Layer by Layer Self-Assembled Polyelectrolyte Multilayers with

Layer by Layer Self-Assembled Polyelectrolyte Multilayers with Embedded Phospholipid ... We describe a method to embed phospholipid vesicles into poly...
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Langmuir 2004, 20, 4835-4839

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Layer by Layer Self-Assembled Polyelectrolyte Multilayers with Embedded Phospholipid Vesicles Marc Michel,† Dominique Vautier,‡ Jean-Claude Voegel,‡ Pierre Schaaf,*,† and Vincent Ball† Institut Charles Sadron, Centre National de la Recherche Scientifique, Unite´ Propre 22, 6 rue Boussingault, 67083 Strasbourg Cedex, France, and Institut National de la Sante´ et de la Recherche Me´ dicale, Unite´ 595, Faculte´ de Me´ decine, 11 rue Humann, 67085 Strasbourg Cedex, France Received January 30, 2004. In Final Form: March 26, 2004 We describe a method to embed phospholipid vesicles into polyelectrolyte multilayers built up by the alternate deposition of polyanions and polycations. Before deposition, the vesicles are rigidified by polycation adsorption onto their surface avoiding their fusion once deposited on the multilayer surface. The vesicles adsorb to form a compact and “hard” monolayer as imaged by atomic force microscopy. The thickness of the adsorbed vesicle layer, of the order of 250 nm, is very close to the diameter of the vesicles in solution. This work should open the route to the buildup of multilayer films containing phospholipid vesicles that could act as “reservoirs” for drugs or enzymatic nanoreactors.

I. Introduction Among the different coating methods of solid surfaces, the layer by layer (LBL1) deposition of oppositely charged polyelectrolytes is of increasing interest.2-4 This is explained by different reasons: Polyelectrolyte multilayers can be deposited on almost any kind of surface, whatever its shape, provided it carries a nonzero surface charge density.2-4 The multilayers can easily be functionalized and even multifunctionalized, with potential applications ranging from electro-optical devices5,6 and separation membranes7,8 to active biomaterial coatings9,10 and nanoreactors.10,11 Up to now, functionalization was achieved by the incorporation of active molecules such as proteins12,13 or DNA,14 of inorganic particles such as carbon * Corresponding author. E-mail: [email protected]. † Institut Charles Sadron. ‡ Institut National de la Sante ´ et de la Recherche Me´dicale. (1) List of abbreviations used in this paper: LBL, layer by layer; unmodifiedLUV, unmodified large unilamellar vesicles; modified LUV, modified large unilamellar vesicles; QCM-D, quartz crystal microbalance with dissipation; POPG, 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylDL-glycerol; POPC, 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylcholine; PEI, branched polyethylene imine; PAH, polyallylamine; PGA, poly-L-glutamic acid; PDL, poly-(D-lysine); PL, pair of layers, i.e., a polyelectrolyte film made from the consecutive adsorption of two oppositely charged polyelectrolytes. (2) Decher, G. Science 1997, 277, 1232-1237. (3) Hammond, P. T. Curr. Opin. Colloid Interface Sci. 2000, 4, 430442. (4) Bertrand, P.; Jonas, A. M.; Laschewsky, A.; Legras, R. Macromol. Rapid. Commun. 2000, 21, 319-348. (5) Fou, A. C.; Onitsuka, O.; Ferreira, M.; Rubner, M. F.; Hsieh, B. R. J. Appl. Phys. 1996, 79, 7501-7509. (6) Eckle, M.; Decher, G. Nano Lett. 2001, 1, 45-49. (7) Stanton, B. W.; Harris, J. J.; Miller, M. D.; Bruening, M. L. Langmuir 2003, 19, 7038-7042. (8) Krasemann, L.; Toutianoush, A.; Tieke, B. J. Membr. Sci. 2001, 181, 221-228. (9) Jessel, N.; Schwinte, P.; Falvey, P.; Darcy, R.; Haı¨kel, Y.; Schaaf, P.; Voegel, J. C.; Ogier, J. Adv. Funct. Mater. 2004, 14, 174-182. (10) Chluba, J.; Voegel, J. C.; Decher, G.; Erbacher, P.; Schaaf, P.; Ogier, J. Biomacromolecules 2001, 2, 800-805. (11) Onda, M.; Lvov, Y.; Ariga, K.; Kunitake, T. J. Fermentation Bioeng. 1996, 82, 502-506. (12) Brynda, E.; Houska, M. In Protein Architecture: Interfacing Molecular Assemblies and Immobilization Biotechnology; Mo¨hwald, H., Ed.; Marcel Dekker: New York, 2000; pp 251-285. (13) Lvov, Y.; Ariga, K.; Ichinose, I.; Kunitake, T. J. Am. Chem. Soc. 1995, 117, 6117-6123.

