Planar Supported Bilayer Polymers Formed from Bis-Diene Lipids by

Biomacromolecules 2003, 4, 841-849

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Planar Supported Bilayer Polymers Formed from Bis-Diene Lipids by Langmuir-Blodgett Deposition and UV Irradiation John C. Conboy,† Sanchao Liu, David F. O’Brien, and S. Scott Saavedra* Department of Chemistry, University of Arizona, Tucson, Arizona 85721-0041 Received July 17, 2002; Revised Manuscript Received March 26, 2003

Substrate-supported lipid bilayers have been prepared from bis-diene functionalized phosphorylcholine (PC) lipids and polymerized by UV irradiation. The overall bilayer structure is largely preserved upon removal from water, although significant loss of material occurs from the upper leaflet of the bilayer, likely due to desorption at the air/water interface. The morphology and surface structure of the bilayer, as observed by AFM, indicate a substantially different arrangement of the lipids in the hydrated and dehydrated states, presumably due to the loss of water from the near surface region. These changes have been correlated with infrared spectral shifts sensitive to the conformation of the hydrocarbon chains. Protein adsorption studies show that rehydrated, polymerized bilayers retain a degree of resistance to BSA adsorption intermediate between model hydrophobic and fluid PC lipid bilayer surfaces. The degree of protein adsorption is correlated with desorption of material from the upper leaflet of the bilayer upon drying, which produces voids at which hydrophobically driven protein adsorption occurs. Introduction At the transducer/sample interface of an ideal receptorbased biosensor, specific binding of an analyte to an immobilized receptor can be detected without interference from the numerous other nontarget molecules present in the sample matrix. To realize this situation, two major challenges must be overcome: (i) To preserve receptor bioactivity, it must be immobilized at the transducer surface in an environment that both preserves its native conformation and correctly orients its binding site toward the sample matrix.1,2 (ii) In most optical and electrochemical biosensors, the physical transducer is an oxide or noble metal surface. These surfaces promote nonspecific, sometimes irreversible adsorption of proteins, which can interfere with the specific interaction desired in a protein immobilization scheme and, when the sample matrix contains nontarget proteins, cause “fouling” of the transducer surface.3,4 An attractive strategy that addresses both challenges is to create a mimic of a cell membrane,5,6 which is a structure designed by nature to both provide a suitable environment for bioactive presentation of receptors and to be highly resistant to nonspecific adsorption of soluble, nontarget proteins.7 The inherent resistance of cell membranes to protein adsorption and platelet adhesion is at least partially attributable to the hydrated phosphorylcholine (PC) moiety, which is the most prevalent lipid headgroup in the outer leaflet of most animal cell membranes.7,8 A number of other strategies to control protein adsorption have been extensively studied (see ref 4 for reviews). * To whom correspondence should be addressed. Phone: (520) 6219761. Fax: (520) 621-8407. E-mail: [email protected]. † Current address: Department of Chemistry, University of Utah, 315 S. 1400 E., Salt Lake City, UT 84112.

Planar supported lipid bilayers (PSLBs) have been extensively studied as models of the lamellar assembly of lipids and associated protein receptors in a cell membrane.9-15 PSLBs are commonly prepared using either the LangmuirBlodgett Schaefer (LBS) or vesicle fusion methods.9,10 The major advantage of LBS deposition is that the packing density and lipid composition of the film can be precisely controlled. With the appropriate choice of composition and deposition conditions, lateral organization and membrane asymmetry can be specified. In contrast, vesicle fusion is restricted to deposition of fluid-phase lipids and cannot be used to create asymmetric bilayers; however, technically, it is a much simpler method to implement than LBS deposition. Regardless of the deposition method, in a fluid PSLB formed from naturally occurring lipids, the molecules are only weakly associated with each other and the underlying substrate surface.14 These structures consequently lack the chemical, mechanical, and/or thermal stability required for technological implementation (e.g., as a nonfouling coating in a reusable biosensor). Improving the structural stability of supported lipid films while maintaining their inherent biocompatibility has therefore been a focus of research efforts since the early 1980s.4b,16,17 One strategy to create robust lipid assemblies is covalent polymerization of the lipid monomers. Studies of bilayer vesicles and mono- and multilamellar films have demonstrated that lipid polymerization can enhance the structural stability and decrease the solubility of these supramolecular structures.16-19 Much of the early work in this area was focused on lipids bearing diacetylene-functionalized chains.16,18 However, this moiety is not an ideal candidate to stabilize a PSLB because its polymerization is highly sensitive to reactive group packing.20 The topotactic nature of the reaction dictates that these lipids can only be polymerized in the gel phase, and a low degree

10.1021/bm0256193 CCC: $25.00 © 2003 American Chemical Society Published on Web 04/24/2003

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alternate preparation and polymerization methods. LBS deposition and UV photopolymerization were used to create cross-linked PSLBs using two lipids bearing polymerizable groups at different positions in the acyl chains (this variable is known to affect polymerizations in bilayer vesicles).18 The films were characterized using ellipsometry, contact angle measurements, atomic force microscopy, UV absorbance and FT-IR/ATR spectroscopies, and X-ray photoelectron spectroscopy. In addition, total internal reflection fluorescence spectroscopy was used to examine protein adsorption properties. The results of these studies demonstrate a correlation between extent of polymerization, air stability, defect density, and resistance to nonspecific protein adsorption. Experimental Section

Figure 1. Structures of the polymerizable lipids bis-SorbPC and bisDenPC.

