Kayisdagi, Istanbul, Turkey. E-mail address: [email protected] (S. Bucak). 1 Present address: Millipore Corporation, USA. 2 Present address: Vanderbilt University, Department of Chemical and Biomolecular Engineering, Nashville, TN 37235, USA. 0021-9797/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.jcis.2010.04.087
obtained by the covalent linking of chemically modified phospholipids with silica [13] or gold. Another type of lipid monolayer, also called immobilized artificial membrane (IAM), can be obtained by the direct covalent bonding of phospholipid analogs on silica particles [14,15]. Such lipid/membrane systems are geometrically well defined and are potentially suitable for the aforementioned quantitative studies. Supported lipid membranes have been used widely in several applications such as preparation of biosensors for environmental monitoring, drug development, medical diagnostics and industrial process control. Immobilized artificial membrane (IAM) chromatography has been used successfully to measure drug–membrane interactions and drug membrane permeability. In addition to IAM, immobilized liposome chromatography (ILC) was developed by Lundahl’s research group in Sweden to study drug–membrane interactions [16–20]. Free lipid vesicles have been combined with ultrafiltration or precipitation to separate protein mixtures [21– 24]. Bayerl and Bayerl coated macroporous silica with lipid bilayers and used these treated beads as the stationary phase for gel permeation and ion exchange chromatography [25]. Another conceptually novel chromatography process, dubbed phase transition chromatography (PTC), was also developed by Bayerl’s research group. Although SLB have been extensively used in several applications, the mechanism of vesicle adsorption onto flat (or near flat) surfaces is still under investigation. Lipid molecules in the supported bilayers can exchange with those in solution, just as in free vesicle suspensions, although this lipid transfer between bilayers on different substrates occurs at a much slower rate than that between free vesicles. It is inferred that lipid monomers dissociate from the surface lipid layers or vesicles and fuse with other bilayer patches. A surface may stabilize the adsorbed SUVs by increasing
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the activation energy barrier for lipid desorption from the supported membrane, while inhibition of collisions between SUVs and supported bilayers also reduces the lipid exchange rate [26]. Bayerl et al. observed a difference in the phase transition temperature of vesicle bilayers in aqueous media and solid supported bilayers [27], which they concluded was due to increased demixing of two lipids in the medium in the two-phase region with increasing curvature due to lowering of the lateral pressure. The formation of supported lipid bilayers on silica nanoparticles was observed within a few minutes of contact time by cryo-electron microscopy [28]. They have shown that bilayer formation was observed with negatively charged, neutral and positively charged lipids but not with highly negatively charged particles where rupture of vesicles did not take place. Zhadanov and Kasemo [29] suggested that the support-induced deformation of an adsorbed vesicle is further enhanced by the adsorption of vesicles in its vicinity. AFM studies have shown fusion of adsorbed vesicles [30]. Phospholipid bilayer formation at bare silica surface was also studied in situ with neutron reflection [31]. Although the first few minutes could not be captured (which is believed to be the adsorption of individual phospholipid vesicles on the bare silica surface), the next stage where growth of bilayer covered area by fusion of single spots was observed until a homogenous stable and fluid phospholipid bilayer covered the surface. Adsorption onto different surfaces [32], effect of hydrophobicity of the lipid tails on adsorption [33] and effect of presence of salt in adsorption of charged lipids [34] have also been studied. In has been found that in order to deposit onto solid, oppositely charged surfaces, charged bilayers also have certain requirements of bilayer fluidity [35]. DODAB vesicles are also found in the rigid gel state at room temperature [36] and closed DODAB vesicles are difficult to disrupt for bilayer deposition so that previously disrupted vesicles or bilayer fragments had to be used in order to optimize gel bilayer deposition on silica [37]. In this paper, we investigate the mechanism of the formation of solid supported bilayers (SSB) via spontaneous adsorption of vesicles onto spherical silica surfaces. Formation kinetics and stability under various conditions of charge density and salt is explored. The mobility of phospholipids is varied by varying the temperature and the kinetic behavior is interpreted to gain more insight into the mechanism of the adsorption. To our knowledge, this is the only work which investigates the temperature dependence of lipid mobility to correlate with adsorption efficiency and coverage. 2. Materials At the working pH of 7, neutral phospholipid dimyristoyl-sn-glycero-3-phosphocholine (DMPC) 99.5% purity, and negatively charged phospholipid dimyristoyl-sn-glycero-3-phosphoglycerol (DMPG) 99% were purchased from Genzyme Pharmaceuticals, Cambridge, USA. The fluorophore lipid NBD-PE (99% purity) was purchased from Molecular Probes, Inc. Tris Hydrochloride (99.91% purity) was supplied by Mallinckrodt GenAR, Inc. NucleosilÒ spherical silica gel with a 4000 Å pore size and 10 m particle size was purchased from Macherey–Nagel, Inc. The total surface area for these particles was determined to be 10.7 l2/g using the BET method. All chemicals were used without further purification. Milli-Q water was used for the preparation of all solutions.
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dissolving them in a chloroform/methanol mixture to achieve complete mixing. NBD-PE fluorescence probe molecules were dissolved in chloroform and added to the lipid mixture in amounts not exceeding 0.05 mol.% in total lipid concentration. A thin film of mixed lipids was obtained by evaporation of the solvents from the medium using a rotary evaporator. The thin film was kept under vacuum for a few hours to ensure complete dryness. Tris buffer at pH 7 was added to the thin film and the temperature was maintained at around 50 °C (which is well above the phase transition temperature of the lipids) overnight for complete hydration. Multilamellar vesicles were obtained on vortexing the solution for a few minutes. This solution was then sonicated with a tip sonicator (Branson Sonifier 450 at 30% duty cycle) for a period that depended on the lipid mixture composition. The breaking down of multilamellar vesicles to form small unilamellar vesicles (SUV) with an average diameter of 50 nm was apparent from the change in the optical density of the dispersions; the dispersions become almost optically clear. 3.2. Preparation of bilayers on solid support The direct fusion technique [38] was employed for the preparation of supported lipid membranes. Because of their high bending curvature, SUVs are internally unstable and can be assembled readily onto highly hydrophilic surfaces via incubation and adsorption. This technique can also be applied to the preparation of supported lipid monolayers, provided that vigorous stirring is used during the initial stages of the incubation. In the adsorption kinetics experiments, 50 mg dry silica beads were added to 5 mL SUV dispersions with a total lipid concentration of 1.2 mmol dm3. The mixture was left for the required amount of time under fast magnetic stirring. During the adsorption, temperature was controlled using a jacketed glass beaker. For equilibrium experiments, 5 mL vesicle dispersions containing various total lipid concentrations were mixed with 50 mg silica gel and stirred for at least 24 h to allow adsorption equilibration. After the adsorption, the mixture was transferred into 15 mL conical centrifuge tubes and centrifuged for 10 min at 4000 rpm at the required temperature using an Eppendorf Centrifuge 5810 R. This procedure resulted in the precipitation of silica gel with adsorbed lipid. The supernatant was carefully removed by a pipette and transferred to small vials to await analysis. The extent of silica coating was quantified by measuring the NBD-PE fluorescence intensities of the initial SUV and supernatant solutions using a SpexFluoroMax fluorescent spectrophotometer. The excitation and emission wavelengths were 463 and 530 nm, respectively. This indirect measurement of adsorbed lipid on silica surfaces assumes that the fluorescent lipid molecules behave similarly to the normal phospholipid molecules and are uniformly distributed in the mixed lipid vesicles. In the desorption experiments, the coated beads obtained from the centrifugation were rinsed with fresh Tris buffer, stirred gently for a minute, and centrifuged; the supernatant was decanted. This cycle was repeated six times to remove excess lipid vesicles adhered to the membrane. The coated beads were then re-suspended in Tris buffer with or without NaCl and the mixture was stirred continuously for different periods of time. After centrifugation, the supernatant fluorescence intensity was measured to determine the detached lipid amount.
