Covalent Coupling and Characterization of Supported Lipid Layers

Polymer-supported lipid monolayers and bilayers present an opportunity to develop rugged cell mimetic ... Macromolecular Symposia 2016 365 (1), 49-58 ...
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Covalent Coupling and Characterization of Supported Lipid Layers Sally L. McArthur,*,†,‡ Michael W. Halter,‡ Viola Vogel,†,‡ and David G. Castner†,‡ National ESCA and Surface Analysis Center for Biomedical Problems (NESAC/Bio), Departments of Bioengineering and Chemical Engineering, University of Washington, Box 351750, Seattle, Washington 98195-1750, and University of Washington Engineered Biomaterials (UWEB), Department of Bioengineering, University of Washington, Box 351721, Seattle, Washington 98195-1721 Received November 29, 2002. In Final Form: June 2, 2003 To investigate protein and cell surface interactions, robust and increasingly complex model surfaces need to be developed to mimic specific aspects of the cell membrane structure. Polymer-supported lipid monolayers and bilayers present an opportunity to develop rugged cell mimetic surfaces that may be stable in a wide range of in vivo and in vitro applications and characterization techniques. We have investigated the stability of dimyristoylphosphatidylethanolamine (DMPE) monolayers grafted to poly(hydroxyethyl methacrylate) (pHEMA) via 1,1′-carbonyldiimidazole (CDI) chemistry using X-ray photoelectron spectroscopy (XPS), time-of-flight secondary ion mass spectrometry (ToF-SIMS), and fluorescence microscopy. The results illustrate that it is possible to covalently couple the amine-containing lipid headgroup to the carboxyl and hydroxyl groups of the pHEMA and retain a proportion of the lipids at the surface after the samples are sonicated in ethanol. Most importantly, the cross-linking efficiency and retention of the resulting lipid layer is higher if the lipid is transferred to CDI-activated HEMA by Langmuir-Blodgett deposition rather than by adsorption of the lipid directly from solution. Both factors are critical if these monolayers are to form a stable and reproducible model system.

Introduction The membranes that define the boundaries of biological cells consist of lipids, proteins, and carbohydrates, all of which have specific functions in the stabilization of cell membranes, cell communication, and signaling.1 To investigate events occurring at biomaterial interfaces, increasingly complex model systems are being sought. The optimal system would be a cell mimetic surface formed with fluid lipid bilayers, transmembrane and membrane proteins, and a variety of other biomolecules. However, simpler models made up of ordered lipid layers with varying extents of lateral mobility and differing headgroup functionality may also give valuable insight into protein/ lipid and cell/surface interactions. These simple systems need to be reproducible and robust as many of the techniques developed to study biomaterial surface interactions require specific conditions such as ultrahigh vacuum (UHV). In addition, there is a desire to have models that will be suitable for in vivo studies and stable enough to act as sensing units in biosensors and microfluidic devices. The most common lipid monolayer and bilayer systems discussed in the literature are created using Langmuir transfer techniques or vesicle fusion directly onto glass substrates. Under these conditions, a thin film of water lubricates the interface between the glass and the bilayer and permits free lateral diffusion, a factor that is vital to the biological function of cell membranes.2,3 While * Corresponding author. Current address: Department of Engineering Materials, University of Sheffield, Mappin St., Sheffield S1 3JD, UK. E-mail: [email protected]. † NESAC/Bio. ‡ UWEB. (1) Plant, A. L. Langmuir 1999, 15, 5128. (2) Dufrene, Y. F.; Lee, G. U. Biochim. Biophys. ActasBiomembr. 2000, 1509, 14.

there has been some recent success culturing cells on these types of bilayers,4 they are generally regarded to have poor long-term stability1 and cannot be transferred through the air-water interface without disrupting the layer structure.3 Recent approaches for creating more rugged model systems have involved either hybrid bilayers where the inner leaflet is formed from alkane thiol on gold,1,5,6 or deposition of the lipids on a polymer support.3,7,8 The polymer creates a soft, hydrated cushion that acts as both a self-lubricating surface and spacer. As well as permitting a fraction of the inner leaflet lipids to be coupled to the surface to improve the surface stability, the polymer cushion provides a capacity for self-healing that is critical in the prevention of nonspecific interactions.3 To create robust model systems suitable for in vitro and in vivo studies of protein/lipid interactions and compatible with a wide range of analysis techniques, we have followed the polymer support route in our surface design. While both poly(ethylene imine) (PEI) and dextran are more commonly used as supports,3,8 we chose poly(hydroxyethyl methacrylate) (pHEMA) to provide the highly hydrated polymer network. The critical advantage of HEMA lies in the fact that while it can be used as a bulk material, the presence of hydroxyl and carboxyl groups within HEMA makes it possible to covalently couple it to a variety of substrates and to have a proportion of the lipids attached (3) Sackmann, E.; Tanaka, M. Trends Biotechnol. 2000, 18, 58. (4) Groves, J. T.; Mahal, L. K.; Bertozzi, C. R. Langmuir 2001, 17, 5129. (5) Tegoulia, V.; Rao, W.; Kalambar, A.; Rabolt, J.; Cooper, S. Langmuir 2001, 17, 4396. (6) Lahiri, J.; Kalal, P.; Frutos, A.; Jonas, S.; Schaeffler, R. Langmuir 2000, 16, 7805. (7) Majewski, J.; Wong, J. Y.; Park, C. K.; Seitz, M.; Israelachvili, J. N.; Smith, G. S. Biophys. J. 1998, 75, 2363. (8) Wong, J. Y.; Majewski, J.; Seitz, M.; Park, C. K.; Israelachvili, J. N.; Smith, G. S. Biophys. J. 1999, 77, 1445.

