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Langmuir 1999, 15, 3866-3874
Formation and Stability of Complex Membrane-Mimetic Monolayers on Solid Supports Theodore M. Winger,† Peter J. Ludovice,† and Elliot L. Chaikof*,†,‡ School of Chemical Engineering, Georgia Institute of Technology, and Laboratory of Biomolecular Materials Research, Department of Surgery, Emory University School of Medicine, Atlanta, Georgia 30322 Received August 6, 1998. In Final Form: March 12, 1999
Membrane-mimetic phospholipid films with or without cholesterol and varying concentrations of one or more synthetic lipopeptide conjugates were formed on an alkylsilane self-assembled monolayer on glass by a process of lipid vesicle fusion. Using a combination of radiochemical titration with 125I-labeled lipopeptides and atomic force microscopy imaging in an aqueous environment, we have characterized the kinetics of multicomponent monolayer formation and film stability as a function of temperature, lipid alkyl chain length, cholesterol content, and the concentration of solution phase bovine serum albumin. These data confirm that multicomponent membrane-mimetic monolayers can be produced in a predictable fashion by vesicle fusion to an alkylated glass substrate. The stability of these systems, however, is compromised when operating above the phase transition temperature of the lipid component, as well as in the presence of albumin.
Introduction Despite the suggestion that supported lipid monolayers will lead to the development of new classes of molecular sensors, diagnostic devices, or medical implants with improved performance characteristics and the potential significance of these systems for the characterization of ligand-receptor and cell-cell binding, stability analyses of these systems in an aqueous environment have been limited. Most if not all of these intended applications will entail operation under relatively complex conditions which will inevitably include both specific and nonspecific interfacial interactions between the supported monolayer and aqueous phase biomacromolecules. As a consequence, these effects could well compromise the structural and functional characteristics of the membrane-mimetic film to a degree dependent upon the composition of the aqueous phase, as well as the time scale of the intended application. For example, albumin will initially adsorb to any surface placed in contact with blood or serum.1,2 The capacity of this protein to bind and become incorporated into phospholipid films as a function of time and protein concentration has been documented. It is likely that this process would potentially affect the integrity of other membranemimetic associated components. To date, supported lipid surfaces have been produced either by vesicle-based techniques or through the use of a Langmuir trough. The former approach consists of first assembling a layer of closely packed hydrocarbon chains on a substrate followed by exposure to either a dilute solution of emulsified lipids or unilamellar lipid vesicles.3-6 * Corresponding author. Present address: 1364 Clifton Road, N.E., Box M-11, Laboratory for Biomolecular Materials Research and Department of Surgery, Emory University, Atlanta, Georgia 30322. Phone: (404) 727-8413. Fax: (404) 727-3660. E-mail:
[email protected]. † Georgia Institute of Technology. ‡ Emory University School of Medicine. (1) Brash, J. L.; Lyman, D. J. J Biomed. Mater. Res. 1969, 3, 175. (2) Brash, J. L.; Uniyal, S. J. Polym. Sci. Polym. Symp. 1979, 66, 377-389. (3) Spinke, J.; Yang, J.; Wolf, H.; Liley, M.; Ringsdorf, H.; Knoll, W. Biophys. J. 1992, 63, 1667-1671.
The latter methodology is based upon Langmuir-Blodgett (LB) or Langmuir-Schaeffer techniques which involve a process of controlled dipping of an alkylated substrate through an organic amphiphilic monolayer.7 Overall, we have favored the utilization of vesicle fusion to generate supported membrane-mimetic films because of their ease of use, scalability for the production of large sample numbers, and the potential adaptability of this approach to nonplanar substrates or porous materials.8-11 However, regardless of one’s preferred methodology for film formation, the continued interest in these systems is derived from the inherent flexibility of self-assembly processes in producing chemically heterogeneous structures. This may be achieved by incorporating synthetic or genetically engineered amphiphilic conjugates with pendent groups which either consist of peptide or carbohydrate sequences or generic high affinity binding groups capable of binding to appropriately tagged biomolecules.3,12-18 Presumably, the performance of these systems will depend on both the choice of the bioactive component and its surface con(4) Seifert, K.; Fendler, K.; Bamberg, E. Biophys. J. 1993, 64, 384391. (5) Florin, E. L.; Gaub, H. E. Biophys. J. 1993, 64, 375-383. (6) Wenzl, P.; Fringeli, M.; Goette, J.; Fringeli, U. Langmuir 1994, 10, 4253-4264. (7) Ulman, A. An introduction to ultrathin organic films from Langmuir-Blodgett to self-assembly; Academic Press: New York, 1991. (8) Marra, K. C.; Winger, T. M.; Hanson, S. R.; Chaikof, E. L. Macromolecules 1997, 30, 6483-6487. (9) Marra, K. C.; Kiddani, D. D. A.; Chaikof, E. Langmuir 1997, 13, 5697-5701. (10) Winger, T. M.; Chaikof, E. L. In Materials Science of the Cell; Plant, A., Vogel, V., Ed.; MRS Publications: Pittsburgh, 1998. (11) Winger, T. M.; Chaikof, E. L. Langmuir 1998, 14, 4148. (12) Duschl, C.; Liley, M.; Corradin, G.; Vogel, H. Biophys. J. 1994, 67, 1229-1237. (13) Heyse, S.; Vogel, H.; Sa¨nger, M.; Sigrist, H. Protein Sci. 1995, 4, 2532-2544. (14) Groves, J. T.; Wu¨lfing, C.; Boxer, S. G. Biophys. J. 1996, 71, 2716-2723. (15) Winger, T. M.; Ludovice, P. J.; Chaikof, E. L. Biomaterials 1995, 16, 443-449. (16) Winger, T. M.; Chaikof, E. L. Langmuir 1997, 13, 3256-3259. (17) Sun, L.; Chaikof, E. L. Bioconjugate Chem. 1997, 8, 567-571. (18) Plant, A. L.; Brigham-Burke, M.; Petrella, E. C.; O’Shannesay, D. J. Anal. Biochem. 1995, 226, 342-248.