3. Experimental
3.3. Determination of phase transition temperature
3.1. Preparation of vesicle solution
Differential scanning calorimetry (DSC) was used to observe phase transitions in these systems, as determined by the additional heat added (i.e. the corresponding excess power) required to maintain a constant rate of temperature rise, which provides a direct
Multilamellar vesicle suspensions (MLV) were prepared by weighing a proportion of dry lipids in a round-bottom flask, and
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measure of the energy change associated with the transition. DSC experiments were performed using a SETERAM MicroCal high sensitivity VP-DSC unit.
4. Results and discussion We have investigated the properties of supported lipid bilayers prepared by incubating silica beads with vesicles of varying compositions, to show that their structure, phase transition properties, and stability can be manipulated to provide the desired functionalities. Quantitative adsorption kinetic and equilibrium experiments give insight into the formation mechanisms of lipid bilayers on bare silica beads. These results are discussed in the sections that follow.
4.1. Bilayers at equilibrium Equilibrium adsorption isotherms for pure DMPC and mixed DMPC/DMPG vesicles deposited as bilayers on the silica supports when vesicles of various lipid compositions were incubated with the silica beads are shown in Fig. 1. The broken horizontal lines indicate the loading that would be obtained for complete bilayer coverage of the bead surfaces. Complete coverage is calculated by assuming only the outer surface of silica particles (10 lm in diameter) to be0 covered by lipid molecules with an individual projected area of 50 Å A. The results given in Fig. 1, obtained in the absence of added salt, indicate that a complete coating of the solid support was obtained with pure, neutrally charged DMPC vesicles, even at low lipid concentrations (>0.2 mM). With negatively charged (DMPG) mixed lipid vesicles at the same total lipid concentration, however, bilayer deposition on the solid support was incomplete, although greater coverage of the solid support was achieved for any given lipid mixture with increasing total lipid concentration. The role played by electrostatic interactions in mediating bilayer deposition is evident from the trends shown in Fig. 2, in which the adsorption of mixed vesicles to silica surfaces is seen to depend strongly on ionic strength. With increasing salt concentration, the equilibrium adsorbed lipid amount increased due to the Debye screening of the repulsive electrostatic interactions. For a 0.02 M NaCl solution, the Debye screening length is 21.5 Å, which is larger than the water layer thickness between the silica surface and the lipid bilayer (10–20 Å) [39], and thus there was an electrostatic barrier to the vesicle adsorption. At the higher NaCl concentration of 0.05 M, the Debye length reduces to 13.6 Å, which
Fig. 1. Equilibrium of lipid bilayer formation on silica beads via SUV adsorption Experimental conditions: vesicles suspended in 10 mMTris–HCl, pH 7.0 buffer solutions; 5 ml SUV dispersion incubated with 0.05 g nucleosil silica beads at room temperature under vigorous stirring for at least 24 h to reach equilibrium.
Fig. 2. Ionic strength effect on equilibrium of lipid bilayer formation on silica beads Experimental conditions: vesicles containing 20% DMPG and 80% DMPC were suspended in 10 mMTris–HCl, pH 7.0 and different concentration of NaCl buffer solutions; 5 ml SUV dispersion was incubated with 0.05 g nucleosil silica beads at room temperature under vigorous stirring for at least 24 h.
is smaller than the typical water film thickness, and thus the electrostatic repulsion was effectively screened.
4.2. Stability of supported bilayers The physical stability of these noncovalently assembled lipid films was investigated by washing the lipid coated beads with aqueous solutions of different salt strength. The silica-supported bilayers in lipid-free aqueous solution were unstable only when negatively charged lipids were incorporated into the membranes, as is evident from Fig. 3, in which the desorption of lipids from the silica surfaces prepared with 80/20 DMPC/DMPG vesicles on washing with buffer solutions. Although a small fraction of the lipids detach from the beads in the absence of the salt, the presence of 0.1 salt solution suppresses this desorption entirely. Increasing ionic strength of the buffer solution provides a greater inhibition of lipid desorption by shielding the electrostatic repulsions between the lipids and the charged silica surfaces. When the negatively charged lipid DMPG is present in the starting vesicle dispersion, some will reside in the inner leaflet of the coated solid support at a distance of approximate 20 Å from the silica surface, and will be repelled by the surface. Subsequent washing of the supported bilayer may facilitate desorption of lipids as a result of this repulsive force. In the presence of NaCl,
Fig. 3. Lipid loss from supported bilayer membrane via desorption Experimental conditions: fully coated silica beads were gently washed to remove excess vesicles (wash about six times); the beads were then incubated with 5 ml 10 mM Tris–HCl, pH 7.0 and with or without NaCl buffer at room temperature under vigorous stirring.
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Fig. 4. Differential Scanning Calorimetry results of (a) 100% DMPC, (b) 95%DMPC and 5%DMPG, (c) 90%DMPC and 10%DMPG, (d) 80%DMPC and 10%DMPG, (e) 100% DMPG.
the negative charge on the bare silica surface is screened, enhancing the bilayer stability by preventing lipid loss from the bilayer. 4.3. Phase transitions of supported lipid membranes Phospholipid membranes undergo a reversible temperaturedependent phase transition from an ordered (gel) state to a more disordered ‘‘fluid” (liquid–crystalline) state or vice versa. Such phase transition behavior is characterized primarily by a main phase transition temperature Tm [40] determined using DSC. Published studies of single DMPC and DPPC bilayers on monodispersed spherical glass or silica beads show that the main transition
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temperatures are about 2 °C lower than those of the corresponding MLVs and that the formation of the rippled phase is suppressed, probably due to lateral stresses induced in the membrane. The two leaflets exhibit a cooperative main phase transition, indicating that they interact strongly. The lateral stress, which increases with substrate curvature, has a significant effect on phase behavior, resulting in the shift of solidus and liquidus points during the transition [27,41]. The phase transition behavior of the supported membranes in this work is in general similar to that of MLVs and results in the formation of ordered layers on the substrate surfaces. The phase transitions for vesicle solutions evident in the DSC traces shown in Fig. 4 indicate that although the two lipids used both have C14 chains, they do not exhibit the same phase transition temperature. Vesicles composed of 100% DMPC show a transition peak at 24.6 °C, whereas the 100% DMPG vesicles have a transition peak at 28.7 °C. Vesicles with different lipid compositions transition from gel to fluid state at temperatures between 24.6 and 28.7 °C. Two separate transition peaks can be seen for the 95/5 DMPC/DMPG system, while at other compositions, broadening obscures the splitting of the peaks. Below the transition temperature, lipids are in a more gel-like state where the lateral movements, flip-flops and bending are more restricted, and thus we anticipate that the adsorption kinetics will be very different than at temperatures where the lipids are more fluid. 4.4. Kinetics of lipid membrane formation Several approaches have been utilized to monitor the kinetics of lipid bilayer formation and monolayers on both planar and curved
Fig. 5. Time courses of lipid bilayer formation on silica beads via SUV adsorption at different temperatures Experimental conditions: vesicles suspended in 10 mMTris–HCl, pH 7.0 buffer solutions; 5 ml of 1.2 mM SUV dispersion incubated with 0.05 g nucleosil silica beads at (a) T = 35 °C, (b) T = 25 °C, (c) T = 15 °C and (d) comparison of 35 °C and 15 °C for uncharged vesicles under vigorous stirring.