nanotubes,15 or of chemically modified polyelectrolytes10 or simply active polyelectrolytes16 into the multilayers during their buildup. Microcapsules,17 selective membranes,16 and active biocoatings9,12,18,19 were prepared along this line. In the field of biomaterials, films possessing antiadhesive20 and antimicrobial properties were, for example, reported. Positively charged chitosan and anionic dextran sulfate films presented anticoagulation or procoagulation properties depending upon the nature of the final layer.21 One could also prepare films with antiinflammatory properties by inserting anti-inflammatory drugs.9 It was also shown that proteins embedded in multilayers are recognized by cells which respond to their presence.22 However, the deposition of intact vesicles into polyelectrolyte multilayers could greatly enhance the loading capacity of the multilayers for potential drug release and could allow encapsulation of biomolecules into their native aqueous environment without a direct contact with the polyelectrolytes constituting the multilayer. However, the lack of stability of the vesicles made from phospholipids does not allow the vesicle deposition without spontaneous disruption. A first approach toward the deposition of intact vesicles on surfaces of poly(vinyl sulfate)/poly(diallyldimethylammonium chloride) multilayers as well as their embedding in such multilayer architectures was reported by Katagiri et al.23 These authors used vesicles made from N,N-dihexadecyl-N′-(3-triethoxysilyl)propylsuccina(14) Pei, R.; Cui, X.; Yang, X.; Wang, E. Biomacromolecules 2001, 2, 463-468. (15) Mamedov, A.; Kotov, N. A.; Prato, M.; Guldi, D. M.; Wicksted, J. P.; Hirsch, A. Nat. Mater. 2002, 1, 190-194. (16) Rmaile, H. H.; Schlenoff, J. B. J. Am. Chem. Soc. 2003, 125, 6602-6603. (17) Caruso, F.; Trau, D.; Mo¨hwald, H.; Renneberg, R. Langmuir 2000, 16, 1485-1488. (18) Mendelsohn, J. D.; Yang, S. Y.; Hiller, J.; Hochbaum, A. I.; Rubner, M. F. Biomacromolecules 2003, 4, 96-106. (19) Yang, S. Y.; Mendelsohn, J. D.; Rubner, M. F. Biomacromolecules 2003, 4, 987-994. (20) Elbert, D. L.; Herbert, C. B.; Hubell, J. A. Langmuir 1999, 15, 5355-5362. (21) Serizawa, T.; Yamaguchi, M.; Matsuyama, T.; Akashi, M. Biomacromolecules 2000, 1, 306-309. (22) Jessel, N.; Atalar, F.; Lavalle, Ph.; Mutterer, J.; Decher, G.; Schaaf, P.; Voegel, J. C.; Ogier, J. Adv. Mater. 2003, 15, 692-695.

10.1021/la049736q CCC: $27.50 © 2004 American Chemical Society Published on Web 05/13/2004

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Figure 1. Size distribution of the unmodified (0 and dashed line) and modified vesicles (b and full line) as obtained with quasi elastic light scattering.