10.1021/la980990q CCC: $18.00 © 1999 American Chemical Society Published on Web 04/28/1999
Membrane-Mimetic Monolayers on Solid Supports
centration. In this report, we sought to characterize the formation of mixed monolayers of phospholipids and synthetic lipopeptides supported on alkylated glass surfaces and identify parameters that influence the longterm stability of these systems. Specifically, the kinetics of monolayer formation and the capacity to form surfaces with predictable concentrations of peptide pendants were defined using radiotracer techniques and atomic force microscopy. In addition, we considered the effect of temperature, phospholipid alkyl chain length, cholesterol content, and the concentration of solution phase bovine serum albumin on the stability of lipopeptide surface concentration. Significantly, the utilization of radiolabeled lipopeptides afforded a high degree of sensitivity with respect to the change in surface concentration of these low abundance membrane-mimetic components. Our data confirm that multicomponent membrane-mimetic monolayers can be produced in a predictable fashion by vesicle fusion to an alkylated glass substrate. The stability of these systems, however, is compromised when operating above the phase transition temperature of the lipid component, as well as in the presence of albumin. Experimental Methods Materials. Dipalmitoylphosphatidylcholine (DPPC), dimyristoylphosphatidylcholine (DMPC), and (4-p-maleimidophenyl)butyrate dipalmitoylphosphatidylethanolamine (DPPE-MPB) were obtained from Avanti Polar Lipids and stored at -20 °C. DPPC and DMPC were dried before use by overnight exposure to P2O5 in a desiccator under a 20 mmHg vacuum. Thiolterminated peptides, GRGDY-NH-CH2CH2-SH (P1) and WQPPRARIGY-NH-CH2CH2-SH (P2), were synthesized at the Microchemical Facility at Emory University and stored at -20 °C immediately following HPLC purification. Silica plates (20 × 20 cm; 250 µm thickness) used for thin-layer chromatography (TLC) were purchased from Whatman. The TLC plates were developed with iodine for the detection of all compounds. Free amines were detected with a ninhydrin spray (0.2 M in absolute ethanol, Alltech) and free thiols by reaction with Ellman’s reagent, and a molybdenum blue spray reagent (Alltech) was used to develop phosphorus-containing compounds. Fine silica (silica gel Davisil, grade 633, 200-425 mesh, 60 Å) was purchased from Aldrich Chemicals and dried overnight at 200 °C before use. The 15 mL polypropylene columns used for lipopeptide purification by liquid chromatography were obtained from Bio-Rad. For lipopeptide radiolabeling, Na125I (5 mCi in 0.01 N NaOH) was obtained from ICN and Iodobeads purchased from Pierce Chemicals. Desalting and removal of free iodine required 5 mL HiTrap columns (Pharmacia Biotech) and DispoDialyzer dialysis tubes (1000 MWCO) from Spectrum Microgon. Microscope borosilicate glass coverslips (S/P Cover Glass, 24 × 40 × 0.21 mm) were purchased from Baxter Scientific and Multi-Terge, an alkaline chelating detergent, obtained from E. M. Diagnostic Systems. All other chemicals and solvents (HPLC grade) were from Aldrich Chemicals. Dust-Off puff-duster cans (compressed gas filtered to 0.1 µm) were purchased from Falcon Safety Products, Inc. Distilled water was deionized and of ultrafiltered grade (18 MΩ cm, Continental). The buffers used were 250 mM sodium phosphate at pH 6.2 (buffer A), 1 wt % of octyl-β-D-glucoside (OG) in 250 mM sodium phosphate at pH 6.2 (buffer B), and 20 mM sodium phosphate (pH 7.4) with 1 wt % octyl-β-D-glucoside (OG) (buffer C). The solvents used for lipopeptide purification by liquid chromatography were hexane/chloroform/2-propanol/glacial acetic acid/ water 18:10:62:1:9 v/v/v/v/v (solvent A) and hexane/chloroform/ 2-propanol/glacial acetic acid/water 15:5:60:3:17 v/v/v/v/v (solvent B). Lipopeptide Synthesis. A mixture of 5.33 mL of purified chloroform, 2.66 mL of methanol, 400 µL of deionized water, and 120 µL of aqueous triethylamine (0.16 M) was prepared in a 1 dram glass vial. Then, 23 µmol of DPPE-MPB (954 g/mol) were weighed into a glass vial and quantitatively transferred with the above solvent mixture into a clean 8 mL glass Kimax tube which contained 1 equiv of thiol-terminated peptide. The reaction was
Langmuir, Vol. 15, No. 11, 1999 3867 monitored by TLC using chloroform/methanol/acetic acid/water (CMAW) 4/2/0.05/0.4 v/v/v/v as the eluent. After 15 h, the reaction mixture was concentrated on a SpeedVac (1 h at 23 °C and 10 min at 65 °C), frozen after the addition of 4 mL of deionized water, and lyophilized, which afforded a fluffy white solid. Lipopeptide purification was performed by liquid chromatography over a 9 × 1.5 cm silica bed. Briefly, a slurry of silica in 2-propanol/ hexane 1/1 v/v was added to the column and subsequently conditioned with 10 mL of 2-propanol, followed by 10 mL of eluents A/B 4/1 v/v. The lyophilized reaction mixture was dissolved in 1 mL of A/B 4/1 v/v and transferred onto the column, and elution was conducted using a solvent gradient. Specifically, 12 mL of A/B 4/1, 20 mL A/B 1/1, 9 mL A/B 1/2, and 30 mL B were serially added to the column, and 3 mL fractions collected and monitored by TLC. Appropriate fractions were combined, concentrated to 1 mL on a SpeedVac (1 h at 45 °C), frozen in the presence of 16 mL of deionized water/2-propanol 3/1 