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Surface Analysis of Erodible Multilayered Polyelectrolyte Films: Nanometer-Scale Structure and Erosion Profiles Nathaniel J. Fredin, Jingtao Zhang, and David M. Lynn* Department of Chemical and Biological Engineering, University of WisconsinsMadison, 1415 Engineering Drive, Madison, Wisconsin 53706-1607 Received March 5, 2005. In Final Form: April 15, 2005 Atomic force microscopy (AFM) and scanning electron microscopy (SEM) coupled with ellipsometry have been used to characterize the microscale and nanoscale structures of erodible multilayered films fabricated from degradable polyamine 1 and either sodium poly(styrene sulfonate) (SPS) or plasmid DNA. Striking differences were found in the topography, structures, and erosion profiles of these two materials upon incubation in PBS buffer at 37 °C. For films fabricated from SPS, AFM data are consistent with an erosion process that occurs uniformly without the generation of holes or pits over large, micrometer-scale areas. By contrast, films fabricated from plasmid DNA undergo structural rearrangements to present surfacebound particles ranging in size from 50 to 400 nm. Additional characterization of these particulate structures by SEM suggested that they are interpenetrated with or fused to underlying polyelectrolyte layers on the silicon surface, providing a potential mechanism to manipulate the adhesive forces with which these particles are bound to the surface. The erosion profile observed for polymer 1/SPS films suggests that it may be possible to design assemblies that release two film components with well-defined release kinetics. In the context of gene delivery, the presentation of condensed DNA as nanoparticles at these surfaces may be advantageous with respect to stimulating the internalization and processing of DNA by cells. A quantitative understanding of the factors influencing the fabrication, structure, and erosion profiles of these materials will be useful for the design of multilayered assemblies for specific applications in which controlled film erosion or the release of therapeutic materials is desired.
Introduction The alternating, layer-by-layer adsorption of polycations and polyanions on surfaces is a versatile method for the fabrication of multilayered thin films.1-4 The conformal deposition of strategic combinations of natural and synthetic polyelectrolytes on a variety of different substrates has yielded nanostructured assemblies of interest in numerous biotechnical contexts.5-16 The primary benefits of these materials in biological applications often stem as much from the stability of these materials under (1) Decher, G. Science 1997, 277, 1232-1237. (2) Peyratout, C. S.; Dahne, L. Angew. Chem., Int. Ed. 2004, 43, 3762-3783. (3) Bertrand, P.; Jonas, A.; Laschewsky, A.; Legras, R. Macromol. Rapid Commun. 2000, 21, 319-348. (4) Hammond, P. T. Adv. Mater. 2004, 16, 1271-1293. (5) Yu, A. M.; Liang, Z. J.; Caruso, F. Chem. Mater. 2005, 17, 171175. (6) Berg, M. C.; Yang, S. Y.; Hammond, P. T.; Rubner, M. F. Langmuir 2004, 20, 1362-1368. (7) Ghan, R.; Shutava, T.; Patel, A.; John, V. T.; Lvov, Y. Macromolecules 2004, 37, 4519-4524. (8) Richert, L.; Boulmedais, F.; Lavalle, P.; Mutterer, J.; Ferreux, E.; Decher, G.; Schaaf, P.; Voegel, J. C.; Picart, C. Biomacromolecules 2004, 5, 284-294. (9) Mendelsohn, J. D.; Yang, S. Y.; Hiller, J.; Hochbaum, A. I.; Rubner, M. F. Biomacromolecules 2003, 4, 96-106. (10) Michel, M.; Vautier, D.; Voegel, J. C.; Schaaf, P.; Ball, V. Langmuir 2004, 20, 4835-4839. (11) Salloum, D. S.; Schlenoff, J. B. Biomacromolecules 2004, 5, 10891096. (12) Salloum, D. S.; Olenych, S. G.; Keller, T. C. S.; Schlenoff, J. B. Biomacromolecules 2005, 6, 161-167. (13) Thierry, B.; Winnik, F. M.; Merhi, Y.; Silver, J.; Tabrizian, M. Biomacromolecules 2003, 4, 1564-1571. (14) Thierry, B.; Winnik, F. M.; Merhi, Y.; Tabrizian, M. J. Am. Chem. Soc. 2003, 125, 7494-7495. (15) Gergely, C.; Bahi, S.; Szalontai, B.; Flores, H.; Schaaf, P.; Voegel, J. C.; Cuisinier, F. J. G. Langmuir 2004, 20, 5575-5582. (16) Thierry, B.; Kujawa, P.; Tkaczyk, C.; Winnik, F. M.; Bilodeau, L.; Tabrizian, M. J. Am. Chem. Soc. 2005, 127, 1626-1627.