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surfaces. Direct optical and electrochemical monitoring methods that include total internal reflection fluorescence microscopy (TIRFM), surface plasmon resonance (SPR), ellipsometry, and impedance spectroscopy were not suitable for our work as they can only be applied to flat surfaces. We therefore determined the adsorbed amounts directly from the decrease in the supernatant concentration of fluorescently labeled lipids after vesicle solutions had been incubated with the silica beads for given periods of time. The kinetics of vesicle adsorption and bilayer formation on macroporous silica beads are shown in Fig. 5 for different vesicle compositions below and above their transition temperatures. The dashed line represents the calculated lipid density on the surface for a single phospholipid bilayer, assuming an average molecular area of 50 Å2. Within a few hours the mass of the adsorbed neutral lipid DMPC approached that expected for the formation of a single bilayer on the silica surface. In the presence of 10% or more DMPG, the surface coverage was limited even after 24 h of incubation. Slower adsorption rates were observed with increasing DMPG content, with two kinetic regimes in which a slow initial process was followed by a more rapid adsorption of the vesicles. The ionic strength of the solution can play an important role in the deposition kinetics of mixed vesicles onto the silica surface. At high salt concentrations (e.g., 0.1 M) there is no kinetic barrier to the adsorption of charged vesicles, as shown in Fig. 5b for the 80% DMPC system. Temperature has a significant effect on the kinetics of lipid adsorption. At all temperatures, the full adsorption of pure DMPC vesicles onto silica gel were complete within an hour, while that of DMPG-containing systems was incomplete and the adsorption rates were much slower. The kinetics of adsorption and equilibrium coverage decreased with a lowering of the temperature from 35 °C to 25 °C, close to their transition temperatures, and were significantly reduced when the lipid bilayers were well below these transition temperatures, as is evident from the results shown for different temperatures in Fig. 5a–c. Fig. 5d provides a closer comparison of the DMPC adsorption progress with time at two temperatures, showing more clearly the retardation in adsorption rate below the transition temperature even for these uncharged vesicles. 4.5. Mechanism of formation of lipid membranes Vesicle fusion allows the deposition of closed membranes of macroscopic lateral dimensions (i.e. millimeter to centimeter scale) that would be difficult to obtain in other ways. An appreciation of the mechanisms of lipid deposition in which lipids that are aggregated in vesicles spontaneously coat the silica surface at the cost of losing their vesicle integrity is important for the design of
systems that exploit these phenomena. Vesicles prepared via sonication are forced mechanically to form structures with a very high bending energy that can be relieved by fusion to form surfaces of lower curvature, and, eventually, flat lamellar phases. Thus, in the presence of an almost flat surface (10 mm diameter silica particles); vesicles will be inclined to spread over the surfaces to alleviate the bending energy. It has been suggested that there are four sequential steps involved in the deposition of a lipid bilayer on a hydrophilic surface [42]: (i) vesicle approach to the surface, (ii) vesicle adhesion, (iii) vesicle rupture, and (iv) subsequent spreading and fusion of the adsorbed vesicle bilayers. The first three steps can proceed readily with sufficiently attractive surfaces, while lateral bilayer spreading depends critically on the degree of the hydration of the substrate. Experimental studies using reflection interference contrast microscopy (RICM) demonstrated two distinctive bilayer spreading behaviors: spreading by a single open edged bilayer and spreading of a membrane fold or lobe by means of a tank-tread-like rolling process [43]. Rolling is the dominant mechanism for surfaces that strongly dehydrate the membrane and lead to a thin film of bound water. In contrast, on surfaces that are very hydrophilic but do not dehydrate the membrane, a thin water layer forms between the membrane and the solid surface, allowing the bilayer to spread by sliding, and thereby to form a continuous supported bilayer. According to our results, a possible mechanism of vesicle adsorption onto silica is suggested. The adsorption experiments reported above indicate clearly that the kinetics is affected significantly by whether the temperature is above or below the transition temperature. For uncharged DMPC lipid vesicles, the adsorption is rapid, but faster above the transition temperature than below, which we attribute to the change in fluidity of the bilayer. For charged systems, adsorption behavior is dictated to some extent by the repulsive interactions between the anionic lipids in the outer envelope of the vesicle bilayer and the negatively charged silica surface (f-potential of –21 mV). The adsorption can, nevertheless, occur because of the mobility of the lipids within this bilayer, which allows the charged lipids to migrate to regions further removed from the approach face of the vesicle to mitigate the charge interactions with the silica surface (Fig. 6a). The adsorbed vesicle ruptures and spreads over the surface to form a disc-like lamellar structure in which the negatively charged lipids preferentially reside at the edges where the curvature is highest and repulsive interactions are minimized (Fig. 6b). Further incoming vesicles can continue to adsorb on the bare silica surface, or they can adsorb on and be assimilated by the previously adsorbed bilayer discs. Initially, vesicle adsorption on the bare silica predominates, but as more of the surface becomes covered the probability of adsorption on the previously formed discs
Fig. 6. Schematic representation of the proposed mechanism of vesicle adsorption on silica. Red represents negatively charged whereas blue represents neutral surfaces and lipids: (a) approach of vesicles; (b) initial stages of vesicle adsorption onto silica; (c) pathways of further vesicle adsorption onto silica and (d) saturation of silica surface with adsorbed lipids. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
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Fig. 7. f-potential of vesicles Experimental conditions: 2 mM vesicle suspension in 10 mMTris–HCl at pH 7.0.