v/v, and lyophilized overnight. The lipopeptide typically eluted in pure form between fractions 20 and 30. The final product (coded LP1 and LP2) was stored at -20 °C in a desiccated environment. Lipopeptide Iodination. Stocks of buffer A and buffer B were prepared, and dissolved oxygen was displaced by degassing and sonicating for 10 min followed by argon bubbling for 5 min. Iodobeads were washed in buffer B and dried on filter paper. Iodobeads and the HiTrap column were conditioned by flushing with 25 mL of buffer A followed by 10 mL of buffer B at a rate of 5 mL/min. Typically, radiolabeling was performed by placing five Iodobeads in a 2 mL vial, followed by 350 µL of buffer B and 2.5-5 mCi of Na125I. Then, 5 min were allowed for effective generation of the reactive iodinating species, after which 200 µL from a 1 mM lipopeptide stock solution in buffer B were added. The reactor was shaken every 15 min for 1 h, and the reaction was terminated by transferring the liquid phase of the reaction mixture onto a HiTrap column. Elution was conducted with 5 mL of buffer B at a flow rate of 5 mL/min. The eluate was collected in 0.5 mL fractions and monitored for radioactivity and UV absorbance, and fractions corresponding to the lipopeptide peak (1.5-3.0 mL) were pooled and stored at 4 °C. To ensure the complete elimination of all residual free iodine, radiolabeled lipopeptides were dialyzed against buffer B until the CPM (counts per minute) depletion rate matched the value of 0.01163 day-1, corresponding to the natural rate of decay of 125I. Preparation of Mixed Lipopeptide/Phospholipid Liposomes by Dialysis. A stock of 450 µM DPPC micellar suspension in 1 wt % OG was prepared by dissolving 4.96 mg DPPC in 2 mL EtOH/CHCl3 1/1 v/v. The solution was evaporated to dryness with a stream of argon and further dried at 45 °C for 30 min on a SpeedVac, which was followed by the addition of 15 mL of buffer C. A micellar suspension containing either one or both lipopeptides in 1 wt % OG and 250 mM sodium phosphate (pH 6.2) was mixed with the OG/phospholipid stock at varying ratios. Characteristically, lipopeptide concentrations ranged from 0 to 3 mol %. Each mix (e1 mL) was placed in a 5 mL SpectraPor DispoDialyzer cartridge along with a small Teflon-coated magnetic stirrbar (8 × 1.5 mm). All mixes were dialyzed against 100 mL of buffer C for 30 h at 23 °C to remove unbound radioactive iodine when radiolabeled lipopeptides were utilized. The detergent was subsequently removed by further dialysis against 3 × 100 mL of 20 mM sodium phosphate at pH 7.4, allowing contact times of 24-30 h. The dialyzed vesicles were typically used within 1 day in fusion experiments. Preparation of Alkylated Glass. Coverslips were puffdusted and transferred to a class 100 cleanroom. A solution of Multi-Terge/d.i. water 1/8 v/v was applied, and the slides were rinsed with water and chloroform. Surfaces were blown dry with nitrogen and exposed to an argon plasma (500 ( 10 mTorr) for 9 min at 100 ( 1 W. Slides were then immersed for 60 min in an octadecyltrichlorosilane (OTS) reaction mixture which consisted of 248 mL of bicyclohexyl, 24.8 mL of hydrated CHCl3, and 27 mL of a solution of 63 mM OTS in anhydrous CCl4 (95%, stored in a desiccator over P2O5 at room temperature). Samples were subsequently rinsed and sonicated (∼47 kHz, 80 W) for 10 min in chloroform, rinsed once again with water, and dried with nitrogen.11 Mixed Monolayer Formation by Vesicle Fusion. Upon completion of vesicle formation by detergent dialysis, suspensions
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were adjusted to a final total lipid concentration of 140 µM and a final NaCl concentration of 150 mM in 20 mM sodium phosphate (pH 7.4). An amount of 70 µL of each vesicle mixture was pipetted into individual polystyrene microwells (Corning Costar) each containing a 3.5 × 3.5 mm OTS/glass sample which was maintained to the well bottom. Microwells were transferred into a sealed humidity-saturated chamber and stored at 50 °C without agitation. After varying vesicle fusion periods, wells were extensively rinsed by flushing each well continuously with 15 mL of buffer (150 mM NaCl in 20 mM sodium phosphate at pH 7.4) using an electrically operated pipet, followed by 12 repeated rinses, each consisting of 500 µL of buffer. Samples were then transferred under buffer to new microwells, and the surface bound radioactivity was determined using a 1274 Riagamma γ counter (LKB Instruments Inc.). All samples were prepared in quadruplicate. Considerations in the Analysis of Membrane-Mimetic Film Composition Using Radiolabeled Lipopeptides. A geometrical model of a multicomponent membrane-mimetic film on an alkylated glass substrate is presented in Figure 1. The sample exposed to fusion buffer consists of a square hydrophobic (OTS-coated) face of area a1 and four sides each of area a2 and of thickness d. Unlike the major face a1, the sides are hydrophilic since samples were cut from glass coverslips after surface alkylation. Edge effects were determined to be significant because of the relatively small substrate size. Therefore, due to differences in surface chemistry, this model assumes the self-assembly of an odd number of phospholipid monolayers on a1 and an even number on a2. The total area occupied by fused phospholipids is Bn, where
Bn ) na1 + (n + 1)4dxa1
(n ) 1, 3, 5 ...)