mide which were stabilized by the formation of a polymerized silica layer at the vesicle surface by spontaneous condensation of triethoxysilyl groups in water. We present here an alternative route to adsorb and embed vesicles made only from phospholipids in polyelectrolyte multilayers without vesicle disruption. Instead of stabilizing the vesicles with an inorganic layer, we use phospholipid vesicles which are stabilized by adsorption of polyelectrolytes. These polyelectrolytes can be of the same nature as those incorporated in the multilayers. II. Materials and Methods Throughout this study, we used 10 mM Tris (tris(hydroxymethyl)aminomethane, Gibco BRL) buffer at pH ) 7.4 with either 0.015 or 0.15 M NaCl (Prolabo, France). The lipids used to prepare the vesicles, all purchased from Sigma and used without further purification, were 1-palmitoyl2-oleoyl-sn-glycero-3-phosphatidylcholine (POPC) [(L-R-lecithin, type XVI-E from egg yolk, Sigma, ref P-3556, lot 22K5223], cholesterol [(5-cholesten-3-ol), Sigma, ref C-8667, lot 050K304], and 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidyl-DL-glycerol (POPG) [β-oleoyl-γ-palmitoyl (C18:1[cis]-9/C16:0), Sigma, ref P-6956, lot 112K5210]. The polyelectrolytes used to build the polyelectrolyte multilayers were branched polyethylene imine (PEI, Aldrich, catalog number 15,188-2, molecular weight ) 750 000 g mol-1), polyallylamine (PAH, Aldrich, catalog number 28,322-3, molecular weight ) 65 000 g mol-1), and poly-L-glutamic acid (PGA, Sigma, ref P-4886, viscosimetric molecular weight ) 17 000 g mol-1). Poly-(D-lysine) (PDL) (Sigma, ref P-4408, lot 020K5940, viscosimetric molecular weight ) 27 200 g mol-1) was used to prepare modified large unilamellar vesicles. Vesicle Preparation. According to Nebel et al.,24 the liposomes were prepared by dissolving a mixture of 50 mg of POPC, 2.5 mg of POPG, and 2.5 mg of cholesterol in 5 mL of chloroform. POPG was used to confer a negative charge to the vesicles at pH 7.4, the pH at which the experiments were performed, and cholesterol was used for its ability to rigidify membranes. The lipid solution was dried under a flux of nitrogen and stored under vacuum overnight. Ten milliliters of 15 mM NaCl, 10 mM Tris buffer at pH 7.4 was then added for lipid hydration. The obtained turbid suspension was then subjected to 15 freeze and thaw cycles and finally extruded 10 times through a 0.22 µm Millex GV membrane (Millipore). This method is known to form unilamellar vesicles,25 and these vesicles will be called unmodified large unilamellar vesicles (unmodifiedLUV). The z average diameter of the vesicles lies around 200 nm with a distribution ranging from 100 to 300 nm (Figure 1), and the zeta potential is -54.9 ( 3.6 mV (Zetasizer 3000 HS, Malvern Instruments). For deposition experiments, the unmodified LUV solutions were (23) (a) Katagiri, K.; Hamasaki, R.; Ariga, K.; Kiguchi, J. Langmuir 2002, 18, 6709-6711. (b) Katagiri, A.; Hamasaki, R.; Ariga, K.; Kiguchi, J. J. Am. Chem. Soc. 2002, 124, 7892-7893. (24) Nebel, S.; Ganz, P.; Seelig, J. Biochemistry 1997, 36, 28532859. (25) Traikia, M.; Warschawski, D. E.; Recouvreur, M.; Carthaud, J.; Devaux, P. F. Eur. Biophys. J. 2000, 29, 184-195.