increases (Fig. 6c). In this instance, the net effect is for the disc to grow in size. This adsorption will be faster than onto bare silica because the repulsive interactions are reduced owing to the large uncharged surface offered by the discs. At some point the surface-toedge area ratio increases sufficiently that the edges cannot accommodate the majority of the negative charges, and the surfaces of the discs become more strongly charged, thereby providing a less favorable environment for further adsorption. At the same time, the discs begin to interact with each other, and when the surface coverage reaches a critical value, further increases in coverage lead to increasingly unfavorable repulsive interactions between the discs, thereby inhibiting further adsorption and coverage (Fig. 6d). Below the phase transition temperature, the mobility of lipids within the bilayer is greatly restricted and hence there will be no charge distribution and amelioration of the charge interactions between the vesicle and the charged silica surfaces. The fusion of the vesicle with the surface and subsequent spreading is retarded due to the rigidity of the gel-like lipids in the bilayer, as noted above for the uncharged vesicle adsorption. Fig. 7 shows the zeta potential of bare silica along with lipidcoated silica and of vesicles with different lipid compositions. The f-potential of the DMPC-coated surface is the same as that of DMPC vesicles, which shows that complete coverage is achieved and no bare silica is exposed. At the other end of the spectrum, when 100% DMPG (negatively charged at pH 7) is used, no adsorption is observed. The f-potential of the DMPG-incubated silica (19.6) is essentially that of bare silica (21.0), which indicates lack of adsorption. In the presence of negatively charged lipids, the zeta potential values are similar to that of bare silica, which is consistent with the limited coverage observed in adsorption equilibrium and kinetic studies. 5. Conclusions The adsorption of small unilamellar pure and mixed lipid vesicles on silica beads with their subsequent redistribution over the surfaces of these beads to form solid-supported lipid bilayers has been investigated for a pair of lipids, one (DMPC) uncharged and the other (DMPG) negatively charged. The equilibrium surface coverage of the beads depends on the relative charged state of the vesicles – complete coverage can be attained quickly with uncharged vesicles, while even with only 10–20% charged lipids in the mixtures, incomplete coverage of the negatively charged beads is observed. It is evident that the equilibrium properties of the supported lipid bilayers are strongly dependent on electrostatic interactions, which can be screened by increasing the ionic strength of the solution to which the lipid vesicles and bilayers are exposed. The kinetics of the adsorption process, which follows a twostage process with an initial slow stage followed by a second, faster
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stage before equilibrium adsorption is reached, shed some light on the effects of vesicle charge on the formation of the supported bilayers. A negatively charged vesicle can adsorb on the silica surface because the anionic lipids are able to redistribute on the vesicle surface away from the approach front to reduce electrostatic repulsions, but the rate of adsorption is still mediated and slowed by charge–charge interactions. Once adsorbed, the vesicle opens up and spreads over the surface to form a disc-like lipid bilayer on the bead surface. The negatively charged lipids accumulate on the edges of the discs, thus leaving the uncharged surfaces on the disc available for the adsorption of more vesicles and the growth of the discs. The rate of vesicle adsorption accelerates as the discs grow and more uncharged surfaces become available. This growth phase is arrested by the final need for anionic lipids to be accommodated on the disc surfaces as the edge-to-surface ratio (inversely proportional to the disc size) decreases and the edges cannot take up all the incoming anionic lipids, thereby inhibiting the adsorption of the vesicles, and by the charged interactions between the edges of neighboring discs as they grow and approach each other. These processes are affected also by the mobility of the lipids within the bilayers, which is considerably lower below than above their gel transition temperatures. When the lipids are in ‘gel’ state, the reduced mobility hinders the organization of lipids in the bilayer. When negatively charged lipids are present in the vesicle, without being able to organize laterally and/or across the bilayer, the charge redistribution as the vesicle approaches the negatively charged silica surface becomes extremely challenged and this leads to very little to no coverage. When there is no electrostatic repulsion between the vesicles and the silica surface, reorganization is not required and even in ‘gel’ state, vesicles can approach and spread on the surface. As shown in this study, investigation of kinetics of adsorption at different mobility of lipids clearly indicates that reorganization of lipids is actually the mechanism of vesicle adsorption and determining factor in the percent coverage of lipids on silica surfaces. References [1] K. Akamatsu, Radiat. Phys. Chem. 78 (2009) 1179. [2] T.P. Herringson, J.G. Altin, J. Control. Release 139 (2009) 229. [3] D. Zucker, D. Marcus, Y. Barenholz, A. Goldblum, J. Control. Release 139 (2009) 73. [4] J.-H. Wang, E.L. Reinherz, Curr. Opin. Struct. Biol. 10 (2000) 656. [5] S.L. Johnson, T.M. Bayerl, D.C. McDermott, G.W. Adam, A.R. Rennie, R.K. Thomas, E. Sackmann, Biophys. J. 59 (1991) 289. [6] F.M. Linseisen, M. Hetzer, T. Brumm, T.M. Bayerl, Biophys. J. 72 (1997) 1659. [7] M. Kasbauer, T.M. Bayerl, Langmuir 15 (1998) 2431. [8] A.L. Plant, Langmuir 9 (1993) 2764. [9] A.L. Plant, M. Gueguetchkeri, W. Yap, Biophys. J. 67 (1994) 1126. [10] A.L. Plant et al., Anal. Biochem. 226 (1995) 342. [11] C.W. Meuse, S. Krueger, C.F. Majkrzak, J.A. Dura, J. Fu, J.T. Connor, A.L. Plant, Biophys. J. 74 (1998) 1388. [12] J.B. Hubbard, V. Silin, A.L. Plant, Biophys. Chem. 75 (1998) 163. [13] H. Lang, C. Duschl, M. Gratzel, H. Vogel, Thin Solid Films 210/211 (1992) 818. [14] C. Pidgeon et al., Anal. Biochem. 194 (1991) 163. [15] Y.Y. Cheng, J.C. Song, L. Hanlan, C. Pidgeon, Adv. Drug Deliver. Rev. 23 (1996) 229. [16] Z. Yanxiao, Z. Cheng-Ming, L. Yi-Ming, S. Hjerten, P. Lundahl, J. Chromatogr., A 749 (1996) 13. [17] F. Beigi, P. Lundahl, J. Chromatogr., A 852 (1999) 313. [18] F. Beigi, Q. Yang, P. Lundahl, J. Chromatogr., A 704 (1995) 315. [19] P. Lundahl, F. Beigi, Adv. Drug Deliver. Rev. 23 (1997) 221. [20] Q. Yang, X.-Y. Liu, S.-I. Ajiki, M. Hara, P. Lundahl, J. Miyake, J. Chromatogr., B 707 (1998) 131. [21] N.J. Lynch, P.K. Kilpatrick, R.G. Carbonell, Biotechnol. Bioprocess Eng. 50 (1996) 169. [22] J.D. Powers, P.K. Kilpatrick, R.G. Carbonell, Biotechnol. Bioprocess Eng. 33 (1989) 173. [23] J.D. Powers, P.K. Kilpatrick, R.G. Carbonell, Biotechnol. Bioprocess Eng. 36 (1990) 506. [24] J.D. Powers, P.K. Kilpatrick, R.G. Carbonell, Biotechnol. Bioprocess Eng. 44 (1994) 509. [25] T.M. Bayerl, S. Bayerl, US Patent, USA, 1997. [26] H.M. Reinl, T.M. Bayerl, Biochemistry 33 (1994) 14091.
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Journal of Colloid and Interface Science www.elsevier.com/locate/jcis
Dynamics of supported lipid bilayer deposition from vesicle suspensions Seyda Bucak *, Chen Wang 1, Paul E. Laibinis 2, T. Alan Hatton Department of Chemical Engineering, Massachusetts Institute of Technology, Cambridge, MA 02139, USA
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Article history: Received 10 March 2010 Accepted 29 April 2010 Available online 4 May 2010 Keywords: Solid-supported membranes Liposomes Surface coverage Adsorption kinetics
a b s t r a c t The equilibrium and dynamic adsorption behavior of mixed lipid (DMPC and DMPG) vesicles on a hydrophilic silica bead substrate indicate that the adsorption is a two-step process. Vesicles initially adsorb and spread on the silica surface as lamellar bilayer discs, with a rate mediated by the charge–charge interactions between vesicles and the silica beads. Vesicle adsorption occurs more readily on the growing discs than on the bare silica surface, and consequently the adsorption rate increases as the discs grow. Eventually, the growth is arrested because of unfavorable charge interactions between the discs as they cover more of the surface, and the equilibrium condition of partial surface coverage is attained. Ó 2010 Elsevier Inc. All rights reserved.