(1)
and n is the number of supported monolayers. The lipopeptide surface concentration can be expressed in terms of either moles per unit area of exposed substrate or as moles in the outermost monolayer. The latter expression is derived by dividing the total number of lipopeptide molecules detected on the sample by the equivalent monolayer area (Bn),
(ΓLP,Bn) )
(CPM/sample) BnR1 exp[-λt]
(2)
where ΓLP,Bn is the lipopeptide surface concentration (moles/unit area of equivalent monolayer), R1 is the specific activity of the lipopeptide (CPM/mole) at t ) 0, λ is the half-life-related decay constant for 125I, and t is the time elapsed since the initial determination of the specific activity. In a two-component system consisting of a single lipopeptide (LP1) and DPPC, an alternate formulation of ΓLP,Bn can be expressed in terms of the total number of molecules (NT), the molecular area of each species (A1 and APC), and their respective mole fractions (x1 and xPC), such that
(ΓLP1,Bn) )
NTx1 ANNT[x1A1 + (1 - x1)APC]
) x1
AN[x1A1 + (1 - x1)APC]
(3)
where AN is Avogadro’s number. Therefore, upon rearrangement of eq 2, experimental data (CPM/sample) derived from this system can be expressed as
(CPM/sample) ) BnR1 exp[-λt]
x1 AN[x1A1 + (1 - x1)APC]
(4)
which facilitates both the presentation of the data as CPM/sample as a function of the lipopeptide mole fraction (x1) and multiple least-squares linear regression analysis (Microsoft Excel, version 5.0) to determine the best fit for the adjustable parameters n, A1, and APC. Extension of this analytical model to more complex multicomponent systems requires the utilization of generalized expressions for ΓLP,Bn. For example, in a system consisting of DPPC and a series of lipopeptides (1, 2, 3, ... i), ΓLPi,Bn can be
Figure 1. Geometrical model for lipid mono/multilayer formation on glass samples of mixed hydrophobic and hydrophilic surface types. Area a1 is hydrophobic (OTS-coated), area A2 is hydrophilic (borosilicate glass). Only a1 and a2 faces were accessible to phospholipid vesicles during the fusion process. expressed in terms of the total number of molecules (NT), the molecular area of each species (Ai), and their respective mole fractions (xi), such that
(ΓLPi,Bn) )
xi
[∑x A + (1 - ∑ x )A
AN
i
i
i
PC
]
(5)
Likewise, upon rearrangement of eq 2, experimental data (CPM/ sample) derived from a multicomponent system can be expressed as i
(CPM/sample) ) Bn
∑ (Γ
LPi,Bn)Ri
exp[-λt]
(6)
1
which facilitates multiple least-squares linear regression analysis to determine the best fit for the adjustable parameters n and Ai. Determination of the Long-Term Stability of Supported Lipid Monolayers. Substrate-supported planar lipid monolayers were produced for each of the radiolabeled (LP)x: (DPPC)100-x systems (x ) 0.1, 0.5, 1, 2, and 3 mol %) and incubated in either phosphate buffered saline (PBS; 150 mM NaCl in 20 mM sodium phosphate at pH 7.4) or PBS containing bovine serum albumin (BSA; Sigma) in concentrations ranging from 1 to 5 wt %. Samples were transferred under the respective aqueous environment into new microwells at predetermined time intervals, and the surface-associated radioactivity was measured immediately following transfer. Stability assays at temperatures above 23 °C were performed while keeping the microwells in a thermostated humidity-saturated oven. All samples were prepared in quadruplicate. Atomic Force Microscopy. Images were obtained using a Nanoscope IIIA instrument from Digital Instruments fitted with an E-head scanner. Contact AFM cantilevers with a nominal spring constant 0.05 N/m and featuring a pyramidal Si3N4 oxidesharpened tip (0.6 µm) were purchased from Park Scientific Instruments. To enable AFM imaging in water of fused lipid monolayers, a sample holder was designed for the piezoelectric stage which accommodated vesicle fusion, rinsing, and image analysis without the need for sample transfer; thus avoiding the risk of exposure to air. Circular hydrophilic wells (1 mm deep, 8 mm i.d., 10 mm o.d.) were produced on a microscope slide by injection molding using medical grade silicon rubber (Technical Products Inc.) and a 16-well chamber slide template (Nunc) followed by a 7 day cure at 37 °C in a humid environment. The wells were separated from each other with a glass cutter and a 4 × 4 mm OTS/glass sample was glued to the bottom of each well with silicone rubber. All AFM samples of supported lipid monolayers were prepared in duplicate following the vesicle fusion and substrate rinsing protocols described above. Data were acquired 5 min, 40 min, and 4 h after vesicle fusion in an immersed contact regime, using the constant height mode, with a horizontal resolution set at 512 × 512 pixels and a scanning line speed of 4 Hz. The probe force was maintained at 1-5 nN throughout the imaging experiment. Characteristically, three different areas, separated by at least 10 µm, were imaged. The average roughness (Ra) was calculated using the system software and was the average of triplicate measurements.