Michel et al. diluted by a factor of 30 in 15 mM NaCl, 10 mM Tris buffer at pH 7.4 before adsorption on a multilayer ending with PAH. Modified LUVs were prepared from an unmodified LUV solution after a 15-fold dilution in the 15 mM NaCl, 10 mM Tris buffer. This solution was then slowly dropped into the same volume of a 0.5 mg mL-1 PDL solution prepared in the same Tris buffer. Both the PDL chain length (Mw ) 27 200 g mol-1) and its concentration (0.5 mg mL-1) were chosen to be small enough to avoid significant vesicle bridging. Moreover, the ionic strength (15 mM NaCl) of the buffer was also chosen to be small enough to avoid a too strong screening of the repulsive electrostatic forces between identically charged vesicles. Because of the negative charge of the unmodified LUV, PDL is assumed to adsorb onto the vesicles. This was verified by the zeta potential, which became positive with values of the order of +(65.2 ( 3.4) mV after PDL adsorption. Moreover, the size distribution of the vesicles remained comparable to that of unmodified LUVs with diameters ranging typically from 150 to 350 nm (see Figure 1). This behavior was reproducible over more than 10 experiments and clearly shows the absence of significant vesicle aggregation induced by the PDL adsorption. Characterization Methods. We used quartz crystal microbalance with dissipation (QCM-D, Q-Sense-AB, Go¨teborg, Sweden), ellipsometry (PLASMOS SD series) working at constant incidence angle, and atomic force microscopy (AFM, Nanoscope IV, Digital Instruments) to characterize the vesicle incorporation and the film structure. QCM-D is a technique in which a quartz crystal is excited at its fundamental frequency (about 5 MHz) and observation takes place at the first, third, fifth, and seventh overtones (denoted ν and corresponding to 5, 15, 25, and 35 MHz, respectively).26,27 A decrease in ∆f/ν is usually associated, in a first approximation, with an increase of the mass coupled to the quartz. Here, we use QCM-D only for qualitative analysis. Due to the complexity of our films, we do not expect Sauerbrey’s relation to be valid.28 A more refined quantitative analysis would require the use of the viscoelastic model of films developed by Voinona et al.29,30 The validity of such a model for our system is however questionable and would need much further investigation. Recent studies showed nevertheless that QCM-D is able to discriminate between a deposition process in which the vesicles remain intact on the surface and a process in which the deposition is accompanied by a vesicle rupture on the surface and the formation of a lipid bilayer.31-33 Okahata and co-workers also used QCM to investigate the phase transition and the hydration capability of the polar headgroups of the phospholipids. These observations were obtained from the change in ∆f with temperature over a wide temperature range.34,35 Unfortunately, such changes cannot be followed with our experimental setup. It is however expected that our vesicles are highly hydrated due to the hydrophilic nature of the phosphatidylcholine and phosphatidylglycerol end groups. Ellipsometry is a widely known optical technique which allows the measurement of film thickness. We use a PLASMOS SD 2300 (Munich, Germany) instrument working at constant incidence angle. The technique is thus merely sensitive to the optical mass nd where n is the film refractive index and d is its optical thickness. We impose to n a value of 1.465 in order to calculate the film thickness. Our instrument allows only determination of the thickness of films in contact with air. The films were thus always dried before measurement. (26) Rodahl, M.; Kasemo, B. Sens. Actuators, B 1996, B37, 111-116. (27) Ho¨o¨k, F.; Rodahl, M.; Brzezinski, P.; Kasemo, B. J. Colloid Interface Sci. 1998, 208, 63-67. (28) Sauerbrey, G. Z. Phys. 1959, 155, 206-222. (29) Rodahl, M.; Ho¨o¨k, F.; Fredriksson, C.; Keller, C. A.; Kroser, A.; Brzezinski, P.; Voinova, M.; Kasemo, B. Faraday Discuss. 1997, 107, 229-246. (30) Voinova, M. V.; Rodahl, M.; Jonson, M.; Kasemo, B. Phys. Scr. 1999, 59, 391-396. (31) Reimhult, E.; Ho¨o¨k, F.; Kasemo, B. Langmuir 2003, 19, 16811691. (32) Richter, R.; Mukhopadhyay, A.; Brisson, A. Biophys. J. 2003, 85, 3035-3047. (33) Seantier, B.; Breffa, C.; Felix, O.; Decher, G. Nano Lett. 2004, 4, 5-10. (34) Okahata, Y.; Kimura, K.; Ariga, K. J. Am. Chem. Soc. 1989, 111, 9190-9194. (35) Ariga, K.; Okahata, Y. Langmuir 1994, 10, 2272-2276.