1. Introduction Over the past two decades, special attention has been paid to vesicles made from both natural and synthetic phospholipids with the realization of their potential applications as drug delivery carriers and as model systems for the study of cell–cell and cell–membrane interactions in the immune system [1–4]. A number of problems were noted, however, when quantitative studies of the properties and behavior of the artificial lipid membrane systems were required. These include the stability of the prepared liposome dispersion over time due to vesicle fusion, the variable number of bilayers and water content in each compartment, and the change of phase structures under different conditions [5]. A well-defined lipid bilayer system is thus necessary to facilitate investigations on e.g., lipid–protein interactions. Lipid membranes supported on solid substrates can be prepared to meet these needs, with potential use also in more practical applications. The number of lipid layers in supported lipid membranes can be controlled by employing a suitable substrate and preparation method. Two types of membranes that are particularly interesting are the so-called supported lipid bilayer (SLB) and supported lipid monolayer (SLM). An SLB is formed on a hydrophilic surface such as silica, quartz, oxidized silicon, and glass consists of two leaflets of phospholipids [6]. An SLM can be assembled on silane-derivatized silica surfaces [6,7] or thiol-coated gold surfaces [8–12], or it can be
* Corresponding author. Present address: Yeditepe University, Department of Chemical Engineering, Kayisdagi, Istanbul, Turkey. E-mail address: [email protected] (S. Bucak). 1 Present address: Millipore Corporation, USA. 2 Present address: Vanderbilt University, Department of Chemical and Biomolecular Engineering, Nashville, TN 37235, USA. 0021-9797/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.jcis.2010.04.087
obtained by the covalent linking of chemically modified phospholipids with silica [13] or gold. Another type of lipid monolayer, also called immobilized artificial membrane (IAM), can be obtained by the direct covalent bonding of phospholipid analogs on silica particles [14,15]. Such lipid/membrane systems are geometrically well defined and are potentially suitable for the aforementioned quantitative studies. Supported lipid membranes have been used widely in several applications such as preparation of biosensors for environmental monitoring, drug development, medical diagnostics and industrial process control. Immobilized artificial membrane (IAM) chromatography has been used successfully to measure drug–membrane interactions and drug membrane permeability. In addition to IAM, immobilized liposome chromatography (ILC) was developed by Lundahl’s research group in Sweden to study drug–membrane interactions [16–20]. Free lipid vesicles have been combined with ultrafiltration or precipitation to separate protein mixtures [21– 24]. Bayerl and Bayerl coated macroporous silica with lipid bilayers and used these treated beads as the stationary phase for gel permeation and ion exchange chromatography [25]. Another conceptually novel chromatography process, dubbed phase transition chromatography (PTC), was also developed by Bayerl’s research group. Although SLB have been extensively used in several applications, the mechanism of vesicle adsorption onto flat (or near flat) surfaces is still under investigation. Lipid molecules in the supported bilayers can exchange with those in solution, just as in free vesicle suspensions, although this lipid transfer between bilayers on different substrates occurs at a much slower rate than that between free vesicles. It is inferred that lipid monomers dissociate from the surface lipid layers or vesicles and fuse with other bilayer patches. A surface may stabilize the adsorbed SUVs by increasing
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the activation energy barrier for lipid desorption from the supported membrane, while inhibition of collisions between SUVs and supported bilayers also reduces the lipid exchange rate [26]. Bayerl et al. observed a difference in the phase transition temperature of vesicle bilayers in aqueous media and solid supported bilayers [27], which they concluded was due to increased demixing of two lipids in the medium in the two-phase region with increasing curvature due to lowering of the lateral pressure. The formation of supported lipid bilayers on silica nanoparticles was observed within a few minutes of contact time by cryo-electron microscopy [28]. They have shown that bilayer formation was observed with negatively charged, neutral and positively charged lipids but not with highly negatively charged particles where rupture of vesicles did not take place. Zhadanov and Kasemo [29] suggested that the support-induced deformation of an adsorbed vesicle is further enhanced by the adsorption of vesicles in its vicinity. AFM studies have shown fusion of adsorbed vesicles [30]. Phospholipid bilayer formation at bare silica surface was also studied in situ with neutron reflection [31]. Although the first few minutes could not be captured (which is believed to be the adsorption of individual phospholipid vesicles on the bare silica surface), the next stage where growth of bilayer covered area by fusion of single spots was observed until a homogenous stable and fluid phospholipid bilayer covered the surface. Adsorption onto different surfaces [32], effect of hydrophobicity of the lipid tails on adsorption [33] and effect of presence of salt in adsorption of charged lipids [34] have also been studied. In has been found that in order to deposit onto solid, oppositely charged surfaces, charged bilayers also have certain requirements of bilayer fluidity [35]. DODAB vesicles are also found in the rigid gel state at room temperature [36] and closed DODAB vesicles are difficult to disrupt for bilayer deposition so that previously disrupted vesicles or bilayer fragments had to be used in order to optimize gel bilayer deposition on silica [37]. In this paper, we investigate the mechanism of the formation of solid supported bilayers (SSB) via spontaneous adsorption of vesicles onto spherical silica surfaces. Formation kinetics and stability under various conditions of charge density and salt is explored. The mobility of phospholipids is varied by varying the temperature and the kinetic behavior is interpreted to gain more insight into the mechanism of the adsorption. To our knowledge, this is the only work which investigates the temperature dependence of lipid mobility to correlate with adsorption efficiency and coverage. 2. Materials At the working pH of 7, neutral phospholipid dimyristoyl-sn-glycero-3-phosphocholine (DMPC) 99.5% purity, and negatively charged phospholipid dimyristoyl-sn-glycero-3-phosphoglycerol (DMPG) 99% were purchased from Genzyme Pharmaceuticals, Cambridge, USA. The fluorophore lipid NBD-PE (99% purity) was purchased from Molecular Probes, Inc. Tris Hydrochloride (99.91% purity) was supplied by Mallinckrodt GenAR, Inc. NucleosilÒ spherical silica gel with a 4000 Å pore size and 10 m particle size was purchased from Macherey–Nagel, Inc. The total surface area for these particles was determined to be 10.7 l2/g using the BET method. All chemicals were used without further purification. Milli-Q water was used for the preparation of all solutions.