Membrane-Mimetic Monolayers on Solid Supports
Results and Discussion Lipopeptide Synthesis. Two thiol-terminated peptides H-Gly-Arg-Gly-Asp-Tyr-NHCH2CH2SH (GRGDYNHCH2CH2SH) and H-Trp-Gln-Pro-Pro-Arg-Ala-Arg-IleGly-Tyr-NHCH2CH2SH (WQPPRARIGY-NHCH2CH2SH), integrin receptor and heparan sulfate binding sequences, respectively, were synthesized for conjugation studies. Respective molecular weights were 625.7 and 1302.6 g/mol as determined by MALDI mass spectrometry. Although coupling yields after lipid conjugation were >80% by TLC, overall yields were 31% and 48%, respectively, after purification. Rf values in CMAW 4/2/0.05/0.4 v/v/v/v were as follows: DPPE-MPB (0.82 ( 0.04), LP1 (0.25 ( 0.03), LP2 (0.36 ( 0.03). The presence of a free N-terminus on the conjugate was verified by positive reaction with ninhydrin on the TLC plate. The same spot turned blue upon reaction with molybdenum blue, indicating the presence of the conjugated phospholipid moiety, and the thiol-active Ellman’s reagent did not react with either one of the conjugates, confirming the proper coupling orientation of the two peptides. Additional confirmation of the structures of LP1 and LP2 was provided by MALDI mass spectrometry which yielded the expected molecular weights of 1624.5 (MLP1 + 2Na) and 2279.0 g/mol (MLP2 + Na), respectively. Both lipopeptides demonstrated strong absorbance in the UV spectra, and concentration calibration curves were determined at 220 nm with molar extinction coefficients () of 14828 M-1 cm-1 (LP1 e 100 µM) and 38921 M-1 cm-1 (LP2 e 50 µM), respectively. Lipopeptide Radiolabeling and Formation of Multicomponent Vesicles. Both LP1 and LP2 were characterized after cold labeling with NaI127 in order to ensure that oxidative iodination did not affect the integrity of the lipopeptide. Notably, side products were not detected on TLC and Rf values of the conjugates were unchanged after iodination. However, several peaks were noted on MALDI mass spectrometry consistent with the generation of several oxidized species with molecular weights close to that of the starting materials. For example, the MALDI mass spectrum of the iodinated LP1 demonstrated peaks at 1723 g/mol, which can be attributed to the monoiodinated sulfoxide M + O + I, as well as at 1617 and 1633 g/mol, which likely represent M + Na + O (sulfoxide) and M + Na + 2O (sulfone). In addition, a peak was also observed at 1649 g/mol which we suspect corresponds to M + H + 3O, the hydrolyzed sulfone following ring opening hydrolysis of the maleimide group. Similar results were noted following the iodination of LP2. Thus, while subtle changes may occur following oxidative iodination, the peptide and phospholipid moieties remain covalently linked and, as such, confirm the feasibility of preparing radioactive conjugate probes using an oxidative iodination protocol. Characteristically, the specific activity following radioiodination with NaI125 ranged between 2 and 10 CPM/ fmol of lipopeptide. Mixed micelles of LP1/DPPC containing 0.1, 0.5, 1, 2, and 3 mol % LP1 were prepared as described. Radiometric titration indicated that the true lipopeptide contents of these formulations were 0.13, 0.43, 0.84, 1.77 and 3.02 mol %, respectively. In prior reports, extrusion has been our preferred method for the preparation of well-defined unilamellar liposomes; however, dialysis was chosen in this series of investigations in order to minimize exposure to radioactivity during lipopeptide handling. Others have confirmed that dialysis of OG micelles yield liposomes which are similar in both size (180-240 nm) and structure (uni- and oligolamellar) to those produced by extrusion through a membrane of 200 nm porosity.19,20 Nonetheless, due to the tendency of zwitterionic vesicles to aggregate
Langmuir, Vol. 15, No. 11, 1999 3869
Figure 2. Kinetic study of vesicle fusion to OTS/substrates was performed using a 130 µM suspension of LP1/DPPC (1:99 mol %) liposomes in 150 mM NaCl at 50 °C.
and undergo changes in diameter upon storage, all dialyzed vesicles were used within 1 day of storage at room temperature. Kinetics of Mixed Lipopeptide/Phospholipid Vesicle Fusion and Film Formation: Analysis by Correlative Radiotitration and Atomic Force Microscopy. A kinetic study of vesicle fusion to OTS/substrates was performed using a suspension of LP1:DPPC (1:99 mol %) liposomes. Fusion in the presence of 150 mM NaCl at 50 °C was assessed after incubation periods ranging between 2 min and 18 h. Experimental data was regressed to a model of lipid film formation in order to estimate values for n, A1, and APC. As demonstrated in Figure 2, the formation of the expected monolayer reached completion after 4-6 h of vesicle fusion. Of note, after longer fusion times (18-20 h), a 3-SAM arrangement was observed, consistent with the tendency of phospholipids to form multilayered vesicles, tubes,21 or stacked planar surfaces22,23 by osmotic self-assembly. While previous authors have reported that supported multilayered phosphatidylcholine lipid assemblies peel upon exposure to a large excess of water,24,25 this was not observed in our system. The supported lipid trilayer containing 1 mol % lipopeptide was resistant to extensive rinsing. Of note, the addition of up to 50 mol % cholesterol to this system did not alter the kinetics of monolayer formation (data not shown). The kinetics of supported monolayer formation were characterized by parallel AFM studies. Fusion was terminated at 5 min, 40 min, and 4 h, surfaces imaged, and associated roughness (Ra) and film thickness analyses (19) Mimms, L. T.; Zampighi, G.; Nozaki, Y.; Tanford, C.; Reynolds, J. A. Biochemistry 1981, 20, 833-40. (20) New, R. R. C. Liposomes: A Practical Approach; Oxford University Press: New York, 1992. (21) Lasic, D. D. J. Colloid Interf. Sci. 1990, 140, 302-304. (22) Lipowsky, R. Curr. Opin. Struct. Biol. 1995, 5, 531-540. (23) Chernomordik, L.; Kozlov, M. M.; Zimmerberg, J. J. Membrane Biol. 1995, 146, 1-14. (24) Netz, R. R.; Lipowsky, R. Phys. Rev. Lett. 1993, 71, 3596-3599. (25) Hartung, J.; Helfrich, W.; Klo¨sgen, B. Biophys. Chem. 1994, 49, 77-81.