10.1021/la026928h CCC: $25.00 © 2003 American Chemical Society Published on Web 08/23/2003

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at the interface. A further benefit of using HEMA is that its manufacturing technology is already well developed for a number of medical device industries including contact lenses, and as such its interactions with biological systems have been well characterized.9-14 In addition, while HEMA is not a nonfouling material per se, specific HEMA-based polymers currently used in soft contact lenses have been shown to adsorb low levels of protein in vivo and in vitro.15,16 This may prevent nonspecific interactions from affecting the polymer cushion and aid in the maintenance of the ordered lipid structure under implant conditions. Here we investigate methods for coupling HEMA onto a glass substrate and subsequently coupling 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine (DMPE) onto the HEMA under a variety of conditions. Fluorescence microscopy was used to image the resulting lipid structures and to compare the organization of the lipid layers. The efficacy of the reaction chemistry and the stability of the resulting surfaces were investigated using X-ray photoelectron spectroscopy (XPS) and time-of-flight secondary ion mass spectrometry (ToF-SIMS). Due to the complex nature of the ToF-SIMS spectra, principal component analysis (PCA) was used to compare the positive and negative ion spectra and to determine the fragments responsible for variation between spectra, an approach previously used to characterize polymer surfaces17,18 and adsorbed protein films.19 The results of this study illustrated that it was possible to chemically couple the lipid to a HEMA substrate and that the process enabled the retention of the lipid at the interface under harsh conditions. Materials and Methods Surface Modification. Hydroxyethyl methacrylate (HEMA), methacrylic acid (MAA), benzoin methyl ether (BME), 1,1′carbonyldiimidazole (CDI) (Sigma, St. Louis, MO), ethylene glycol dimethacrylate (EGDMA) (Fluka Chemicals, Basel, Switzerland), 3-methacryloxypropyldimethylchlorosilane (Gelest Inc., Tullytown, PA), and 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine (DMPE) (Avanti, Alabaster, AL) were purchased and used as received without further purification. All glass cover slips (25 mm, Corning No. 1) were first cleaned in Piranha solution and then stored under water until used. Caution! Piranha solution is a very strong oxidizer and is extremely dangerous to handle in the laboratory; gloves, goggles, and face shields are needed for protection. Silane surface modification was achieved using a method developed previously in our laboratory by Dannenberger et al.20 Clean glass cover slips were treated with a 10-15% solution of 3-methacryloxypropyldimethylchlorosilane in methylene chloride (MeCl) (Fisher Research, Pittsburgh, PA) for 1 h. Samples were bath sonicated three times in MeCl to remove any residual silane (9) Schultz, C. L.; Kunert, K. S.; White, R. J. Ind. Microbiol. Biotechnol. 2000, 24, 113. (10) Willcox, M. D. P.; Harmis, N.; Cowell, B. A.; Williams, T.; Holden, B. A. Biomaterials 2001, 22, 3235. (11) Franklin, V. J.; Bright, A. M.; Tighe, B. TRIP 1993, 1, 9-16. (12) Dalton, P. D.; Flynn, L.; Shoichet, M. S. Biomaterials 2002, 23, 3843. (13) Karakecili, A. G.; Gumusderelioglu, M. J. Biomater. Sci. Polym. Ed. 2002, 13, 185. (14) Tunney, M. M.; Jones, D. S.; Gorman, S. P. Int. J. Pharm. 1997, 151, 121. (15) Jones, L.; Evans, K.; Sariri, R.; Franklin, V.; Tighe, B. CLAO 1997, 23, 122. (16) McArthur, S. L.; McLean, K. M.; St. John, H. A. W.; Griesser, H. J. Biomaterials 2001, 22, 3295. (17) McArthur, S. L.; Wagner, M. S.; McLean, K. M.; Hartley, P. G.; Griesser, H. J.; Castner, D. G. Surf. Interface Anal. 2002, 33, 924. (18) Delcorte, A.; Bertrand, P.; Arys, X.; Jonas, A.; Wischerhoff, E.; Mayer, B.; Laschewsky, A. Surf. Sci. 1996, 366, 149. (19) Wagner, M. S.; Castner, D. G. Langmuir 2001, 17, 4649. (20) Dannenberger, O.; Boeckl, M.; Bassuk, J. A.; Valint, P. L.; Sasaki, T.; V. Vogel. AVS Trans. 1998, 45.