of conversion is a typical result.18 Furthermore, because bisdiacetylene lipids exist in the solid-analogous phase at room temperature, PSLB self-assembly by vesicle fusion is precluded.21 In contrast, lipids with alkene-functionalized chains (e.g., acryloyl, methacryloyl, sorbyl, and dienoyl) can be polymerized in either the gel or liquid crystalline phases to a higher degree of conversion than bis-diacetylene lipids.18a In early work, Regen and co-workers17 used UV light to polymerize mono- and diacrylate-functionalized lipids adsorbed on poly(ethylene), forming supported lipid films of near monolayer thickness. Their water contact data were higher (less hydrophilic) than expected for a substrate coated with a wellpacked array of PC groups,22,23 suggesting incomplete coverage and/or significant film disorder allowed the overlying water droplet to come into contact with exposed lipids chains. More recently, Chaikof and co-workers24 used vesicle fusion to form a mono-acryloyl lipid monolayer on an alkylsilane SAM. Reaction produced linear polymers in the upper leaflet of the hybrid bilayer. Although enhanced stability during extended incubation in water was observed, significant lipid desorption occurred when the assembly was exposed to a surfactant. In contrast, polymerization of bissubstituted lipids produces cross-linked bilayers with greatly enhanced stability relative to linear polymerizations. For example, cross-linked unilamellar vesicles are insoluble in surfactant solutions and organic solvents.18,25 The position of the polymerizable moieties in the lipid chains affects their reactivity as well as the physicochemical properties of the (poly)lipid. In a recent communication and subsequent papers,22 we described redox-initiated radical polymerization of a selfassembled PSLB, composed of bis-sorbyl phosphatidylcholine (bis-SorbPC, Figure 1),26 that was stable to transfer across the air/water interface, insoluble in surfactants and organic solvents, and as resistant to nonspecific protein adsorption as a fluid PSLB. In this study, we focus on

Materials. Bis-SorbPC was prepared by a modification of the procedure reported by Lamparski et al.27 The synthesis of bis-dienoyl phosphatidylcholine (bis-DenPC) was adapted from that reported by Dorn et al.28 The lipid structure was established by 1H NMR and HRMS. In addition, the purity was confirmed by the presence of only one spot on TLC. 1-Palmitoyl-2-oleoylphosphatidylcholine (POPC) was purchased from Avanti Polar lipids for use as a control in protein adsorption studies. The water used in the polymerization and protein adsorption experiments, hereafter referred to as deionized water, was obtained from a Barnstead Nanopure system with a measured resistivity of 17.9 MΩ cm and total organic content specified as less than 10 ppb. Silicon wafer substrates were purchased from Wacker and had a 20 ( 5.6 Å thick native oxide layer after cleaning (described below). The substrates for the protein adsorption experiments were fused silica slides (4 × 2.5 cm2, Dynasil). Si ATR crystals (Harrick Scientific) were used as substrates for FT-IR/ATR spectroscopy. Fluorescein-labeled bovine serum albumin (FITC-BSA) with a labeling ratio of 11.2:1 was purchased from Sigma and used without any further purification. Fluorescein-labeled dextran (10 000 MW, 2.9:1 labeling ratio) was purchased from Molecular Probes. Preparation of Polymerized Bis-SorbPC Bilayers. PSLBs were prepared on Si wafers, fused silica slides, and Si ATR crystals. All substrates were treated as follows: They were first sonicated in 50% isopropyl alcohol/50% water (v/v), rinsed in deionized water, treated with 30% H2O2/70% concentrated H2SO4 for 30 min, and then rinsed repeatedly in deionized water. The cleaned substrates were then sonicated in a 0.1 M solution of AlCl3 for 30 min, rinsed with deionized water, sonicated for 15 min in deionized water, and then rinsed again and dried under flowing N2. This procedure produced hydrophilic surfaces with a sessile water contact angle of 9.6 ( 3.5°. Treatment of the substrates with AlCl3 was found to result in better transfer of lipid films, possibly by binding and stabilizing the lipid layer.29 PSLBs of bis-SorbPC and bis-DenPC were prepared by LBS deposition.11a,13 A phospholipid monolayer was spread on a Nima model 611D LB trough using benzene as the spreading solvent and deionized water as the subphase. The pressure-area isotherm for a Langmuir monolayer of bis-

Planar Supported Bilayer Polymers

Figure 2. Pressure-area isotherm for bis-SorbPC at the air/water interface.

SorbPC is shown in Figure 2. The isotherm for bis-DenPC (not shown) was similar to that reported by Grainger et al.30 Film depositions were performed at a surface pressure of 35 mN/m, corresponding to approximately 60 Å2/molecule. The inner leaflet of the bilayer was deposited by withdrawing the substrate from the subphase at a rate of 10 mm/min. Transfer ratios of 98 ( 5% were obtained. The second leaflet of the bilayer was deposited by passing the substrate coated with the previously deposited monolayer horizontally through the air-water interface at constant pressure. After formation, the unpolymerized bilayer was maintained in an aqueous environment at all times. All depositions were carried out at 25 °C. Other surface pressures and subphase compositions were explored, but the conditions stated above produced the most stable and reproducible films. Polymerization of the lipid bilayer was performed in situ by exposure to UV radiation from a low-pressure mercury pen lamp (Fisher Scientific) with a rated intensity of 4500 µW/cm2 at 254 nm. A 230 nm long-pass filter (UG5, Schott Glass) was used to remove the shorter wavelength UV that was found to cause degradation of the lipid film. The lamp was held 7.5 cm from the substrates which were illuminated for 4 min. Prior to initiating polymerization, the solution containing the PSLB was purged with air for 30 min. After UV irradiation, the (poly)PSLB was removed from solution, rinsed several times with deionized water, and dried with a stream of nitrogen. Kinetic Measurements. Polymerization rate experiments were performed using a Spectral Instruments 440 UV-vis spectrometer. PSLBs were deposited on one side of four quartz slides which were then assembled into a stack, separated by 2 mm thick, 25 mm OD Viton O-rings, and mounted in a fluid cell without exposing the bilayers to air. This arrangement allowed absorbance measurements to be made on PSLBs in a transmission geometry with adequate sensitivity. Absorbance spectra were collected after exposure to UV illumination at various time intervals. The kinetic data were retrieved from the spectra by integrating the absorbance peak at 260 nm after baseline correction. Ellipsometry and Contact Angle Measurements. Thickness measurements were performed on dried PSLBs depos-