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Figure 3. Wet contact mode AFM images of LP1/DPPC (1:99 mol %) self-assembled on alkylated glass coverslips after (A, B) 5 min, (C, D) 40 min, and (D, E) 4 h of vesicle fusion at 50 °C in 150 mM NaCl, 20 mM sodium phosphate (pH 7.4). Images (A; Ra 2.8 Å, 2 × 2 µm2), (C; Ra 2.8 Å, 5 × 5 µm2), and (E; Ra 1.8 Å, 5 × 5 µm2) represent the top view of the SAM along with the corresponding roughness analysis, while the respective cross sections and average line roughness are shown in images (B; Ra 2.8 Å), (D; Ra 2.25 Å), and (F; Ra 1.4 Å).
determined (Figure 3). All surfaces were void of contaminants, including residual liposomes which would have confounded the interpretation of the radiometric data presented above. Thus, AFM is in agreement with our observations, in this and other studies, that the rinse method developed herein is exhaustive.11 Of greater
significance is the evolution of film morphology and topography during the 4 h observation period. With increased surface roughness (Figure 3A; Ra ) 2.8 Å over 2 × 2 µm2), 5 min samples exhibited a “blurred” appearance compared to the topography of the underlying OTS layer (Ra ) 1.5 Å over 10 × 10 µm2). After a 40 min fusion
Membrane-Mimetic Monolayers on Solid Supports
Langmuir, Vol. 15, No. 11, 1999 3871
Figure 4. Surface-associated radioactivity as a function of LP1 (A) or LP2 (B) lipopeptide content in the DPPC supported monolayer. Modeled molecular areas for LP1, LP2, and DPPC are 173, 171, and 70 Å2, respectively. Under these conditions, the model fits the experimental data for a lipid monolayer (n ) 1) on OTS. Based upon the determined model parameters, these data may also be expressed as fmol of LP1 (C) or LP2 (D) per cm2 of equivalent monolayer.
period, samples demonstrated a high density of serpentine surface defects; consistent with incomplete monolayer formation. While surface defect depth was typically 10 Å and the average area surface roughness was not significantly altered compared with the 5 min time point (Figure 3C; Ra ) 2.8 Å over 5 × 5 µm2); average surface roughness between observed defects had decreased with respect to that noted on 5 min samples (Figure 3D, Ra ) 2.25 Å over 6.5 µm line, vs Figure 3B, Ra ) 2.8 Å over 2 µm). Due to greater phospholipid surface coverage, 40 min OTS samples were likely less hydrophobic than those samples investigated earlier in the fusion process. We believe that this phenomenon is at the origin of the blurred images obtained at earlier fusion times. Hydrophobic surfaces can be conveniently imaged by dry contact AFM in air; however, AFM in a wet environment is often easier to perform on hydrophilic surfaces. Radiometric kinetic studies suggest that only a partial phospholipid monolayer is formed after a fusion time of 40 min. Therefore, we speculate that the measured 10 Å depth of observed surface defects in effect spans the partial monolayer. After a 4 h fusion period, samples exhibited a very smooth surface, with an average roughness of 1.8 Å over 5 × 5 µm2 (Figure 3E). This was significantly smoother compared with those images obtained at 5 and 40 min timepoints, in agreement with the occurrence of a topography-smoothing molecular self-assembly process. Defects were rare but were once again characterized by depths of approximately 10 Å. Of note, optimal comparative analysis of area and line roughness among all samples would require fields of view and linear AFM examinations of identical dimension. Given the length of a DPPC molecule (ca. 20 to 24 Å above and below its Tm of 41 °C, respectively),13,20 a film thickness of 10 Å suggests that the self-assembled mole-
cules are tilted at an angle of 60° from the normal. This is inconsistent with a prior report from our group which documented the formation of a supported single component DPPC monolayer on OTS/glass with a measured depth of ca. 20 Å.11 Overall, literature values derived from surface plasmon resonance measurements for the thickness of supported DMPC and DPPC monolayers doped with 5 mol % biotin-DPPE have ranged from 16 to 36 Å.3,13 While alternate interpretations of our film thickness data include either intercalation of the DPPC and OTS alkyl chains or lateral spreading of the DPPC alkyl chains in order to maximize their interaction with the underlying OTS substrate, further investigations would be required to unambiguously interpret the basis for the observed film thickness. Nevertheless, the evolution of surface topography in association with our radiometric data strongly supports the formation of a complete phospholipid monolayer during a vesicle exposure period of 4-6 h. Generation of DPPC Membranes of Varying Lipopeptide Composition: Controlled Film Formation by Vesicle Fusion. To test the generality of the transfer of lipopeptide-containing molecular assemblies from liposomes to planar hydrophobic surfaces, mixed vesicles containing 0.1, 0.3, 1, 2, or 3 mol % of either radiolabeled LP1 or LP2 were prepared and fused with OTS/glass substrates at 50 °C for 6 h. After completion of the fusion step, wells were exhaustively rinsed and equilibrated in buffer and the surface-bound radioactivity was measured (Figure 4). It is assumed that the mole fraction of lipopeptide in the supported film is identical to that in the generated liposomes. Regression of these data to our model of film formation is depicted and allowed us to estimate values for n, A1, A2, and APC. A parameter sensitivity study showed that the best fit of the experimental data for the
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Figure 6. LP1/DPPC monolayer stability at 23 °C during prolonged incubation in 150 mM NaCl in 20 mM sodium phosphate (pH 7.4). Figure 5. Surface-associated radioactivity as a function of LP1 and LP2 lipopeptide content in a ternary system with DPPC. Modeled molecular areas for LP1, LP2, and DPPC are 173, 171, and 70 Å2, respectively, with a monolayer (n ) 1) providing the best fit for the experimental data.