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Figure 1. Schematic of pHEMA and DMPE grafting chemistry. (a) Reaction of CDI with HEMA. (b) Reaction of DMPE with CDI activated HEMA. and dried overnight under vacuum. For HEMA surface modification, a 12 µL droplet of the base HEMA monomer mix was placed on a clean glass microscope slide, and then the silanetreated cover slip was gently placed on top. The base monomer mix was made by the addition of 78.4 mL of HEMA, 0.104 mL of EGDMA, 2.6 mL of MAA, 172 mg of BME, and 13.8 mL of N′,N-dimethylformamide (DMF) (Aldrich Chemicals Milwaukee, WI) and vortexing. The polymer coatings were cured under a UV lamp (254 nm) for 2 h and then immediately immersed in water for >72 h. This enabled the HEMA-coated cover slip to be removed from the microscope slide using a razor blade. Surfaces were bath sonicated for 5 min in water, then in 50/50 water/EtOH, and then in water, and they were finally dried under a nitrogen stream before further analysis. The pHEMA layers on glass were reacted with CDI, which activates hydroxyls and carboxylates present in the polymer film and enables subsequent reaction with amine-containing lipids. Samples were treated with 0.25 M CDI in 1,4-dioxane (Fisher Research, Pittsburgh, PA) for 2 h at 37 °C. Samples were rinsed three times in dioxane, then sonicated in dioxane for 5 min, and dried under a nitrogen stream. For solution treatment with DMPE lipids, samples were immersed in solutions of 1 mg/mL DMPE in 50/50 EtOH/water for 37 h at 37 °C while stirring. Samples were rinsed three times in water, then sonicated in water for 5 min, and dried under a nitrogen stream. For Langmuir-Blodgett (LB) transfers, pure DMPE monolayers were spread from chloroform (Aldrich Chemicals, Milwaukee, WI) solutions (approximately 1 mg/mL) at the air/water interface in a Teflon trough (8 × 18 × 0.7 cm) with a dipping well at 22 ( 1 °C. The surface pressure was monitored by the Wilhelmy plate technique.21 During vertical withdrawal of the substrate, the surface pressure (20 mN/m) was maintained using a computer controlled feedback loop. Transfer ratios were approximately 1. Samples were subsequently either dried in a nitrogen stream or sonicated in ethanol for 5 min and then dried. A schematic of the reaction chemistry is presented in Figure 1. Surface Analysis. XPS analysis was performed using a SSI X-Probe equipped with a monochromated Al KR source at a power (21) Adamson, A. W.; Gast, A. P. Physical chemistry of surfaces; Wiley and Sons: New York, 1997.

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Table 1. XPS Atomic Compositions of the Surfaces after Each Stage of the Grafting Process C 1s

O 1s

N 1s

P 2p

Si 2p

O:C

N:C

HEMA (theory) HEMA HEMA + DMPE HEMA + LB DMPE HEMA + LB DMPE (EtOH sonicated)

66.0 69.3 ( 0.3 69.5 ( 0.8 79.9 ( 0.9 69.1 ( 0.6

33.0 29.6 ( 0.5 30.3 ( 0.9 18.0 ( 0.4 30.9 ( 0.6

0.0 0.0 ( 0.0 0.0 ( 0.0 0.6 ( 0.5 0.0 ( 0.0

0.0 0.0 ( 0.0 0.3 ( 0.1 1.5 ( 0.3 0.0 ( 0.0

0.0 1.1 ( 0.3 0.0 ( 0.0 0.0 ( 0.0 0.0 ( 0.0

0.5 0.43 ( 0.01 0.44 ( 0.02 0.23 ( 0.01 0.45 ( 0.01

0.0 0.00 ( 0.00 0.0 ( 0.0 0.01 ( 0.01 0.00 ( 0.00

HEMA + CDI HEMA + CDI + DMPE HEMA + CDI + LB DMPE HEMA + CDI + LB DMPE (EtOH sonicated)