Polyelectrolyte Multilayers with Embedded Vesicles Concerning AFM imaging, the QCM-D crystals were rinsed with distilled water in order to avoid sodium chloride crystallization, dried in air, and imaged with silicium nitride cantilevers (k ) 0.03 N/m, Park Scientific). Buildup of the Polyelectrolyte Multilayers. The experiments were performed as follows: For QCM-D experiments, the films were deposited on silicacovered quartz crystals (Q-Sense-AB, Go¨teborg, Sweden). These crystals were cleaned directly in the QCM-D experimental cell with Hellmanex at 2% (v/v) for 30 min, rinsed with deionized water (Milli-Q Plus, Millipore, F ) 18.2 MΩ cm), put in contact with a 0.1 M HCl solution for 10 min, and finally extensively rinsed with the 0.15 M NaCl, 10 mM Tris buffer solution. The films were constructed in situ as follows: a PEI solution at 5 mg mL-1 dissolved in the 0.15 M NaCl, 10 mM Tris buffer was injected in the experimental cell and thus brought in contact with the adsorption surface for 10 min. The same buffer solution, the PGA solution (1 mg mL-1 dissolved in the 0.15 M NaCl, 10 mM Tris buffer), the buffer solution, the PAH solution (1 mg mL-1 dissolved in the 0.15 M NaCl, 10 mM Tris buffer), and again the same buffer solution were successively put in contact with the surface for 10 min. This allowed the formation of a PGA-PAH bilayer. A contact time of 10 min between each solution and the surface was sufficient to reach a constant value in the adsorbed amounts.36 The unmodified LUVs which are negatively charged were adsorbed over a 40 min period on a PEI-(PGA-PAH)2 film, whereas the modified LUVs, which are positively charged, were adsorbed for 30 min on a PEI-(PGAPAH)2-PGA film. As will be seen in the results section, these durations are sufficient to reach steady-state values in both the frequency and dissipation. Before the vesicle deposition, the film was put in contact with a 15 mM NaCl, 10 mM Tris buffer into which the vesicles were suspended. After the vesicle deposition, the surface was rinsed again with the 15 mM NaCl, 10 mM Tris, pH 7.4 buffer and subsequently with the 150 mM NaCl, 10 mM Tris buffer, before continuation of the alternate polyelectrolyte adsorption. In the case of the unmodified LUVs (respectively modified LUVs), we deposited a PAH-(PGA-PAH)2 (respectively (PGA-PAH)2) film on top of the vesicles. For AFM imaging, the PEI-(PGA-PAH)2-unmodifiedLUVsPAH-(PGA-PAH)2 and the PEI-(PGA-PAH)2-PGA-modifiedLUVs-(PGA-PAH)2 modified quartz crystal surfaces were stored under buffer for 1-7 days before imaging under dry conditions with contact mode AFM. Before imaging, the QCM crystals were rinsed with distilled water in order to avoid sodium chloride crystallization, dried in air, and imaged with new cantilevers. For ellipsometry experiments, silicon wafers (Polylabo, Strasbourg, France) were cleaned with a 1:1 CH2Cl2/MeOH mixture and then with a 10 mM H2SO4 solution. They were then extensively rinsed with deionized water and dried under a nitrogen stream. The adsorption time of the polyelectrolyte solutions and the rinsing steps in 150 mM NaCl, 10 mM Tris buffer were all equal to 10 min. These adsorption and buffer rinse steps were made by dipping the silicon wafer into the polyelectrolyte or buffer solutions. Before the adsorption of the modifiedLUVs, a PEI-(PGA/PAH)2-PGA multilayer was made and the layer thickness was measured after each deposition step. To adsorb modified LUVs on a silicon wafer already covered by a PEI-(PGA/PAH)2-PGA multilayer, some cautions were taken. Indeed, one expects the presence of a lipid monolayer at the air/water interface of the vesicle solution. To avoid the deposition of a Langmuir-Blodgett film onto the multilayer during the dipping of the silicon wafers into the solution, we avoided direct exposure of the multilayer with the air/water interface. To this aim, the vesicle solution was injected in large excess into an aqueous solution (15 mM NaCl, 10 mM Tris, pH 7.4) in which the silicon wafer was already immersed. After 30 min of contact, the lipids in the solution at the air/water interface were eliminated by addition of a large excess of 15 mM NaCl, 10 mM Tris buffer. We used a similar deposition and rinsing procedure for the deposition of the additional PGA and PAH layers. The wafer was withdrawn from the aqueous solution once the first PGA/PAH (36) Boulmedais, F.; Ball, V.; Schwinte, P.; Frisch, B.; Voegel, J.-C.; Schaaf, P. Langmuir 2003, 19, 440-445.

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Figure 2. Adsorption kinetics followed by QCM-D of unmodified LUVs (dashed line) onto a PEI-(PGA-PAH)2 multilayer and of modified LUVs (solid line) onto a PEI-(PGA-PAH)2PGA polyelectrolyte multilayer. Panel A: normalized frequency change. Panel B: corresponding changes in dissipation. The time ∆t ) 0 corresponds to the injection of the large unilamellar vesicles. The buffer injection after the vesicle adsorption has reached a plateau is labeled in panels A and B with an arrow. For means of clarity, only the normalized frequency shifts ∆f/ν corresponding to the third harmonic (ν ) 3, 15 MHz) of the crystal are represented. The inset of panel A represents the normalized frequency shift corresponding to ν ) 3 during the buildup of the PEI-(PGA-PAH)2-unmodifiedLUVs-PAH(PGA-PAH)2 film. The arrows labeled with C correspond to the injection of the polycation, and the arrows labeled with A to the injection of the polyanion.

bilayer was deposited on top of the vesicle layer and the last PGA/PAH layer pair was deposited by the dipping method described previously.