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ited on Si substrates using a Rudolph Manual Nulling Ellipsometer. Measurements were made using the 632.8 nm line of a HeNe laser at an incident angle 70° from the surface normal, assuming a uniform refractive index profile of 1.457 for the PSLB. Static water contact angle measurements were performed with a water-cooled CCD camera (Princeton Instruments Model 512TK) and the contact angle retrieved via imaging analysis software (Scion Image).31 X-ray Photoelectron Spectroscopy (XPS). XPS was used to examine the elemental composition of (poly)PSLBs. Measurements were performed on bilayers deposited on silicon wafers using a Kratos 165 Ultra Imaging XPS equipped with a 165 mm mean radius hemispherical analyzer and an eight channeltron detection system. The base pressure in the analyzer chamber was ca. 5 × 10-9 Torr. X-rays from an Al ΚR source at 1486.6 eV were used for excitation. Electrons were collected in the constant analyzer energy (CAE) mode with a pass energy of 50 keV. Integration times were 0.25 s, co-added four times, for a total of 1.0 s at an interval of 0.1 eV. The areas under the XPS peaks were determined by numerical integration after baseline correction. Relative peak area ratios were calculated using published photoionization cross-sections32 after accounting for the transmission properties of the analyzer. Atomic Force Microscopy (AFM). The surface morphology of (poly)PSLBs was examined using atomic force microscopy (AFM) performed in tapping mode on a Nanoscope III (Digital Instruments, Santa Barbara, CA). Oxide sharpened silicon nitride tips were used for studies of samples both in air and immersed in water. In the latter case, samples were mounted in the standard Digital Instruments solution cell and allowed to equilibrate under deionized water for 0.5-1.5 h before image acquisition was initiated, using contact-cantilevers in a tapping mode configuration. To minimize distortion by contact forces between the tip and the sample, the minimum force required to obtain optimum resolution was used. The driving amplitude applied was the automated instrument default, between 100 and 400 mV. After the tip was engaged and imaging commenced, the setpoint value was increased until contact with the surface was lost and then incrementally decreased until optimal resolution was achieved. Setpoint values scaled with drive amplitude values and were generally between 0.2 and 1.2 V, which usually resulted in “softer” imaging than the default engagement settings. The samples were not altered by the AFM measurement, as noted by the invariance of successive AFM scans. The images presented below are representative of scans from different locations on the sample, different samples, and using different tips to image the surfaces. FT-IR Spectroscopy. FT-IR spectra of the C-H stretching region of PSLBs deposited on the upper face of a Si ATR crystal were obtained on a Nicolet 550 spectrometer equipped with a Harrick Scientific ATR accessory. Spectra of hydrated bilayers were obtained by placing a water layer on the surface of the (poly)PSLB-coated crystal. The spectrometer and sample compartment were purged with dry air. The spectra collected were an average of 128 scans at 1.0 cm-1 resolution over a frequency range of 4000-400 cm-1. Background spectra of the bare ATR crystal were

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recorded in dry air and with a 2 mm thick water layer on the upper surface of the crystal. TIRF Spectroscopy. Protein adsorption studies were performed by total internal reflection fluorescence (TIRF) spectroscopy. The optical arrangement consists of two rightangle quartz prisms mounted in a TIRF liquid flow cell.33 One prism is used to couple the excitation light at 488 nm from an air-cooled Ar+ laser into a fused silica slide mounted in the cell. After propagating the light through the slide by total internal reflection, the other prism is used to outcouple the excitation, thereby reducing scattered light in the cell volume. Index matching fluid (n ) 1.463, Cargille) was used to make the incoupling and outcoupling processes more efficient. Increasing concentrations of FITC-labeled BSA dissolved in buffer (50 mM phosphate, pH 7.2, containing 100 mM NaCl) were injected into the flow cell and allowed to equilibrate with the surface for 30 min prior to each measurement, which was determined experimentally to be a sufficient time for a steady-state response to be observed. The flow cell was mounted on a Nikon Diaphot inverted microscope. Fluorescence emission was back-collected through the quartz slide with a 4× objective, optically filtered, and detected with a photomultiplier tube. The excitation light was modulated at a frequency of 2.5 kHz, and phase sensitive detection was used to retrieve the fluorescence intensity. The experiment was interfaced to a PC for data collection. All experiments were performed at 25 °C.33 A Langmuir model was used to extract the apparent equilibrium association constant of BSA and the surface under investigation. The form of the Langmuir isotherm used was [BSA]Ka )

F Fmax - F

(1)

where F is the measured fluorescence, [BSA] is the bulk concentration of BSA injected into the flow cell, Ka is the equilibrium association constant, and Fmax is the maximum fluorescence at surface coverage saturation. Both Ka and Fmax were determined by a nonlinear least-squares analysis of the experimental data. A modified form of a numerical quantitation method33,34 was used to determine the relative surface coverage of adsorbed BSA from the fluorescence data. Calibration was performed by measuring the fluorescence from standard solutions of FITC-labeled dextran injected into the flow cell. By taking the ratio of the surface to solution phase total fluorescence, the surface concentration of adsorbed BSA can be determined by the following: φsf2ΓBSA Fs ) Fb φblkcdextrandp

(2)

where Fs is the fluorescence detected from the surface bound BSA, Fb is the fluorescence detected from the dissolved FITC-dextran, φsf and φblk are the quantum yields of FITC at the surface and in the bulk respectively, ΓBSA is the surface concentration of BSA, cdextran is the bulk concentration of FITC-dextran, and dp is the characteristic evanescent penetra-

tion depth. The latter quantity is given by dp )