64.6 ( 1.1 66.5 ( 0.5 78.2 ( 0.4 68.6 ( 2.0

Covalent Coupling 24.6 ( 1.5 10.8 ( 0.5 27.5 ( 1.0 6.1 ( 0.6 17.6 ( 0.2 3.0 ( 0.2 23.5 ( 2.4 7.7 ( 0.3

0.0 ( 0.0 0.0 ( 0.0 1.2 ( 0.2 0.1 ( 0.3

0.0 ( 0.0 0.0 ( 0.0 0.0 ( 0.0 0.0 ( 0.0

0.38 ( 0.03 0.41 ( 0.02 0.22 ( 0.00 0.34 ( 0.04

0.17 ( 0.01 0.09 ( 0.01 0.04 ( 0.00 0.11 ( 0.00

of 200 W (1000 µm spot). Elements present on the surface were identified from a survey scan. For further analysis and quantification, spectra from the individual peaks were collected at 150 eV pass energy (res 4). High-resolution spectra were collected at 50 eV (res 2). All data were collected at 55° (from the surface normal). A value of 285 eV for the binding energy of the main C 1s component (CHx) was used to correct for charging of specimens under irradiation.22 Multiple samples were analyzed from each batch, and the results were averaged. F-tests and Student’s t-tests were used to evaluate the statistical variations between each of the sets of data, with p < 0.01 considered significant. High-resolution C 1s spectra were fitted using a Shirley background subtraction and a series of Gaussian peaks. ToF-SIMS analysis of all surfaces was performed using a PHI Model 7200 Reflectron time-of-flight secondary ion mass spectrometer (Physical Electronics, Eden Prairie, MN) equipped with an 8 keV Cs+ primary ion source. Positive and negative ion spectra were acquired by rastering the ion beam over a 200 µm × 200 µm sample area with data collected in the 0-200 m/z range. The primary ion dose was less than 1012 ions/cm2 to maintain static ToF-SIMS conditions,23 and charge neutralization was achieved with a pulsed electron flood gun. The mass resolution (m/∆m) at C2H3+ (m/z ) 27) was typically above 5000. Spectra were calibrated to the CH3+, C2H3+, C3H5+, and C7H7+ or a combination of CH-, CHO-, CN-, and CNO- peaks prior to further analysis. A minimum of three samples and three points per sample were analyzed for each sample type. Principal component analysis (PCA) of the spectra was performed using the PLS Toolbox v 2.0 (Eigenvector Research, Manson, WA) for MATLAB (The Mathworks Inc, Natick, MA). Prior to multivariate analysis, a peak set was created assigning a centroid peak value and area of integration window for every peak detected in each spectrum. The areas of these peaks were then calculated and normalized to the total intensity of all peaks in each individual spectrum to correct for differences in total secondary ion yield between spectra. Data were mean centered (set at the origin) to ensure that the variance in the data set was due to differences in sample variances rather than differences in sample means.24 Fluorescence Imaging. The Langmuir monolayers were imaged at the air/water interface by adding 1-2 mol % of the probe1-myristoyl-2-[12-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]dodecanoyl]-sn-glycero-3-phosphoethanolamine to the DMPE. Images were then acquired using a fluorescence microscope (Nikon) with an attached CCD (Photometrix, CoolSnap). To image the DMPE monolayers after they were Langmuir-Blodgett transferred to glass, HEMA, and CDI activated HEMA, 1-2 mol % of the probe 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-2-1,3-benzoxadiazol-4-yl) was added to the DMPE and the surfaces were imaged using a fluorescence microscope (Nikon, TE 200) with an attached CCD (Princeton Instruments, TEK512). (22) Beamson, G.; Briggs, D. High resolution XPS of organic polymers. The Scienta ESCA00 database; John Wiley and Sons: Chichester, 1992. (23) Marletta, G.; Catalano, S. M.; Pignataro, S. Surf. Interface Anal. 1990, 16, 407. (24) Vandeginste, B. G. M.; Massart, D. L.; Buydens, L. M. C.; deJong, S.; Lewi, P. J.; Smeyers-Verbeke, J. Handbook of chemometrics and qualimetrics: Part b; Elsevier Science Publishers B. V.: Amsterdam, 1998.

Figure 2. XPS high-resolution C 1s profiles of (a) HEMA, (b) HEMA + CDI, (c) HEMA + CDI + adsorbed DMPE, (d) HEMA + CDI + LB DMPE, and (e) HEMA + CDI + LB DMPE (EtOH sonicated).