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dissolving them in a chloroform/methanol mixture to achieve complete mixing. NBD-PE fluorescence probe molecules were dissolved in chloroform and added to the lipid mixture in amounts not exceeding 0.05 mol.% in total lipid concentration. A thin film of mixed lipids was obtained by evaporation of the solvents from the medium using a rotary evaporator. The thin film was kept under vacuum for a few hours to ensure complete dryness. Tris buffer at pH 7 was added to the thin film and the temperature was maintained at around 50 °C (which is well above the phase transition temperature of the lipids) overnight for complete hydration. Multilamellar vesicles were obtained on vortexing the solution for a few minutes. This solution was then sonicated with a tip sonicator (Branson Sonifier 450 at 30% duty cycle) for a period that depended on the lipid mixture composition. The breaking down of multilamellar vesicles to form small unilamellar vesicles (SUV) with an average diameter of 50 nm was apparent from the change in the optical density of the dispersions; the dispersions become almost optically clear. 3.2. Preparation of bilayers on solid support The direct fusion technique [38] was employed for the preparation of supported lipid membranes. Because of their high bending curvature, SUVs are internally unstable and can be assembled readily onto highly hydrophilic surfaces via incubation and adsorption. This technique can also be applied to the preparation of supported lipid monolayers, provided that vigorous stirring is used during the initial stages of the incubation. In the adsorption kinetics experiments, 50 mg dry silica beads were added to 5 mL SUV dispersions with a total lipid concentration of 1.2 mmol dm3. The mixture was left for the required amount of time under fast magnetic stirring. During the adsorption, temperature was controlled using a jacketed glass beaker. For equilibrium experiments, 5 mL vesicle dispersions containing various total lipid concentrations were mixed with 50 mg silica gel and stirred for at least 24 h to allow adsorption equilibration. After the adsorption, the mixture was transferred into 15 mL conical centrifuge tubes and centrifuged for 10 min at 4000 rpm at the required temperature using an Eppendorf Centrifuge 5810 R. This procedure resulted in the precipitation of silica gel with adsorbed lipid. The supernatant was carefully removed by a pipette and transferred to small vials to await analysis. The extent of silica coating was quantified by measuring the NBD-PE fluorescence intensities of the initial SUV and supernatant solutions using a SpexFluoroMax fluorescent spectrophotometer. The excitation and emission wavelengths were 463 and 530 nm, respectively. This indirect measurement of adsorbed lipid on silica surfaces assumes that the fluorescent lipid molecules behave similarly to the normal phospholipid molecules and are uniformly distributed in the mixed lipid vesicles. In the desorption experiments, the coated beads obtained from the centrifugation were rinsed with fresh Tris buffer, stirred gently for a minute, and centrifuged; the supernatant was decanted. This cycle was repeated six times to remove excess lipid vesicles adhered to the membrane. The coated beads were then re-suspended in Tris buffer with or without NaCl and the mixture was stirred continuously for different periods of time. After centrifugation, the supernatant fluorescence intensity was measured to determine the detached lipid amount.
3. Experimental
3.3. Determination of phase transition temperature
3.1. Preparation of vesicle solution
Differential scanning calorimetry (DSC) was used to observe phase transitions in these systems, as determined by the additional heat added (i.e. the corresponding excess power) required to maintain a constant rate of temperature rise, which provides a direct
Multilamellar vesicle suspensions (MLV) were prepared by weighing a proportion of dry lipids in a round-bottom flask, and
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measure of the energy change associated with the transition. DSC experiments were performed using a SETERAM MicroCal high sensitivity VP-DSC unit.
4. Results and discussion We have investigated the properties of supported lipid bilayers prepared by incubating silica beads with vesicles of varying compositions, to show that their structure, phase transition properties, and stability can be manipulated to provide the desired functionalities. Quantitative adsorption kinetic and equilibrium experiments give insight into the formation mechanisms of lipid bilayers on bare silica beads. These results are discussed in the sections that follow.
4.1. Bilayers at equilibrium Equilibrium adsorption isotherms for pure DMPC and mixed DMPC/DMPG vesicles deposited as bilayers on the silica supports when vesicles of various lipid compositions were incubated with the silica beads are shown in Fig. 1. The broken horizontal lines indicate the loading that would be obtained for complete bilayer coverage of the bead surfaces. Complete coverage is calculated by assuming only the outer surface of silica particles (10 lm in diameter) to be0 covered by lipid molecules with an individual projected area of 50 Å A. The results given in Fig. 1, obtained in the absence of added salt, indicate that a complete coating of the solid support was obtained with pure, neutrally charged DMPC vesicles, even at low lipid concentrations (>0.2 mM). With negatively charged (DMPG) mixed lipid vesicles at the same total lipid concentration, however, bilayer deposition on the solid support was incomplete, although greater coverage of the solid support was achieved for any given lipid mixture with increasing total lipid concentration. The role played by electrostatic interactions in mediating bilayer deposition is evident from the trends shown in Fig. 2, in which the adsorption of mixed vesicles to silica surfaces is seen to depend strongly on ionic strength. With increasing salt concentration, the equilibrium adsorbed lipid amount increased due to the Debye screening of the repulsive electrostatic interactions. For a 0.02 M NaCl solution, the Debye screening length is 21.5 Å, which is larger than the water layer thickness between the silica surface and the lipid bilayer (10–20 Å) [39], and thus there was an electrostatic barrier to the vesicle adsorption. At the higher NaCl concentration of 0.05 M, the Debye length reduces to 13.6 Å, which
Fig. 1. Equilibrium of lipid bilayer formation on silica beads via SUV adsorption Experimental conditions: vesicles suspended in 10 mMTris–HCl, pH 7.0 buffer solutions; 5 ml SUV dispersion incubated with 0.05 g nucleosil silica beads at room temperature under vigorous stirring for at least 24 h to reach equilibrium.
Fig. 2. Ionic strength effect on equilibrium of lipid bilayer formation on silica beads Experimental conditions: vesicles containing 20% DMPG and 80% DMPC were suspended in 10 mMTris–HCl, pH 7.0 and different concentration of NaCl buffer solutions; 5 ml SUV dispersion was incubated with 0.05 g nucleosil silica beads at room temperature under vigorous stirring for at least 24 h.
is smaller than the typical water film thickness, and thus the electrostatic repulsion was effectively screened.
4.2. Stability of supported bilayers The physical stability of these noncovalently assembled lipid films was investigated by washing the lipid coated beads with aqueous solutions of different salt strength. The silica-supported bilayers in lipid-free aqueous solution were unstable only when negatively charged lipids were incorporated into the membranes, as is evident from Fig. 3, in which the desorption of lipids from the silica surfaces prepared with 80/20 DMPC/DMPG vesicles on washing with buffer solutions. Although a small fraction of the lipids detach from the beads in the absence of the salt, the presence of 0.1 salt solution suppresses this desorption entirely. Increasing ionic strength of the buffer solution provides a greater inhibition of lipid desorption by shielding the electrostatic repulsions between the lipids and the charged silica surfaces. When the negatively charged lipid DMPG is present in the starting vesicle dispersion, some will reside in the inner leaflet of the coated solid support at a distance of approximate 20 Å from the silica surface, and will be repelled by the surface. Subsequent washing of the supported bilayer may facilitate desorption of lipids as a result of this repulsive force. In the presence of NaCl,
Fig. 3. Lipid loss from supported bilayer membrane via desorption Experimental conditions: fully coated silica beads were gently washed to remove excess vesicles (wash about six times); the beads were then incubated with 5 ml 10 mM Tris–HCl, pH 7.0 and with or without NaCl buffer at room temperature under vigorous stirring.