LP1/DPPC binary system was obtained for n ) 0.97, APC ) 69.80 Å2/molecule, and A1 ) 173.45 Å2/molecule (r2 ) 0.994). The ranges tested were 0.5-3 for n, 30-80 Å2/ molecule for APC, and 50-200 Å2/molecule for A1. Likewise, the best fit for the LP2/DPPC system was obtained utilizing n ) 0.97, APC ) 69.80 Å2/molecule, and A2 ) 170.84 Å2/molecule (r2 ) 0.994). In both cases, the model was very sensitive to the input value for n but not very sensitive to the estimated molecular area of the lipopeptide due to its relatively low content in the supported film. To test the hypothesis that several lipopeptides could be incorporated in a controllable fashion into a supported phospholipid membrane, mixed DPPC liposomes doped with radiolabeled LP1 and LP2 in different ratios were prepared and fused to OTS/glass substrates. Experimental results and regressed data based upon the presented analytical model for film formation are presented (Figure 5). Multiple linear regression analysis revealed that utilization of n ) 0.97, APC ) 69.80 Å2/molecule, A1 ) 173.45 Å2/molecule, and A2 ) 170.84 Å2/molecule in eq 6 best modeled the experimental data. The calculated molecular area of 69.80 Å2/molecule for DPPC deserves comment. A temperature of 50 °C was chosen during the adsorption of lipopeptide-containing vesicles to the OTS monolayer in order to perform vesicle fusion above the main transition temperature of DPPC (41.5-42 °C). Under these conditions, DPPC was in a liquid expanded phase which is characteristically associated with a molecular area of about 60-70 Å2.26 For example, in compressibility studies using dilute multilamellar vesicles, Lis et al.27,28 observed a molecular area of 71.2 Å2 for DPPC at 50 °C in plain water. While this is consistent with our (26) Hauser, H.; Pascher, I.; Pearson, R. H.; Sundell, S. Biochim. Biophys. Acta 1981, 650, 21-51. (27) Lis, L. J.; McAlister, M.; Fuller, N.; Rand, R. P.; Parsegian, V. A. Biophys. J. 1982, 37, 657-666. (28) Lis, L. J.; McAlister, M.; Fuller, N.; Rand, R. P.; Parsegian, V. A. Biophys. J. 1982, 37, 667-672.
observations, it is likely that molecular packing of DPPC on the OTS/glass substrate varied with temperature. Following vesicle fusion at 50 °C and subsequent cooling to room temperature, DPPC passes through its main transition with a probable reduction of the cross sectional area of its hydrocarbon chains. In this regard, Wenzl et al.6 have measured a molecular area for DPPC of 36.545.1 Å2 at 25 °C after deposition on a DPPA monolayer. As such, we would predict that a reduction in PC molecular area would lead in our system to the development of lateral film tension, compensated by either a corresponding hydrocarbon chain tilt or the development of surface defects in the lipid film. As noted above, the presence of rare surface defects, as determined by AFM, is consistent with an increase in chain tilt. Overall, these data support the general use of vesicle fusion methodology for the controlled transfer of lipopeptide-containing DPPC membranes of varying composition onto hydrophobic substrates with the generation of biologically heterogeneous supported monolayers. Mixed Monolayer Stability in Phosphate Buffered Saline: Effect of Incubation Period, Temperature, Alkyl Chain Length, and Cholesterol Content. Stability data for LP1/DPPC films incubated in PBS at 23 °C are presented in Figure 6. We observed a reduction in surface-associated radioactivity in all samples during the first few days of incubation, regardless of their lipopeptide content. A stable plateau, however, was reached after the fourth day of equilibration with PBS. We believe that the initial fall in radioactivity reflects, in part, diffusion of residual iodide bound as a counterion to positive charges in the peptide moiety. Thereafter, little change was noted over the 32 day period during which the surface-bound radioactivity was monitored. The initial fall in radioactivity is somewhat greater in this set of experiments than that subsequently observed since the final optimization of rinsing and dialysis protocols, designed to remove unbound vesicles or residual free iodine, respectively, were not yet complete. It is also worth noting that the LP1 content in the supported DPPC monolayer is consistent with our previous experiments. The 0.13, 0.43, 0.84, 1.77, and 3.02 mol % LP1 in the DPPC matrix yielded final stable surface concentrations of 600 ( 80, 1700 ( 220,
Membrane-Mimetic Monolayers on Solid Supports
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Figure 7. (A) The effect of lipid alkyl chain length on monolayer stability was characterized by comparing membrane-mimetic surfaces formulated with 3 mol % LP1 in either DPPC or DMPC while stored in PBS at 23 and 37 °C. (B) Mixed monolayers (3 mol % LP1 in DPPC) containing cholesterol (33 or 50 mol %) reduced by 50% the loss of lipopeptide when incubated in PBS alone.
2500 ( 180, 4000 ( 1070, and 9700 ( 600 fmol of LP1 per square centimeter of equivalent monolayer (mean ( standard deviation, n ) 4). The effect of lipid alkyl chain length on monolayer stability was characterized by comparing membranemimetic surfaces formulated with 3 mol % LP1 in either DPPC or DMPC while stored in PBS at 23 and 37 °C (Figure 7A). The final number of stacked monolayers was found to be 1.3 for DPPC and 1 for DMPC, using the described mathematical model. Overall, at room temperature both systems exhibited remarkable stability over an extended period of time. With an increase in temperature to 37 °C, the DPPC and DMPC systems demonstrated lipopeptide losses of 18% and 30%, respectively. Further, the rate of lipopeptide loss was directly proportional to the final equilibrium surface concentration. In other words, the rate of surface depletion was greatest in those systems which exhibited the largest total losses. Thus, these data reveal that a temperature increase from 23 to 37 °C had a significant membrane destabilization effect. We speculate that the increased stability of DPPC with respect to DMPC can be attributed to differences in their respective main transition temperatures. A Tm of 41 °C for DPPC allowed this system to remain in its liquidcrystalline state throughout. The DMPC-based system crossed its Tm of 23 °C upon exposure to PBS at 37 °C, thus transitioning from a mixture of coexisting condensedliquid and expanded-liquid phases to a single homogeneous phase of liquid-expanded DMPC.27-29 The change in the (29) Raudino, A.; Zucarello, F.; LaRosa, C.; Buenni, G. J. Phys. Chem. 1990, 94, 4217-4223.