λ 2π(n1 sin θ - n22)1/2 2

2

(3)

where n1 and n2 are the refractive indices of quartz substrate and water, respectively, θ is the incident angle, and λ is the wavelength of the incident light. It was assumed (a) that the quantum yield of FITC bound to BSA is equal to that bound to dextran, i.e., φsf/φblk ∼ 1 and (b) that FITC-dextran does not adsorb to the surface. Support for the latter is provided by the observation that flushing the TIRF flow cell with buffer eliminated the signal due to the FITC-dextran that previously occupied the flow cell volume; however, this does not eliminate the possibility that some weak, reversible adsorption occurred. Under these assumptions, a plot of the bulk concentration of FITC-dextran standards versus the fluorescence intensity yields the proportionality constant necessary to calculate the surface coverage of BSA. The calibration procedure also allows the adsorption isotherms measured on various surfaces to be normalized. Here all surface fluorescence data were normalized with reference to the fluorescence intensity measured after injecting an 18.6 µM FITC-dextran solution into the TIRF flow cell. For unpolymerized PSLBs, after FITC-BSA adsorption was measured, the flow cell was “washed” with a 1% solution of Triton X-100 to remove the lipid bilayer and adsorbed protein films. Calibration with FITC-dextran standard solutions was carried out thereafter. Because (poly)PSLBs could not be removed with Triton X-100, calibration was carried out with the BSA/PSLB films intact. In these cases, the protein film fluorescence was subtracted from that measured when an 18.6 µM FITC-dextran solution occupied the evanescent volume above the protein-coated lipid film. Results and Discussion Polymerization Kinetics. UV-vis absorbance spectroscopy was used to characterize the rate of polymerization of bis-SorbPC PSLBs. The bis-SorbPC monomer has an absorption maximum at 260 nm.35 By monitoring the depletion of the monomer absorbance as a function of the irradiation time, the apparent rate of polymerization was determined (Figure 3). Complete disappearance of the monomer was observed at times greater than 2 min, which was taken as near-quantitative polymerization (>95%) of the bilayer. The actual percent conversion cannot be determined because it is difficult to accurately measure a very small absorbance. The decay of the integrated monomer absorbance was fit to Abs(t) ) A exp(-kt)

(4)

where Abs(t) is the integrated absorbance at time t, A is the amplitude, and k is the rate of the monomer depletion process. A nonlinear least-squares analysis yielded an apparent k ) 18.9 ( 0.96 s. The polymerization process is believed to occur by a single mechanism as indicated by an adequate fit to a first-order model. These results are


Biomacromolecules, Vol. 4, No. 3, 2003 845 Table 1. Contact Angle and Ellipsometric Data for Polymerized Bis-SorbPC and Bis-DenPC Lipid Films Formed on Si Wafers

surface substrate (Si/SiO2) bis-SorbPC (monolayer) bis-SorbPC (bilayer) bis-DenPC (bilayer)

contact angle (degrees) 9.6 ( 3.5 60.4 ( 3.9 41.9 ( 3.1 65.2 ( 2.4

ellipsometric thickness (Å) 19.8 ( 5.6 (SiO2 layer thickness) 26.2 ( 3.1 48.4 ( 4.2 25.2 ( 4.9

Figure 3. Depletion of the UV absorbance of monomeric bis-SorbPC during irradiation of four PSLBs. The kinetic data were retrieved from the spectra by integrating the absorbance peak at 260 nm, after baseline correction, and normalized to the measurement made before exposure to the mercury lamp. Inset: absorbance spectra before and after irradiation for 240 s.

consistent with prior studies of polymerized sorbyl lipid vesicles which showed that 1,4 addition was the predominate product.18b,35 Irradiation of bis-SorbPC bilayer films for more than 2 min was not found to alter the film structure or morphology as observed by AFM and ellipsometry (see below). Purging the solution with Ar for 30 min (rather than air) also did not produce any measurable differences, consistent with previous observations.35 However, irradiation times less than 2 min resulted in substantially reduced conversion of monomer to polymer, consistent with the kinetic data represented in Figure 3. As a result, all of the films discussed below were prepared in solutions purged with air and irradiated for 4 min, which ensured near-complete polymerization of the lipid bilayer. Physical and Chemical Characterization of PSLBs. In the absence of UV illumination to effect polymerization, a bis-SorbPC lipid bilayer produced by LBS deposition is completely desorbed from a silica support during transfer across the air/water interface (i.e., the ellipsometric thickness is negligible). This is consistent with the known instability of a fluid bilayer formed from nonpolymerizable lipids (e.g., POPC).14,22b Air stability was therefore used as a measure of successful preparation of a polymerized lipid film and was essential in order to proceed with further characterization of the films. Stabilization is presumably achieved because the macromolecular lipid adheres to the underlying substrate surface via polydentate, noncovalent interactions. Ellipsometry and sessile water contact angle data obtained for dried (poly)bis-SorbPC bilayers are summarized in Table 1. Also tabulated for comparison are data for bis-SorbPC monolayers polymerized under the same conditions. Assuming fully extended hydrocarbon chains packed at their van der Waals radii, the calculated thickness for a bis-SorbPC bilayer is in the range of 44-54 Å,36 depending on headgroup conformation. The measured thickness, 48 Å, is consistent with the calculated thickness and nearly twice that measured for the polymerized monolayer (26 Å).

Figure 4. XPS data for a bare Si wafer (a-c) and a polymerized bis-SorbPC bilayer deposited on a Si wafer (d-f). Spectra a and b are for the C1s region, b and e are for the Al2p region, and c and f are for the N1s region.