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Fig. 4. Differential Scanning Calorimetry results of (a) 100% DMPC, (b) 95%DMPC and 5%DMPG, (c) 90%DMPC and 10%DMPG, (d) 80%DMPC and 10%DMPG, (e) 100% DMPG.
the negative charge on the bare silica surface is screened, enhancing the bilayer stability by preventing lipid loss from the bilayer. 4.3. Phase transitions of supported lipid membranes Phospholipid membranes undergo a reversible temperaturedependent phase transition from an ordered (gel) state to a more disordered ‘‘fluid” (liquid–crystalline) state or vice versa. Such phase transition behavior is characterized primarily by a main phase transition temperature Tm [40] determined using DSC. Published studies of single DMPC and DPPC bilayers on monodispersed spherical glass or silica beads show that the main transition
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temperatures are about 2 °C lower than those of the corresponding MLVs and that the formation of the rippled phase is suppressed, probably due to lateral stresses induced in the membrane. The two leaflets exhibit a cooperative main phase transition, indicating that they interact strongly. The lateral stress, which increases with substrate curvature, has a significant effect on phase behavior, resulting in the shift of solidus and liquidus points during the transition [27,41]. The phase transition behavior of the supported membranes in this work is in general similar to that of MLVs and results in the formation of ordered layers on the substrate surfaces. The phase transitions for vesicle solutions evident in the DSC traces shown in Fig. 4 indicate that although the two lipids used both have C14 chains, they do not exhibit the same phase transition temperature. Vesicles composed of 100% DMPC show a transition peak at 24.6 °C, whereas the 100% DMPG vesicles have a transition peak at 28.7 °C. Vesicles with different lipid compositions transition from gel to fluid state at temperatures between 24.6 and 28.7 °C. Two separate transition peaks can be seen for the 95/5 DMPC/DMPG system, while at other compositions, broadening obscures the splitting of the peaks. Below the transition temperature, lipids are in a more gel-like state where the lateral movements, flip-flops and bending are more restricted, and thus we anticipate that the adsorption kinetics will be very different than at temperatures where the lipids are more fluid. 4.4. Kinetics of lipid membrane formation Several approaches have been utilized to monitor the kinetics of lipid bilayer formation and monolayers on both planar and curved
Fig. 5. Time courses of lipid bilayer formation on silica beads via SUV adsorption at different temperatures Experimental conditions: vesicles suspended in 10 mMTris–HCl, pH 7.0 buffer solutions; 5 ml of 1.2 mM SUV dispersion incubated with 0.05 g nucleosil silica beads at (a) T = 35 °C, (b) T = 25 °C, (c) T = 15 °C and (d) comparison of 35 °C and 15 °C for uncharged vesicles under vigorous stirring.
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surfaces. Direct optical and electrochemical monitoring methods that include total internal reflection fluorescence microscopy (TIRFM), surface plasmon resonance (SPR), ellipsometry, and impedance spectroscopy were not suitable for our work as they can only be applied to flat surfaces. We therefore determined the adsorbed amounts directly from the decrease in the supernatant concentration of fluorescently labeled lipids after vesicle solutions had been incubated with the silica beads for given periods of time. The kinetics of vesicle adsorption and bilayer formation on macroporous silica beads are shown in Fig. 5 for different vesicle compositions below and above their transition temperatures. The dashed line represents the calculated lipid density on the surface for a single phospholipid bilayer, assuming an average molecular area of 50 Å2. Within a few hours the mass of the adsorbed neutral lipid DMPC approached that expected for the formation of a single bilayer on the silica surface. In the presence of 10% or more DMPG, the surface coverage was limited even after 24 h of incubation. Slower adsorption rates were observed with increasing DMPG content, with two kinetic regimes in which a slow initial process was followed by a more rapid adsorption of the vesicles. The ionic strength of the solution can play an important role in the deposition kinetics of mixed vesicles onto the silica surface. At high salt concentrations (e.g., 0.1 M) there is no kinetic barrier to the adsorption of charged vesicles, as shown in Fig. 5b for the 80% DMPC system. Temperature has a significant effect on the kinetics of lipid adsorption. At all temperatures, the full adsorption of pure DMPC vesicles onto silica gel were complete within an hour, while that of DMPG-containing systems was incomplete and the adsorption rates were much slower. The kinetics of adsorption and equilibrium coverage decreased with a lowering of the temperature from 35 °C to 25 °C, close to their transition temperatures, and were significantly reduced when the lipid bilayers were well below these transition temperatures, as is evident from the results shown for different temperatures in Fig. 5a–c. Fig. 5d provides a closer comparison of the DMPC adsorption progress with time at two temperatures, showing more clearly the retardation in adsorption rate below the transition temperature even for these uncharged vesicles. 4.5. Mechanism of formation of lipid membranes Vesicle fusion allows the deposition of closed membranes of macroscopic lateral dimensions (i.e. millimeter to centimeter scale) that would be difficult to obtain in other ways. An appreciation of the mechanisms of lipid deposition in which lipids that are aggregated in vesicles spontaneously coat the silica surface at the cost of losing their vesicle integrity is important for the design of
systems that exploit these phenomena. Vesicles prepared via sonication are forced mechanically to form structures with a very high bending energy that can be relieved by fusion to form surfaces of lower curvature, and, eventually, flat lamellar phases. Thus, in the presence of an almost flat surface (10 mm diameter silica particles); vesicles will be inclined to spread over the surfaces to alleviate the bending energy. It has been suggested that there are four sequential steps involved in the deposition of a lipid bilayer on a hydrophilic surface [42]: (i) vesicle approach to the surface, (ii) vesicle adhesion, (iii) vesicle rupture, and (iv) subsequent spreading and fusion of the adsorbed vesicle bilayers. The first three steps can proceed readily with sufficiently attractive surfaces, while lateral bilayer spreading depends critically on the degree of the hydration of the substrate. Experimental studies using reflection interference contrast microscopy (RICM) demonstrated two distinctive bilayer spreading behaviors: spreading by a single open edged bilayer and spreading of a membrane fold or lobe by means of a tank-tread-like rolling process [43]. Rolling is the dominant mechanism for surfaces that strongly dehydrate the membrane and lead to a thin film of bound water. In contrast, on surfaces that are very hydrophilic but do not dehydrate the membrane, a thin water layer forms between the membrane and the solid surface, allowing the bilayer to spread by sliding, and thereby to form a continuous supported bilayer. According to our results, a possible mechanism of vesicle adsorption onto silica is suggested. The adsorption experiments reported above indicate clearly that the kinetics is affected significantly by whether the temperature is above or below the transition temperature. For uncharged DMPC lipid vesicles, the adsorption is rapid, but faster above the transition temperature than below, which we attribute to the change in fluidity of the bilayer. For charged systems, adsorption behavior is dictated to some extent by the repulsive interactions between the anionic lipids in the outer envelope of the vesicle bilayer and the negatively charged silica surface (f-potential of –21 mV). The adsorption can, nevertheless, occur because of the mobility of the lipids within this bilayer, which allows the charged lipids to migrate to regions further removed from the approach face of the vesicle to mitigate the charge interactions with the silica surface (Fig. 6a). The adsorbed vesicle ruptures and spreads over the surface to form a disc-like lamellar structure in which the negatively charged lipids preferentially reside at the edges where the curvature is highest and repulsive interactions are minimized (Fig. 6b). Further incoming vesicles can continue to adsorb on the bare silica surface, or they can adsorb on and be assimilated by the previously adsorbed bilayer discs. Initially, vesicle adsorption on the bare silica predominates, but as more of the surface becomes covered the probability of adsorption on the previously formed discs
Fig. 6. Schematic representation of the proposed mechanism of vesicle adsorption on silica. Red represents negatively charged whereas blue represents neutral surfaces and lipids: (a) approach of vesicles; (b) initial stages of vesicle adsorption onto silica; (c) pathways of further vesicle adsorption onto silica and (d) saturation of silica surface with adsorbed lipids. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
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Fig. 7. f-potential of vesicles Experimental conditions: 2 mM vesicle suspension in 10 mMTris–HCl at pH 7.0.