III. Results and Discussion We first used QCM-D to follow the construction of a PEI-(PGA-PAH)2 film. The systematic decrease of ∆f/ν as the film is brought in contact with a polyelectrolyte solution demonstrates the regular film buildup (inset of Figure 2A). Previous studies of the PGA/PAH multilayer system showed that the mass and thickness of these films grow exponentially with the number of deposited bilayers.36 The thickness of a PEI-(PGA-PAH)2 film is 20 nm as found by optical waveguide lightmode spectroscopy (OWLS) (in situ measurements, data not shown) and 4 nm as found by ellipsometry. The large difference between the thickness measured in solution by OWLS and in the dry state by ellipsometry indicates that the multilayer is highly hydrated. The unmodified LUVs which are negatively charged at pH 7.4 were then brought in contact with this PAH-ending film which is expected to be positively charged. Figure 2 shows that the frequency shift decreases monotonically and the dissipation increases correlatively with time while the opposite qualitative evolutions are observed during the rinsing step. Reimhult et al.31 and Richter et al.32 performed similar experiments in which unilamellar egg-yolk phosphatidylcholine vesicles

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Figure 3. AFM images of a PEI-(PGA-PAH)2-unmodifiedLUVs-PAH-(PGA-PAH)2 film. Panel A: deflection image over a 10 × 10 µm2 surface. The root-mean-square roughness over this area is 88 nm. Panel B: 3D view of the same surface over a 2 × 2 µm2 area. Arrows indicate aggregates probably formed by fusion of several unmodified LUVs.

were directly deposited on a quartz crystal whose surface chemistry was previously modified. They found that signal evolutions similar to the one observed in our study are characteristic of vesicle adsorption without rupture. A vesicle deposition followed by a vesicle rupture would first lead to a decrease of ∆f/ν, a subsequent increase and stabilization of the signal. The increase in ∆f/ν is due to the loss of water contained in the vesicles during their rupture. The reverse evolution is observed for the dissipation. In the case of unmodified LUVs, the increase of ∆f/ν and the correlative decrease of the dissipation that we observe during the rinsing stage could thus be due to partial vesicle rupture or vesicle desorption from the multilayer. After vesicle deposition, the multilayer buildup was pursued by further deposition of a (PAH-PGA)2 pair of layers. The regular decrease of ∆f/ν (see inset of Figure 2A) indicates the continuation of the multilayer buildup. Similar results were obtained for the two other harmonic frequencies (25 and 35 Hz). After 1 or 7 days of storage in the 150 mM NaCl, 10 mM Tris buffer, the film was then rinsed with distilled water, dried, and imaged by AFM in the contact mode (Figure 3). No difference could be observed for the samples imaged after 1 and 7 days. Clearly, the surface is covered with vesicles but their shapes are irregular. One observes a great density of defects and inhomogeneities on this surface. Some particles seem to be formed by the fusion of smaller vesicles (see arrows in Figure 3). In addition, the characteristic size of the vesicles is of the order of

Michel et al.