The sessile water contact angles on (poly)bis-SorbPC bilayer and monolayer surfaces are 42 and 60°, respectively. The latter value reflects the presence of the ester groups near the chain termini of bis-SorbPC (Figure 1). Thus, it is expected to be significantly less than the ca. 110° measured on monolayers composed of saturated alkyl chains.31 In the case of the bilayer, the measured contact angle is higher than the ca. 30° reported for films terminated uniformly with PC groups.22,23 This difference suggests some exposure of the hydrophobic core of the bilayer upon dehydration, possibly due to rearrangement of the lipid headgroups and/or desorption of low molecular weight oligomers. However, taken together, the ellipsometry and contact angle data provide strong evidence that polymerization of bis-SorbPC stabilizes the lipid bilayer such that its overall structure is largely preserved after removal from an aqueous environment. Dried (poly)lipid films were also analyzed using XPS. Figure 4 shows spectra of the N1s, C1s, and Al2p regions of an Al3+-treated Si wafer and a wafer coated with a dried (poly)bis-SorbPC bilayer. The presence of the N1s peak, with a binding energy of 402.3 eV, confirms the presence of a lipid film (Figure 4f). The carbon signals near 286 and 289 eV are also assigned to the lipid film (Figure 4d) and are

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Figure 5. AFM images of (a) a dried bis-SorbPC PSLB and (b) the same film immersed in water. Linescan (c) of a scratch etched into a bis-SorbPC PSLB to the underlining substrate by repeated, high force scanning with the AFM tip.

distinguishable from the major peak at ca. 284 eV (Figure 4, parts a and d); the latter signal is due to atmospheric contaminants adsorbed during the period between polymerization and XPS measurements. A carbon to nitrogen (C/N) elemental ratio of 42 ( 6.3:1 is measured for the polymer film. This composition is consistent with a calculated C/N ratio of 40:1 for bis-SorbPC, within the error in the XPS data, which is typically 15%. An Al2p peak at 74.8 eV is also observed (Figure 4b), consistent with the Al3+ surface treatment prior to film deposition. The peak position indicates that the adsorbed Al is present as the trivalent ion rather than an oxide.37 A significant reduction in Al3+ peak intensity is observed after deposition of the bis-SorbPC bilayer (Figure 4e), because of screening of the surface electrons by the 48 Å thick PSLB. AFM was used to examine the surface topography of (poly)bis-SorbPC PSLBs after drying and rehydration. Typical images are displayed in Figure 5, parts a and b, respectively. Figure 5c shows a line scan taken across a “scratch” made in the polymer bilayer through to the underlying substrate. The scratch was produced by repeatedly scanning the AFM tip at high force on the sample surface. The topographical depth determined by scanning perpendicular to the scratch is 48-52 Å, comparable to the thickness determined by ellipsometry. Returning to the images, significantly different morphologies were observed in the dried and rehydrated states. In the dry state (Figure 5a), the bilayer surface is composed of small, irregularly shaped domains, with an apparent diameter of 10-50 nm and separated by voids with an apparent depth of 5-10 Å. Some larger voids are also present, with an apparent diameter of 60 ( 15 nm and an apparent depth of 15-25 Å. The rms roughness was 5.2 ( 1.4 Å, whereas the roughness of the underlying silicon substrate was 2.1 ( 1.6 Å. Upon immersion in water, the surface morphology changed considerably (Figure 5b). The dried, “cracked” surface became much more uniform and the surface roughness decreased to 3.5 ( 0.8 Å. The larger voids present in the dried sample are still apparent although the apparent mean diameter decreased to about 40 nm with an apparent depth of 20 ( 5 Å. Analysis of image data for hydrated (poly)bis-SorbPC PSLBs shows that voids with a depth of 15-20 Å occupied 36 ( 8% of the surface area of the polymer film. The apparent depth of the voids in both hydrated and dried films indicates that these defects are present only in the upper leaflet of the bilayer structure. A probable cause for formation

of these voids is the desorption of low molecular weight oligomers from the upper leaflet during transfer of the bilayer across the air/water interface. In studies of mono-sorbyl lipid vesicles, UV polymerization was found to produce small polymer units composed of only 3-10 monomers.35 Domains of this size are probably small enough to be readily desorbed when a (poly)PSLB is dried. Similar observations were made in studies of (poly)bis-SorbPC PSLBs formed by vesicle fusion.22b Because the polymerizable groups in bis-SorbPC are located near the chain termini (Figure 1), it is possible that cross-linking occurs between the two leaflets of the bilayer. Thus, void formation may occur by desorption if a polymer unit in the upper leaflet is not covalently linked to the lower leaflet. This hypothesis is supported by comparative studies with bis-dienoyl phosphatidylcholine (bis-DenPC, Figure 1), in which the polymerizable groups are located near the glycerol backbone of the lipid. Water contact angle and ellipsometric measurements performed on UV polymerized, dried bis-DenPC PSLBs are summarized in Table 1. Polymerization followed by removal from water yielded a film with a thickness (25 Å) equivalent to only one-half of a lipid bilayer. The higher water contact angle, 65°, was indicative of a surface more hydrophobic than a bis-SorbPC bilayer (42 degree contact angle) and is consistent with a monolayerequivalent (i.e., tail group exposed) structure. The fact that only a monolayer-thick film of bis-DenPC was obtained may be a consequence of its structure, which precludes a covalent reaction between the inner and outer leaflets of the bilayer; this appears to result in desorption of the outer leaflet upon removal from water. In contrast, the structure of bis-SorbPC allows cross-linking between leaflets in a bilayer (assuming chain interdigitation), thereby stabilizing the structure (i.e., preventing desorption of the outer leaflet). The different results obtained with bis-DenPC and bis-SorbPC show that the location of the polymerizable unit in the lipid monomer is an important variable when creation of an air-stable bilayer is the objective. One possible explanation for the uniform appearance and regular spacing of the voids in both hydrated and dried bisSorbPC bilayers may be that polymerization occurs in domains, possibly due to phase segregation, either present in the unpolymerized film or occurring as a consequence of the polymerization process. The main phase transition temperature for bis-SorbPC is 28.8 °C.38 Under the conditions of deposition, the film should be in a coexistence regime,