Results and Discussion XPS analysis indicated the successful attachment of HEMA onto the glass cover slip, with the grafted layer atomic composition comparable to the theoretical composition of HEMA (Table 1). The subsequent reaction of the HEMA with CDI produced a significant increase in the nitrogen content of the surface and corresponding decreases in the carbon and oxygen contents. Analysis of the C 1s spectra indicated peak shifts associated with the introduction of both the C-N and N-COO species associated with imidazole carbamate active intermediates after CDI reaction with hydroxyl groups or acylimidazole active intermediates after reaction with carboxylates (Figure 2, curves a and b, and Table 2). To further investigate the information provided by XPS, ToF-SIMS and principal component analysis (PCA) were used to analyze each stage of the grafting process. Figure 3 shows the results of PCA analysis comparing the ToFSIMS negative ion spectra from HEMA and HEMA + CDI samples. The scores plot for principal component 1 (PC 1) captured 99% of the variation in this data set and clearly separates the two sample types (Figure 3a). Analysis of the corresponding loading plot for PC 1 (Figure 3b) indicated that the key differences in the spectra were due to the presence of nitrogen containing CDI related fragments at the surface of the CDI + HEMA samples,

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Figure 3. (a) Scores and (b) loading plots on PC 1 generated from PCA analysis of negative ion ToF-SIMS spectra from (2) HEMA and (9) HEMA + CDI surfaces. Table 2. XPS High-Resolution C 1s Curve Fit Results after Each Stage in the Grafting Process 284.6 eV CHx

+1.7 eV C-O, C-N

+3.8 eV O-CdO

HEMA (theory) HEMA HEMA + LB DMPE HEMA + LB DMPE (EtOH sonicated)

50.0 55.3 ( 2.1 78.8 ( 0.7 52.9 ( 0.7

33.0 30.8 ( 1.3 13.6 ( 0.4 32.3 ( 0.3

16.7 13.9 ( 0.7 7.6 ( 0.2 14.8 ( 0.4

0.0 0.0 0.0 0.0

HEMA + CDI HEMA + CDI + DMPE HEMA + CDI + LB DMPE HEMA + CDI + LB DMPE (EtOH sonicated)

52.9 ( 1.8 53.8 ( 2.6 75.5 ( 1.0 59.0 ( 2.5

Covalent Coupling 30.1 ( 2.0 29.4 ( 3.2 16.3 ( 2.8 26.3 ( 1.7

9.9 ( 1.6 10.5 ( 0.4 8.8 ( 0.7 9.5 ( 2.5

7.1 ( 1.5 6.3 ( 1.3 0.9 ( 0.2 5.2 ( 0.7

confirming that the reaction chemistry proceeded successfully and did not introduce contamination. To characterize the effect of chemical cross-linking on lipid monolayer transfer and lipid stability, the lipid was deposited onto the surface by two methods. The first method involved adsorption of the lipid from a stirred solution onto HEMA-coated glass with and without CDI. These samples were then analyzed after sonication in water. In the second method, liquid condensed (LC) phase DMPE monolayers were transferred using the LB technique directly onto unmodified HEMA substrates and onto HEMA substrates previously activated with CDI. The resulting samples were analyzed before and after 5 min sonication in ethanol.

+5 eV N-COO

Lipid Adsorption from Solution. Analysis of the XPS-derived atomic composition (Table 1) indicated that if HEMA substrates were exposed directly to a lipid solution, very little if any lipid adsorbed to HEMA, resulting in no significant changes in any of the elements. Once the HEMA was activated with CDI, increases in carbon and decreases in nitrogen and oxygen indicative of lipid adsorption were evident. Analysis of the C 1s spectra from the HEMA + CDI + DMPE samples (Figure 2c and Table 2) showed increases in the hydrocarbon content of the surface, indicative of the adsorption of DMPE at the interface. The presence of lipid at the surface was confirmed by analysis of the related ToF-SIMS spectra using PCA. PC 1 captured 87% of the variation between

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Figure 4. Fluorescent images of DMPE monolayers at the air/water interface (a-c) and on solid supports (d-f) (1-2 mol % dye, T ) 22 °C). The DMPE monolayer at the air/water interface was imaged sequentially during compression at (a) 9, (b) 12, and (c) 20 mN/m. Condensed, ordered domains reject the dye and appear dark. The dye is more soluble in the liquid expanded phase, which appears bright. Monolayers after Langmuir-Blodgett transfer at a constant surface pressure of 20 mN/m are shown in (d-f), where (d) is glass, (e) is HEMA, and (f) is activated HEMA. The bars represent 50 µm.