400-1000 nm. These values have to be compared to the 200 nm corresponding to the mean size of unmodified LUVs measured in solution (Figure 1). They indicate that the vesicles adhere on the surface but that a fraction of them are not stable enough to withstand either the interaction with the polyelectrolyte cushion, with the polyelectrolytes deposited on the top of their surface, or the drying process and undergo fusion. To increase the mechanical stability of the vesicles and hence the homogeneity of the covered surface, we adsorbed a layer of a relatively small (Mw ) 27 200 g mol-1) PDL chains on the unmodified LUV surfaces. According to Ge et al., such modified LUVs should be quite stiffer37 than the unmodified LUVs and one should be able to adsorb them on a polyanion-ending multilayer film without vesicle fusion. These modified LUVs were deposited on a PEI(PGA-PAH)2-PGA film. Similarly to the unmodified LUVs, QCM-D experiments show that the frequency shift decreases and correlatively the dissipation increases monotonically during the deposition of modified LUVs on the multilayer (Figure 2). The plateau value in the frequency shift is identical to that corresponding to unmodified LUVs, but it is reached much more rapidly. Moreover, the dissipation increase is much smaller than that observed during the deposition of unmodified LUVs. This indicates that, while the deposited mass detected by QCM-D is similar for modified LUVs and unmodified LUVs, the film appears much more rigid with modified LUVs as expected from the rigidification of the vesicles by the PDL layer. The slower time evolution of ∆f/ν and D during adsorption of unmodified LUVs seems thus to indicate the occurrence of a slow restructuration over the surface, such as vesicle fusion concomitantly to vesicle adsorption. Also in marked difference with unmodified LUVs is the absence of frequency shift and dissipation change during and after the rinsing step of modified LUVs. All these behaviors suggest the absence of rupture or fusion of the modified LUVs during both the deposition and rinsing steps. As for the unmodified LUVs, we could deposit an additional (PGA-PAH)2 film on top of this vesicle layer as indicated by an increase of the frequency shift similar to that observed for the unmodified LUVs (data not shown). These films, deposited on the QCM-D crystal surfaces, were stored in buffer for 1 or 7 days and were then imaged by AFM in air after intense water rinse and drying with a nitrogen flow (Figure 4). The surface appears very homogeneously covered with vesicles in a compact monolayer. The spherical morphology of the modified LUVs is maintained, and the diameters of the deposited particles lie between 100 and 400 nm (Figure 4A). This is comparable to or only slightly larger than the size of the modified LUVs in solution. The small increase in the observed size could be due to a small “flattening” of the spherical vesicles consecutive to their interaction with the surface or after further polyelectrolyte deposition on top of the vesicle layer. There exist almost no morphological indications pointing toward vesicle fusion. These observations are in marked contrast with those obtained for the surfaces covered with unmodified LUVs where most vesicles are strongly deformed, possess irregular morphologies (Figure 3A), and have a much larger size. Finally, one can also notice that both the multilayers containing unmodified LUVs and modified LUVs appeared very “hard” when imaged by AFM in the contact mode. We could apply on the cantilever the maximum force allowed by the apparatus without damaging the sample. (37) Ge, L.; Mo¨hwald, H.; Li, J. Colloids Surf., A 2003, 221, 49-53.

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Figure 5. Evolution of the film thickness measured by ellipsometry at different steps of the buildup of a PEI-(PGAPAH)2-PGA-modifiedLUVs-(PGA-PAH)2 multilayer. For the sake of clarity, a PGA-PAH pair of layers is denoted by PL. For example, the point corresponding to PEI-PL2 corresponds to the thickness of the film PEI-(PGA-PAH)2 measured after the final PAH layer deposition and drying. The film is then rehydrated, and the buildup is continued for subsequent measurements.

Figure 4. AFM images of a PEI-(PGA-PAH)2-PGAmodifiedLUVs-(PGA-PAH)2 film. Panel A: deflection image over a 10 × 10 µm2 surface. The root-mean-square roughness over this area is 116 nm. Panel B: 3D view of the same surface over a 2 × 2 µm2 area.

waveguide lightmode spectroscopy. However, the vesicles are expected to be filled with water so that the refractive index of the film should be closer to 1.33. Ellipsometry being mainly sensitive to the product nd, the film thickness should then be closer to 240-250 nm. Neglecting the contribution of the polyelectrolyte layers deposited on the vesicles, this thickness is in full agreement with the 200-300 nm size of the modified LUVs measured in solution. This result clearly demonstrates that the vesicles do not rupture to form a lipid bilayer when adsorbing on the multilayer surface. IV. Conclusions

Although AFM gives a topological image of the film, the technique does not provide a measurement of the thickness of the vesicle layer. We used ellipsometry for this purpose. Figure 5 represents the evolution of the film thickness during its buildup. The vesicle deposition and the subsequent deposition of a (PAH-PGA) film lead to an increase of (220 ( 20) nm in the thickness. This is obtained by assuming a film refractive index of 1.465, which is reasonable for the polyelectrolyte multilayer itself according to previous studies performed by means of optical

In this work, we show that it is possible to embed stable phospholipid vesicles into polyelectrolyte multilayers. This is possible by using phospholipid vesicles stabilized through polycation adsorption which cover the surface in a very homogeneous and compact way. This work opens the route to the buildup of multilayer films containing vesicles that could act as “reservoirs” for drugs or enzymatic nanoreactors, which is our main goal. LA049736Q