Figure 6. C-H stretching region in FT-IR/ATR spectra of a bisSorbPC PSLB in contact with air (solid line) and with water (dashed line). To enable visual comparison of the band positions and widths, the background-subtracted spectra were normalized to the peak absorbance of the methylene asymmetric stretch.

with both liquid condensed (LC) and liquid expanded (LE) phases present. However, epifluorescence micrographs (not shown), taken by incorporating 2.0 mol % FITC-labeled phosphoethanolamine in an unpolymerized bis-SorbPC PSLB, exhibited uniform fluorescence. Thus, on a length scale of several microns, the film was uniform. However, this result does not negate the possibility of domain formation on submicron length scales that cannot be visualized optically but are detectable by AFM. Bis-SorbPC bilayers were also examined using FT-IR/ATR spectroscopy. It is well established that the peak frequency and the bandwidth of the methylene symmetric (νs) and asymmetric (νa) stretches are sensitive to changes in the gauche/trans conformer ratio, the number of conformational states, and motional rate of lipid hydrocarbon chains.39-44 For example, the gel to liquid-crystalline phase transition of dipalmitoylphosphatidylcholine (DPPC) is accompanied by an increase of 2.5 cm-1 in the νs peak frequency and an increase of 3 cm-1 in the bandwidth (at 0.75 of maximum intensity).43 Similar shifts are observed for the νa band and for the νs and νa bands of numerous other phospholipids.43,44 Changes in frequency and bandwidth were observed upon hydration of a previously dried (poly)bis-SorbPC PSLB (Figure 6): (1) The νs peak frequency shifted from 2856.1 to 2854.3 cm-1. (2) The νa peak frequency shifted from 2927 to 2924.5 cm-1. (3) The fwhm of the νs band narrowed from 22.8 to 21.3 cm-1. (4) The full width at half-maximum (fwhm) of the νa band narrowed from 44.1 to 37.3 cm-1. Thus, with reference to published data,39-44 these changes are indicative of greater acyl chain order (i.e., a lower gauche/ trans ratio) and a reduced number of conformational states in a hydrated versus a dried (poly)bis-SorbPC PSLB. Based on these data and the AFM images presented in Figure 6, we hypothesize that, when water solvating the PC headgroups at the outer surface of the lipid bilayer is removed, the lipids coalesce in an effort to minimize the headgroup surface area in contact with air. This process increases disorder in the underlying lipid alkyl chains and increases the void content and size. Rehydration of the headgroups

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Figure 7. Adsorption isotherms of FITC-labeled BSA on a POPC monolayer (solid line), a bis-SorbPC PSLB after drying and rehydration (dashed line), and a fluid POPC PSLB (dash-dot line). The lines through the data represent nonlinear least-squares fits to the Langmuir adsorption isotherm. The results are stated in Table 2. Table 2. Affinity Constants (Ka) and Saturation Fluorescence Intensities (Fmax) Obtained for the FITC-BSA Adsorption to a POPC Monolayer, a Bis-SorbPC Bilayer, and a POPC Bilayer surface

Ka

Fmaxa

POPC (monolayer) 9.8 ( 2.9 × 106 1.0 ( 0.067 bis-SorbPC (polymerized bilayer) 9.1 ( 2.1 × 105 0.51 ( 0.049 POPC (bilayer) 4.8 ( 0.32 × 105 0.16 ( 0.0046 a F max and Ka values were obtained from fitting the data to a Langmuir adsorption isotherm (see text). Fmax values were normalized to that obtained for the POPC monolayer, which was assumed to represent monolayer surface coverage of adsorbed BSA. Thus, Fmax values represent relative fractional surface coverages at apparent saturation.

causes expansion of the lipid layer, relaxing the strain in the lipid chains to create a more ordered and homogeneous structure. Protein Adsorption Studies. To assess if the nonspecific protein adsorption characteristics of a fluid lipid bilayer are altered by cross-linking polymerization, drying, and rehydration, comparative studies of FTIC-BSA adsorption to fluid POPC bilayers and rehydrated (poly)bis-SorbPC bilayers were conducted. Measurements were also performed on “tail group-out” POPC monolayers, which served as a model of a hydrophobic surface at which protein adsorption is highly favored. Adsorption isotherms were measured using TIRF spectroscopy.33,45 Representative data for all three surfaces are plotted in Figure 7. Apparent binding affinities (Ka) and relative fractional surface coverages (Fmax) were extracted by fitting the data sets to a Langmuir model (eq 1). The recovered parameters are listed in Table 2. Despite the apparent goodness of fit observed in Figure 7, it is unlikely that BSA adsorption to these surfaces obeys ideal Langmuir behavior. Specifically, the inherent assumptions that only one type of surface site is present, that lateral interactions between adsorbed molecules are absent, and that the adsorption process is reversible are probably not valid. The recovered Ka and Fmax values should therefore be treated as empirical parameters that enable the adsorption isotherms to be compared. Calibrating the TIRF responses (as discussed above) and assuming that a monolayer of FITC-BSA was adsorbed on