the HEMA + CDI and HEMA + CDI + DMPE samples and clearly separated the two sample sets. Analysis of the corresponding loading plot illustrated that the greatest differences between the spectra were due to the presence of lipid related peaks including fragments of the lipid’s hydrocarbon tail in spectra from HEMA + CDI + DMPE samples (see Supporting Information). It should be noted that while the spectra from the adsorbed lipid samples did on average score negatively, there was a large spread in the scores for the individual spectra, indicating a degree of spectral variation within the sample set. Variations within and between the samples could be due to a number of factors including surface roughness and/or random or uneven distribution of the lipid over the HEMA surface. The surface and chemical sensitivity of ToF-SIMS was emphasized in the comparison of spectra from the HEMA and HEMA + DMPE samples. While the XPS analysis was not able to find any significant differences between the samples before and after lipid adsorption, PCA analysis separated the samples in PC 1 capturing 86% of the variation in the data set. The loading plot for PC 1 confirmed that the separation of the samples was due to the presence of adsorbed lipid at the surface of the HEMA + DMPE samples, with fragments of the lipid tail and PO3 detected in the spectra (see Supporting Information). Langmuir-Blodgett Transfer. Langmuir monolayers of DMPE form ordered films at the air/water interface. Monolayer films at the air/water interface that are transferred to solid supports by the LB technique can retain their ordered structure. To confirm that the structure of the DMPE monolayer present at the air/water interface was maintained upon transfer to the HEMA supports studied here by XPS and SIMS, fluorescence images of DMPE monolayers at the air/water interface were compared with images of the monolayers after LB transfer. Fluorescence images of DMPE monolayers at various surface pressures at the air/water interface were collected (Figure 4a-c). During compression, the dye is rejected from the ordered, condensed phase and retained in the liquid expanded phase. When the monolayer is completely condensed, the dye exists preferentially at the domain boundaries. Subsequently, fluorescent images of the fully condensed monolayers (at 20 mN/m) after LB

transfer onto glass, HEMA, or CDI activated HEMA substrates were also collected (Figure 4d-f). LB transferred monolayers of DMPE on glass are expected to have ordered lipid chains and maintain the structure of the initial monolayer at the air/water interface. Notably, the images of the lipid monolayers on the HEMA and activated HEMA substrates are similar to the glass supported lipid monolayer. This indicates that the monolayers on the polymer supports have retained the ordered structure of the original monolayers at the air/water interface. In the images of the solid support monolayers (Figure 4d-f), the edges of the dark condensed domains appear bright because the dye has segregated out of the ordered phase. The solid supported monolayers have significantly more fluorescence at the domain edges compared to monolayers at the air/water interface at the same surface pressure (Figure 4c). Most likely, this increased fluorescence results from melting of the domains ahead of the three-phase line during deposition25 or from reduced fluorescence of the probe in the aqueous environment at the air/water interface.26,27 XPS analysis of LB transferred monolayers directly onto the HEMA surface showed a significant increase in the carbon and a decrease in oxygen content of the surfaces (Table 1). Analysis of the XPS C 1s profiles from each of the samples indicated significant increase in the hydrocarbon contents of the surfaces and corresponding decreases in peaks associated with the C-O and OdC-O components of the HEMA substrate (Table 2). After sonication in ethanol, XPS data indicated that all lipids were removed from the surface, with the atomic composition of the surface inseparable from that of the untreated HEMA substrate (Table 1). Interestingly, analysis of the curve-fitted data from the C 1s spectra indicated that the sonication actually appeared to have reduced the hydrocarbon content of the surface below that of the untreated samples and closer to the theoretical values of HEMA. The result suggests that the sonication in effect completely (25) Riegler, H.; Spratte, K. Thin Solid Films 1992, 210/211, 9. (26) Lin, S.; Struve, W. Photochem. Photobiol. 1991, 54, 361. (27) Chattopadhyay, A. Chem. Phys. Lipids 1990, 53. (28) McLean, K. M.; McArthur, S. L.; Chatelier, R. C.; Kingshott, P.; Griesser, H. J. Colloids Surf. B: Biointerfaces 2000, 17, 23.

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Figure 5. (a) Scores and (b) loading plots on PC 1 generated from PCA analysis of negative ion ToF-SIMS spectra from (0) HEMA, (2) HEMA + 20mN/m DMPE, and (b) HEMA + 20mN/m DMPE (EtOH sonicated) surfaces.

cleaned the surface of the sample, removing some of the adventitious carbon contamination present on the control. PCA analysis of the corresponding ToF-SIMS data confirmed that while LB transfer did transfer lipid to the nonactivated HEMA surface, the lipid layer was not stable under sonication. Figure 5 shows the scores and loading plots produced by comparing the negative ion spectra from the HEMA and HEMA + LB DMPE samples before and after sonication with principal component 1 (PC 1) capturing 99% of the variation between the spectra. The scores plot for PC 1 (Figure 5a) illustrates that this variation separated the HEMA + LB DMPE spectra from those of the unmodified HEMA and the sonicated HEMA + LB DMPE. The loading plot for PC 1 (Figure 5b) indicates that fragments associated with the lipid, including the lipid molecular ion (negative loadings) and HEMA associated fragments (positive loadings), were the source of the greatest variation between spectra. Interestingly, subsequent PCs were not able to separate the HEMA spectra from those of the sonicated HEMA + LB DMPE samples. LB transfer of DMPE onto the CDI activated HEMA led to a significant increase in the carbon content and a decrease in oxygen content of the surfaces (Table 1). Analysis of the C 1s profiles from each of the samples indicated significant increase in the hydrocarbon contents of the surfaces, indicative of lipid transfer (Figure 2d and Table 2). The nitrogen content of the HEMA + CDI surface decreased significantly with the introduction of the lipid