increases (Fig. 6c). In this instance, the net effect is for the disc to grow in size. This adsorption will be faster than onto bare silica because the repulsive interactions are reduced owing to the large uncharged surface offered by the discs. At some point the surface-toedge area ratio increases sufficiently that the edges cannot accommodate the majority of the negative charges, and the surfaces of the discs become more strongly charged, thereby providing a less favorable environment for further adsorption. At the same time, the discs begin to interact with each other, and when the surface coverage reaches a critical value, further increases in coverage lead to increasingly unfavorable repulsive interactions between the discs, thereby inhibiting further adsorption and coverage (Fig. 6d). Below the phase transition temperature, the mobility of lipids within the bilayer is greatly restricted and hence there will be no charge distribution and amelioration of the charge interactions between the vesicle and the charged silica surfaces. The fusion of the vesicle with the surface and subsequent spreading is retarded due to the rigidity of the gel-like lipids in the bilayer, as noted above for the uncharged vesicle adsorption. Fig. 7 shows the zeta potential of bare silica along with lipidcoated silica and of vesicles with different lipid compositions. The f-potential of the DMPC-coated surface is the same as that of DMPC vesicles, which shows that complete coverage is achieved and no bare silica is exposed. At the other end of the spectrum, when 100% DMPG (negatively charged at pH 7) is used, no adsorption is observed. The f-potential of the DMPG-incubated silica (19.6) is essentially that of bare silica (21.0), which indicates lack of adsorption. In the presence of negatively charged lipids, the zeta potential values are similar to that of bare silica, which is consistent with the limited coverage observed in adsorption equilibrium and kinetic studies. 5. Conclusions The adsorption of small unilamellar pure and mixed lipid vesicles on silica beads with their subsequent redistribution over the surfaces of these beads to form solid-supported lipid bilayers has been investigated for a pair of lipids, one (DMPC) uncharged and the other (DMPG) negatively charged. The equilibrium surface coverage of the beads depends on the relative charged state of the vesicles – complete coverage can be attained quickly with uncharged vesicles, while even with only 10–20% charged lipids in the mixtures, incomplete coverage of the negatively charged beads is observed. It is evident that the equilibrium properties of the supported lipid bilayers are strongly dependent on electrostatic interactions, which can be screened by increasing the ionic strength of the solution to which the lipid vesicles and bilayers are exposed. The kinetics of the adsorption process, which follows a twostage process with an initial slow stage followed by a second, faster
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stage before equilibrium adsorption is reached, shed some light on the effects of vesicle charge on the formation of the supported bilayers. A negatively charged vesicle can adsorb on the silica surface because the anionic lipids are able to redistribute on the vesicle surface away from the approach front to reduce electrostatic repulsions, but the rate of adsorption is still mediated and slowed by charge–charge interactions. Once adsorbed, the vesicle opens up and spreads over the surface to form a disc-like lipid bilayer on the bead surface. The negatively charged lipids accumulate on the edges of the discs, thus leaving the uncharged surfaces on the disc available for the adsorption of more vesicles and the growth of the discs. The rate of vesicle adsorption accelerates as the discs grow and more uncharged surfaces become available. This growth phase is arrested by the final need for anionic lipids to be accommodated on the disc surfaces as the edge-to-surface ratio (inversely proportional to the disc size) decreases and the edges cannot take up all the incoming anionic lipids, thereby inhibiting the adsorption of the vesicles, and by the charged interactions between the edges of neighboring discs as they grow and approach each other. These processes are affected also by the mobility of the lipids within the bilayers, which is considerably lower below than above their gel transition temperatures. When the lipids are in ‘gel’ state, the reduced mobility hinders the organization of lipids in the bilayer. When negatively charged lipids are present in the vesicle, without being able to organize laterally and/or across the bilayer, the charge redistribution as the vesicle approaches the negatively charged silica surface becomes extremely challenged and this leads to very little to no coverage. When there is no electrostatic repulsion between the vesicles and the silica surface, reorganization is not required and even in ‘gel’ state, vesicles can approach and spread on the surface. As shown in this study, investigation of kinetics of adsorption at different mobility of lipids clearly indicates that reorganization of lipids is actually the mechanism of vesicle adsorption and determining factor in the percent coverage of lipids on silica surfaces. References [1] K. Akamatsu, Radiat. Phys. Chem. 78 (2009) 1179. [2] T.P. Herringson, J.G. Altin, J. Control. Release 139 (2009) 229. [3] D. Zucker, D. Marcus, Y. Barenholz, A. Goldblum, J. Control. Release 139 (2009) 73. [4] J.-H. Wang, E.L. Reinherz, Curr. Opin. Struct. Biol. 10 (2000) 656. [5] S.L. Johnson, T.M. Bayerl, D.C. McDermott, G.W. Adam, A.R. Rennie, R.K. Thomas, E. Sackmann, Biophys. J. 59 (1991) 289. [6] F.M. Linseisen, M. Hetzer, T. Brumm, T.M. Bayerl, Biophys. J. 72 (1997) 1659. [7] M. Kasbauer, T.M. Bayerl, Langmuir 15 (1998) 2431. [8] A.L. Plant, Langmuir 9 (1993) 2764. [9] A.L. Plant, M. Gueguetchkeri, W. Yap, Biophys. J. 67 (1994) 1126. [10] A.L. Plant et al., Anal. Biochem. 226 (1995) 342. [11] C.W. Meuse, S. Krueger, C.F. Majkrzak, J.A. Dura, J. Fu, J.T. Connor, A.L. Plant, Biophys. J. 74 (1998) 1388. [12] J.B. Hubbard, V. Silin, A.L. Plant, Biophys. Chem. 75 (1998) 163. [13] H. Lang, C. Duschl, M. Gratzel, H. Vogel, Thin Solid Films 210/211 (1992) 818. [14] C. Pidgeon et al., Anal. Biochem. 194 (1991) 163. [15] Y.Y. Cheng, J.C. Song, L. Hanlan, C. Pidgeon, Adv. Drug Deliver. Rev. 23 (1996) 229. [16] Z. Yanxiao, Z. Cheng-Ming, L. Yi-Ming, S. Hjerten, P. Lundahl, J. Chromatogr., A 749 (1996) 13. [17] F. Beigi, P. Lundahl, J. Chromatogr., A 852 (1999) 313. [18] F. Beigi, Q. Yang, P. Lundahl, J. Chromatogr., A 704 (1995) 315. [19] P. Lundahl, F. Beigi, Adv. Drug Deliver. Rev. 23 (1997) 221. [20] Q. Yang, X.-Y. Liu, S.-I. Ajiki, M. Hara, P. Lundahl, J. Miyake, J. Chromatogr., B 707 (1998) 131. [21] N.J. Lynch, P.K. Kilpatrick, R.G. Carbonell, Biotechnol. Bioprocess Eng. 50 (1996) 169. [22] J.D. Powers, P.K. Kilpatrick, R.G. Carbonell, Biotechnol. Bioprocess Eng. 33 (1989) 173. [23] J.D. Powers, P.K. Kilpatrick, R.G. Carbonell, Biotechnol. Bioprocess Eng. 36 (1990) 506. [24] J.D. Powers, P.K. Kilpatrick, R.G. Carbonell, Biotechnol. Bioprocess Eng. 44 (1994) 509. [25] T.M. Bayerl, S. Bayerl, US Patent, USA, 1997. [26] H.M. Reinl, T.M. Bayerl, Biochemistry 33 (1994) 14091.
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