848


the POPC monolayer allowed the recovered Fmax values to be to normalized. Thus, these values represent relative fractional surface coverages of adsorbed BSA at apparent saturation. Before proceeding, we note that using TIRF to determine the surface coverage of an adsorbed protein film can be problematic, as discussed previously by a number of authors.33,34,46 There are several contributing factors. Of most concern here is self-quenching of FITC fluorescence due to energy transfer, which may occur when FITC-labeled protein molecules are closely packed in an adsorbed film. This will cause the ratio φs/φb (in eq 2) to decrease as the surface coverage increases. Returning to the Fmax values listed in Table 2, the net effect of self-quenching is that the values listed for the POPC and bis-SorbPC bilayers may be overestimated. Cognizant of these limitations, the data can still be compared semiquantitatively. Consistent with expectations, the extent of BSA adsorption to a POPC PSLB is considerably different than to a hydrophobic POPC monolayer. The respective Ka values, 4.8 ( 0.32 × 105 and 9.8 ( 2.9 × 106, differed by more than an order of magnitude, and the respective Fmax values were 0.16 and 1 monolayer. These differences show that BSA binds more strongly and at higher density on the hydrophobic surface, which is consistent with a large body of evidence showing that protein adsorption is highly favored at hydrophobic surfaces.3 The Ka and relative surface coverage measured for BSA adsorption to a (poly)bis-SorbPC PSLB after drying and rehydration were 9.1 ( 2.1 × 105 and 0.51 monolayer, respectively, which are intermediate between the values measured for the POPC bilayer and monolayer. Thus, the resistance of a fluid PC lipid bilayer to nonspecific protein adsorption is partially retained after polymerization, drying, and rehydration. The intermediate Ka suggests the possible existence of two types of adsorption sites on (poly)bis-SorbPC; however, fitting the data to a sum of two independent binding processes did not produce a statistically better fit. The greater degree of protein adsorption relative to the POPC bilayer can be attributed to the nonuniformity of the (poly)lipid films. As discussed above, after drying and rehydration, a (poly)bis-SorbPC bilayer contains voids with an apparent depth of approximately one lipid monolayer. These voids expose the hydrophobic core of the bilayer interior, leading to adsorption of soluble proteins. A quantitative correlation between exposed hydrophobic adsorption sites on the polymer surface and the amount of BSA adsorption can be made. As noted above, the voids on a hydrated (poly)bis-SorbPC PSLB occupy 36 ( 8% of the total surface area, as determined by AFM. The amount of BSA adsorbed to the voids should be approximately equal to the difference in relative fractional surface coverage on a (poly)bis-SorbPC bilayer and a POPC bilayer. This difference (0.51 minus 0.16 monolayer) is 0.35 monolayer. Thus, the increase in BSA adsorption on bis-SorbPC, relative to POPC, appears to be quantitatively correlated to the density of hydrophobic voids in the upper leaflet of the (poly)bisSorbPC bilayer. This correlation is consistent with results from our recent studies of bis-SorbPC PSLBs formed by vesicle fusion.22

Conboy et al.

Redox-initiated radical polymerization produced a bilayer with a very low defect density. The amount of nonspecific BSA adsorption to a bilayer formed in this manner was found to be equivalent to that of a fluid PSLB composed of 1-palmitoyl-2-oleoylphosphatidylcholine (POPC), even after the former had been dried and rehydrated. Finally, we note that this comparison also addresses the relationship between lateral lipid mobility and the degree of nonspecific protein adsorption to bilayers. In a recent paper,48 Glasma¨star et al. suggested that the rapid lateral diffusion of PC lipids in a fluid phase bilayer, relative to a gel phase bilayer, may be responsible for the capability of the former to resist nonspecific interactions. Data reported by Semple et al.47 provides support for this hypothesis; they observed that plasma protein binding to gel phase PC liposomes was considerably greater than binding to fluid phase liposomes. However, the data presented in ref 22c clearly demonstrate that fluidity is not required for a phosphorylcholine PSLB to resist nonspecific protein adsorption, because the lipids in a cross-linked bis-SorbPC bilayer are immobile. By extension, we conclude that the greater degree of BSA adsorption to the (poly)bis-SorbPC PSLBs prepared and characterized in this study, relative to the amount adsorbed to POPC, cannot be attributed to the lack of lateral lipid mobility in the (poly)lipid. Summary and Conclusions Substrate supported lipid bilayers have been prepared from the lipid monomer bis-SorbPC and polymerized by UV irradiation. The overall bilayer structure is largely preserved upon removal of the film from water, although significant loss of material occurs from the upper leaflet of the bilayer, probably because of desorption of the relatively lower molecular weight products. Comparative studies with bisDenPC, in which the polymerizable groups are located near the lipid backbone, suggest that inter-leaflet linkages in a (poly)bis-SorbPC bilayer contribute to its structural stability. The morphology and surface structure of (poly)bis-SorbPC PSLBs, as observed by AFM, indicate a different arrangement of the lipids in the hydrated and dehydrated states, presumably because of the loss of water from the near surface region. These changes have been correlated with changes in the conformation of the hydrocarbon chains by FT-IR/ATR. The resistance of a rehydrated bis-SorbPC PSLB to nonspecific adsorption of BSA is intermediate between model hydrophobic and fluid PC bilayer surfaces. The greater protein adsorption relative to a POPC bilayer is correlated with the desorption of material from the upper leaflet of the bilayer, which produces voids at which hydrophobically driven protein adsorption occurs. In comparison to (poly)bis-SorbPC PSLBs formed by vesicle fusion and redox-initiated radical polymerization,22 LBS deposition followed by UV photopolymerization produces PSLBs with a higher defect density and decreased resistance to nonspecific protein adsorption. The difference in deposition method does not appear to be the critical factor, because vesicle fusion followed by UV photopolymerization also produces (poly)PSLBs22b that are significantly less protein resistant than those polymerized by redox-initiation.22c