layer, but the signal was not eliminated and, correspondingly, there was evidence of N-COO related peak shifts in the XPS C 1s profile (Figure 2d and Table 2). While the presence of the lipid on the HEMA substrate was shown to introduce a small amount of nitrogen presumably from the ethanolamine headgroup (see HEMA + LB DMPE data in Table 1), the high energy of the associated peak shift reflects sampling of the underlying CDI activated HEMA substrate. Intact lipid monolayers of DMPE produced at transfer ratios close to unity are expected to be 2-3 nm thick,8 so the HEMA + CDI substrate should be well within the XPS sampling depth (5-6 nm). Critically, the presence of the CDI did not appear to significantly disrupt the formation of the lipid monolayer. Sonication of the samples in ethanol significantly reduced the carbon contents of the HEMA + CDI + LB DMPE surfaces, but did not return them to the same composition as the HEMA + CDI surface. Analysis of the C 1s profiles indicated that there was still lipid present at the interface, with the surfaces having higher proportions of hydrocarbon and lower nitrogen related peak shifts than the HEMA + CDI samples (Figure 2e and Table 2). PCA analysis comparing ToF-SIMS spectra from each step of the HEMA + CDI + LB DMPE grafting and sonication process clearly separated each of the stages with PCs 1 and 2 capturing 89% and 6% of the variation in the positive ion spectra, respectively. Figure 6a shows the resulting scores plot for the first two PCs. Not surprisingly, the greatest variation is seen between the

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Figure 6. (a) Scores on PC 1 and 2 and loading plots for (b) PC 1 and (c) PC 2 generated from PCA analysis of positive ion ToF-SIMS spectra from (b) HEMA + CDI, (2) HEMA + CDI + 20mN/m DMPE and (9) HEMA + CDI + 20mN/m DMPE (EtOH sonicated) surfaces.

spectra from HEMA + CDI and the HEMA + CDI + LB DMPE samples (PC 1). Spectra from samples sonicated

after LB transfer scored between the two extremes and were separated from the other two data sets in PC 2.

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Figure 7. (a) Scores and (b) loading plots for PCA analysis of positive ion ToF-SIMS spectra from (2) HEMA + CDI + 20mN/m DMPE (EtOH sonicated) and (9) HEMA + CDI + 20mN/m DMPE surfaces.

Analysis of the loading plots illustrated that separation in PC 1 (Figure 6b) was due to the presence of lipid related fragments (negative loadings) and nitrogen containing fragments (positive loadings), while in PC 2 (Figure 6c) variation in the intensity of hydrocarbon and nitrogen containing peaks (positive loadings) and hydrocarbon and oxygen containing peaks (negative loadings) separated the sonicated samples from the other two sets of data. The initial approach was able to separate each of the sets of spectra, but the presence of the HEMA + CDI data in the PCA complicated the interpretation of the loading plot. Having established that the sonicated samples did not revert to the HEMA + CDI chemistry, a second PCA analysis was undertaken using only the spectra from the lipid LB transfer surfaces. Figure 7 shows the scores and loading plots for the CDI grafted lipid surfaces before and after sonication. PC 1 captured 94% of the variation in the positive ion data set and clearly separated the two sample sets (Figure 7a). Analysis of the loading plot indicated that the separation was caused by the relative intensities of the lipid and coupling chemistry related fragments (Figure 7b). Comparison of specific regions of the spectra in Figure 8 confirmed that there were lipid related peaks detected on the HEMA + CDI surfaces after sonication. Analysis of the spectra indicated that there was a peak series starting at 495.441 due to ionization of the lipid tails (C31H59O4) seen in the HEMA + CDI + LB DMPE spectra

Figure 8. Comparison of a region of the ToF-SIMS spectra from (a) HEMA + CDI, (b) HEMA + CDI + 20mN/m DMPE, and (c) HEMA + CDI + 20mN/m DMPE (EtOH sonicated) surfaces showing the presence of lipid and CDI related fragments on the surface after EtOH sonication of the grafted lipid films.

and a series starting at 505.37 related to fragmentation of HEMA + CDI (C26H53O7N2). After sonication, both sets of peaks were detected in spectra, indicating that there was still lipid present at the surface of the sample. It is interesting to note that, prior to sonication, there was