Figure 8. The influence of BSA on monolayer stability was characterized at 37 °C by determining the change in lipopeptide surface concentration as a function BSA concentration (0-5 gm %) in PBS. Supported monolayers were formulated with 3 mol % LP1 in DPPC in the presence of either 0 (A), 33 (B), or 50 mol % (C) cholesterol. The presence of cholesterol at either 33 or 50 mol % had relatively little effect on either the rate or magnitude of lipopeptide loss when monolayers were incubated in the presence of BSA.
phase state of the lipid monolayer does not establish a mechanism for lipopeptide loss. Nonetheless, several investigators have documented spontaneous thermodynamic rearrangement of phosphatidylcholine molecules in favor of structures with smaller radii of curvature upon input of thermal,20,22 chemical,30 or mechanical energy.31-34 (30) Batzri, S.; Korn, E. D. Biophys. Acta 1973, 298, 1015-1019. (31) Olson, F.; Hunt, C. A.; Szoka, F. C.; Vail, W. J.; Papahadjopolous, D. Biochim. Biophys. Acta 1979, 557, 9-23. (32) Mayer, L. D.; Hope, M. J.; Cullis, P. R. Biochim. Biophys. Acta 1986, 858, 161-168.
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Therefore, we speculate that with an increase in temperature vesicles may be generated and subsequently lost from the lipid monolayer. It is notable that addition to the lipid monolayer of either 33 or 50 mol % of cholesterol reduced by 50% the loss of lipopeptide (Figure 7B). Mixed Monolayer Stability in Phosphate Buffered Saline: Effect of Solution Phase BSA and Monolayer Associated Cholesterol Content. Lipopeptide/DPPCsupported planar monolayers were formulated with 3 mol % LP1 and incubated at 37 °C in PBS with varying concentrations of BSA (Figure 8A). The magnitude and rate of lipopeptide depletion from the planar membrane increases with the solution concentration of BSA and approaches 40% at the highest protein concentration. Notably, all substrates achieve a new stable surface concentration of lipopeptide after an 8 day incubation period. The influence of increasing protein concentration on lipopeptide loss is not linear, with the largest effect observed for BSA concentrations up to 2 gm % and relatively little change thereafter. The ability of monolayer cholesterol content to reduce lipopeptide loss was characterized. Vesicles containing 3 mol % LP1 and either 33 or 50 mol % cholesterol were produced and fused onto OTS/glass, as described above. Model surfaces were subsequently exposed to PBS with BSA. However, relatively little effect on either the rate or magnitude of lipopeptide loss was observed (Figure 8B,C). Further experimentation, beyond the scope of this report, will be required to determine the mechanism of lipopeptide loss. Nonetheless, we postulate that the loss likely occurs as a consequence of both protein-protein and protein-membrane interactions. In this regard, it is conceivable that binding of BSA to surface-associated peptide pendent groups could result in the desorption of the lipopeptide component and the reestablishment of a new surface equilibrium concentration. Protein-membrane interactions may be of even greater significance. In the late 1960s, Brash and Lyman reported that surfaceadsorbed DPPC was capable of adsorbing human serum albumin.1 Although recent investigations by Malmsteen and Lassen35 have suggested that BSA is adsorbed only weakly onto DPPC surfaces, it is difficult to relate these results to the present study since both rate and binding constants were not determined. However, it is also probable that some of the BSA molecules are incorporated into the lipid layer. For example, Cornell et al.36 found that BSA is incorporated into lipid monolayers of palmi(33) MacDonald, R. C.; MacDonald, R. I.; Menco, B. P. M.; Takashita, K.; Subbarao, N. K.; Hu, L.-R. Biochem. Biophys. Acta 1991, 1061, 297303. (34) Clerc, S. G.; Thompson, T. E. Biophys. J. 1994, 67, 475-477. (35) Malmsten, M.; Lassen, B. Colloid Surf. B:Biointerfaces 1995, 4, 173-184. (36) Cornell, D. G.; Patterson, D. L.; Hoban, N. J. Colloid Interface Sci. 1990, 140, 428.
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toyloleoylphaspahtidylcholine (POPC) and palmitoyloleoylphopshatidylglycerol. Recent investigations by Cho et al.37 have documented that at the air/water interface the extent of BSA incorporation into phosphatidylcholine monolayers is largely dependent upon the lipid molecular packing density. That is, protein adsorption was enhanced at dilute lipid monolayer densities (64-107 Å2/molecule) and reduced in dense lipid monolayers (33 Å2/molecule). Similarly, Terrettaz and colleagues38 found that nonspecific binding of streptavidin to POPC/thiolipid-supported membranes not containing biotin strongly depended on the structure of the lipid monolayer, being highest in the fluid expanded and gas analogue states. Negligible binding was observed to closed packed lipid layers. In the present study, the planar surface onto which the monolayer is adsorbed is of finite area. Thus, the intercalation of BSA with the monolayer could well displace both DPPC and lipopeptide constituents into the solution phase, as a consequence of either edge effects or, perhaps, the induction of planar-to-vesicle transitions with liposome desorption. We believe that our observations bear relevance for both the design of supported membrane-mimetic systems which operate in an aqueous biological milieu, as well as the optimization of site-specific drug delivery by colloidal lipid-based carriers in contact with blood. Limitations of the latter have been attributed to adsorption of albumin and other plasma proteins onto the liposome surface thereby facilitating particle opsonization and the consequent nonspecific uptake by cellular components of the reticuloendothelial system. The data presented herein suggest that targeted liposomal drug delivery may be further limited by surface loss of the targeting conjugate mediated by interaction of the vesicles with serum proteins. As such, these effects may be reduced, but not likely eliminated, by greater control of molecular packing density or by changes in other local properties which alter the electrostatic or steric microenvironment in the region of the pendent group. Ultimately, we suspect that stabilization of membrane-mimetic structures by the generation of polymerized assemblies, as reported elsewhere,8,9 will be required for the introduction of these systems into their promised areas of application. Acknowledgment. Supported by the Whitaker Foundation and the NIH (HL56819). E.L.C. is a ClinicianScientist of the American Heart Association. The authors acknowledge Professor Lawrence A. Bottomley at the Georgia Institute of Technology, Department of Chemistry, for assistance with AFM imaging. LA980990Q (37) Cho, D.; Narsimhan, G.; Franses, E. I. Langmuir 1997, 13, 47104715. (38) Terrettaz, S.; Stora, T.; Duschl, C.; Vogel, H. Langmuir 1993, 9, 1361-1369.