Acknowledgment. This material is based upon work partially supported by the National Science Foundation under Grant Numbers CHE-0108805 to S.S.S. and DMR-9619887 to D.F.O. J.C.C. was partially supported by a NRSA from NIH (GM19914). The authors would also like to thank Prof. Jeanne Pemberton and Mr. Domenic Tiani for the use of the FT-IR spectrometer and ellipsometer. Additional thanks go to the surface science facility at the University of Arizona for XPS analysis. References and Notes (1) (a) Barker, S. A. In Biosensors: Fundamentals and Applications; Turner, A. P. F., Karube, I, Wilson, G. S., Eds.; Oxford: New York, 1987; pp 85-99. (b) Taylor, R. F. Protein Immobilization: Fundamentals and Applications; Marcel Dekker: New York, 1991. (c) Rao, S. V.; Anderson, K. W.; Bachas, L. G. Mikrochim. Acta 1998, 128, 127-143. (2) Some examples of oriented immobilization are (a) P. L. Edmiston, P. L.; Saavedra, S. S. Biophys. J. 1998, 74, 999-1006. (b) Burgess, J. D.; Rhoten, M. C.; Hawkridge, F. M. Langmuir 1998, 14, 24672475. (3) (a) Brash, J. L.; Horbett, T. A. In Proteins at Interfaces II; Horbett, T. A., Brash, J. L., Eds.; ACS Symposium Series 602; American Chemical Society: Washington, DC, 1995; pp 1-23 and references therein. (b) Hlady, V.; Buijs, J. Curr. Opin. Biotech. 1996, 7, 7277 and references therein. (4) (a) Wisniewski, N.; Reichert, M. Colloids Surf. B-Biointerfaces 2000, 18, 197-219. (b) Wisniewski, N.; Moussy, F.; Reichert, M. Fresenius J. Anal. Chem. 2000, 366, 611-621. (5) For reviews, see: (a) Plant, A. L. Langmuir, 1999, 15, 5128-5135. (b) Sackmann, E. Science 1996, 271, 43-48. (6) Examples of lipid membrane-based sensors include the following: (a) Terrettaz, S.; Stora, T.; Duschl, C.; Vogel, H. Langmuir 1993, 9, 1361-1369. (b) Puu, G. Anal. Chem. 2001, 73, 72-79. (c) Stelzle, M.; Weissmu¨ller, G.; Sackmann, E. J. Phys. Chem. 1993, 97, 297481. (d) Song, X. D.; Swanson, B. I. Anal. Chem. 1999, 71, 20972107. (e) Schmidt, C.; Mayer, M.; Vogel, H. Angew. Chem., Int. Ed. 2000, 39, 3137-3140. (f) Cornell, B. A.; Braach-Maksvytis, V.; King, L. G.; Osman, P. D. J.; Raguse, B.; Wieczorek, L.; Pace, R. J. Nature 1997, 387, 580-583. (g) Tien, H. T.; Ottova, A. L. Colloids Surf. A 1999, 149, 217-233. (7) Gennis, R. B. Biomembranes; Springer-Verlag: New York, 1989. (8) (a) Zwall, R. F. A.; Hemker, H. C. Haemostasis 1982, 11, 12. (b) Chapman, D. Langmuir 1993, 9, 39. (9) McConnell, H. M.; Watts, T. H.; Weis, R. M.; Brian, A. A. Biochim. Biophys. Acta 1986, 864, 95-106. (10) Thompson, N. L.; Palmer, A. G., III. Comments Mol. Cell. Biophys. 1988, 5, 39-56. (11) (a) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985, 47, 105113. (b) Brian, A. A.; McConnell, H. M. Proc. Natl. Acad. Sci. 1984, 81, 6159-6163. (12) (a) Salafsky, J.; Groves, J. T.; Boxer, S. G. Biochemistry 1996, 35, 5, 14773-14781. (b) Burgess, J. D.; Rhoten, M. C.; Hawkridge, F. M. Langmuir 1998, 14, 2467-2475. (c) Salamon, Z.; Tollin, G. Biophys. J. 1996, 71, 858-867. (d) Heyse, S.; Ernst, O. P.; Dienes, Z.; Hofmann, K. P.; Vogel, H. Biochemistry 1998, 37, 507-522. (13) (a) Edmiston, P. L.; Saavedra, S. S. J. Am. Chem. Soc. 1998, 120, 1665-1671. (b) Edmiston, P. L.; Saavedra, S. S. Biophys. J. 1998, 74, 999-1006. (c) Fischer, B.; Heyn, S. P.; Egger, M.; Gaub, H. E. Langmuir 1993, 9, 136. (14) Cremer, P. S.; Boxer, S. G. J. Phys. Chem. B 1999, 103, 25542559. (15) (a) Dietrich, C.; Bagatolli, L. A.; Volovyk, Z. N.; Thompson, N. L.; Levi, M.; Jacobson, K.; Gratton, E. Biophys. J. 2001, 80, 14171428. (b) Hillebrandt, H.; Wiegand, G.; Tanaka, M.; Sackmann, E. Langmuir 1999, 15, 8451-8459. (c) Kalb, E.; Frey, S.; Tamm, L. K. Biochim. Biophys. Acta 1992, 1103, 307-316. (d) Majewski, J.; Wong, J. Y.; Park, C. K.; Seitz, M.; Israelachvili, J. N.; Smith, G. S. Biophys. J. 1998, 75, 2363-2367. (16) (a) Hayward, J.; Chapman, D. Biomaterials 1984, 5, 135. (b) Albrecht, O.; Johnston. D. S.; Villaverde, C.; Chapman, D. Biochim. Biophys. Acta 1982, 687, 165-169. (17) (a) Foltynowicz, Z.; Yamaguchi, K.; Czajka, B.; Regen, L. R. Macromolecules 1985, 18, 1394-1401. (b) Regen, S. L.; Kirszensztejn, P.; Singh, A. Macromolecules 1983, 16, 335-338.

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BM0256193

Planar Supported Bilayer Polymers Formed from Bis-Diene Lipids by

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