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evidence of the whole lipid seen in the ToF-SIMS spectra, but after sonication, only smaller fragments of the lipid, related to fragments of the tails, were evident. This reduction in the molecular ion yield could be due to two factors: a reduction in the total lipid present on the surface, reducing the probability that molecular ions are generated, and/or the presence of the covalent bond between the lipid headgroup and the HEMA, making it harder to generate the intact lipid molecular ions. Covalent binding of a species to the surface has been shown to result in a significant decrease in molecular ion yields in MALDIMS experiments.28 Comparison of all of the samples before and after sonication in ethanol indicated that the presence of the covalent coupling via CDI was critical in retaining the lipid at the surface in a range of conditions. If we compare the two lipid application techniques, XPS data indicate that the LB transfer is able to transfer and retain slightly more lipid on the HEMA + CDI surface. Comparison of the ToF-SIMS data is complicated by evidence of lipid association with the HEMA without covalent coupling. In other experiments,29 the anchoring of the DMPE monolayer to the HEMA substrate by the grafting some of the lipids has been shown to result in monolayers with greater stability in solution compared to glass and untreated HEMA. It is interesting to note that no lipid was detected by XPS or ToF-SIMS on the LB transferred untreated HEMA surface after sonication, suggesting that lipid may have been physically trapped in the swollen HEMA network during the extended time allowed for lipid adsorption from solution. This conclusion is supported by contact lens literature11,15 where lipid uptake into the bulk HEMA material has been well established. Ordering of the lipid layer may also be expected to influence the efficacy of the reaction chemistry. In solution the random orientation of the lipid molecules and fluid kinetics limit the probability that the headgroups will be in a suitable position to react with the surface at any point in time. Once bound to the surface, the steric hindrance from the lipid tails act to limit the access of subsequent lipids as they approach the reactive groups on the surface. In addition to this, imidazole carbamate is known to hydrolyze in water,30 limiting the amount of time the HEMA + CDI surface will be active. If, instead, the lipids are transferred to the surface as an oriented closely packed monolayer (i.e., LB deposition), then all the headgroups are in position to be covalently coupled to the surface and the prepacking of the layer ensures that there is little effect from steric hindrance. In this case the density of the coupling is limited only by the number of (29) Halter, M.; Vogel, V. Manuscript in preparation. (30) Hermanson, G. T. Bioconjugate techniques; Academic Press: London, 1996.

McArthur et al.

reactive groups on the surface. XPS data from this study support this hypothesis, with the LB deposited surface appearing to retain a slightly higher level of carbon after sonication compared to the solution adsorbed surface. The preparation of the lipid monolayer via LB deposition has several advantages over a solution phase approach. The orientation and packing of the lipid layer are critical if these systems are to act as a real model for protein/lipid interactions. Gaps or any irreproducibility in the coatings will severely limit their efficacy and predictability. The lipid orientation is also important if this type of structure is to be used as the inner leaflet of a supported lipid bilayer. LB deposition allows a one-step process for the preparation of the inner leaflet with the density and fluidity of the layer being controlled via the transfer parameters, the density of reactive groups on the substrate and the proportion of reactive lipid present in the transfer mixture. Conclusions XPS and ToF-SIMS have been used to characterize the deposition and attachment of DMPE lipid layers onto polymeric HEMA substrates. The important results from these experiments were as follows: 1. LB transfer produced higher lipid concentrations on the HEMA surfaces than solution phase deposition. 2. Ethanol sonication essentially removed all lipid deposited onto the unmodifed HEMA substrate. 3. Independent of the methods used to apply the lipid to the surface, a small, but significant, amount of lipid remained on the CDI-activated HEMA surfaces after ethanol sonication. 4. LB deposition of DMPE onto a CDI-activated HEMA substrate is an effective method for preparing covalently coupled lipid layers. Acknowledgment. We gratefully acknowledge the support of the National ESCA and Surface Analysis Center for Biomedical Problems (NESAC/Bio, NIH Grant RR01296) through the National Center for Research Resources and the University of Washington Engineered Biomaterials (UWEB, Grant NSF EEC 9529161). We thank Oliver Dannenberger, Paul J. Valint, and Joeseph McGee for their contributions to the development of protocols for the silanization of glass and HEMA formulations used in the polymer supports and Sarah Keller and Ben Stottrup for use of their microscope to image monolayers at the air/water interface. Supporting Information Available: Table providing assignments for the ToF-SIMS peaks and figures showing the ToF-SIMS and PCA results for adsorbed DMPE on unmodified and CDI modified HEMA surfaces. This material is available free of charge via the Internet at http://pubs.acs.org. LA026928H