physiological conditions12,16-18 as from the ease with which the thickness, internal composition, and surface properties of these materials can be tailored on the nanometer scale.1-4 There has been increasing recent interest, however, in the design of multilayered films and less stable assemblies for applications in the areas of drug delivery and controlled release. Of particular interest has been the deposition of films onto microcrystalline19-23 and nanocrystalline24 drug “cores” to slow the dissolution and diffusion of small molecules into aqueous environments. Other groups have developed strategies for the reversible absorption of small-molecule dyes into the bulk of a multilayered film.16,25-29 We30,31 and others29,32-43 have focused on the design of polyelectrolyte assemblies that (17) Richert, L.; Lavalle, P.; Vautier, D.; Senger, B.; Stoltz, J. F.; Schaaf, P.; Voegel, J. C.; Picart, C. Biomacromolecules 2002, 3, 11701178. (18) Tryoen-Toth, P.; Vautier, D.; Haikel, Y.; Voegel, J. C.; Schaaf, P.; Chluba, J.; Ogier, J. J. Biomed. Mater. Res. 2002, 60, 657-667. (19) Dai, Z. F.; Heilig, A.; Zastrow, H.; Donath, E.; Mohwald, H. Chem.Eur. J. 2004, 10, 6369-6374. (20) Shi, X. Y.; Caruso, F. Langmuir 2001, 17, 2036-2042. (21) Ai, H.; Jones, S. A.; de Villiers, M. M.; Lvov, Y. M. J. Controlled Release 2003, 86, 59-68. (22) Antipov, A. A.; Sukhorukov, G. B.; Donath, E.; Mohwald, H. J. Phys. Chem. B 2001, 105, 2281-2284. (23) Qiu, X. P.; Leporatti, S.; Donath, E.; Mohwald, H. Langmuir 2001, 17, 5375-5380. (24) Zahr, A. S.; de Villiers, M.; Pishko, M. V. Langmuir 2005, 21, 403-410. (25) Burke, S. E.; Barrett, C. J. Macromolecules 2004, 37, 53755384. (26) Chung, A. J.; Rubner, M. F. Langmuir 2002, 18, 1176-1183. (27) Quinn, J. F.; Caruso, F. Langmuir 2004, 20, 20-22. (28) Hiller, J.; Rubner, M. F. Macromolecules 2003, 36, 4078-4083. (29) Sukhishvili, S. A.; Granick, S. Macromolecules 2002, 35, 301310. (30) Vazquez, E.; Dewitt, D. M.; Hammond, P. T.; Lynn, D. M. J. Am. Chem. Soc. 2002, 124, 13992-13993. (31) Zhang, J.; Chua, L. S.; Lynn, D. M. Langmuir 2004, 20, 80158021. (32) Cho, J.; Caruso, F. Macromolecules 2003, 36, 2845-2851.
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disassemble, disintegrate, or erode in ways that provide a mechanism for the controlled release of the polyelectrolyte components that serve as the structural elements of these materials, that is, those polymers that are incorporated directly into the film structure. Several groups have used changes in environmental conditions such as pH or ionic strength to trigger the disassembly of multilayered films fabricated from strong and/or weak polyelectrolytes.29,32-34,38,41,42 Anzai and coworkers have fabricated assemblies that disintegrate upon the addition of small molecules that disrupt specific receptor/ligand interactions in a multilayered film.36,37 In a separate approach, Akashi and co-workers reported the enzymatic degradation of layer-by-layer films fabricated from chitosan/dextran sulfate39 or DNA/poly(diallyldimethylammonium chloride),40 and, more recently, Etienne et al. reported the enzymatic degradation of multilayered assemblies fabricated from hyaluronic acid and chitosan.35 We recently described the fabrication of multilayered assemblies incorporating a hydrolytically degradable synthetic polycation.30,31 We reported in an initial publication that films fabricated from polymer 1 and sodium
poly(styrene sulfonate) (SPS) erode gradually upon incubation in phosphate-buffered saline (PBS),30 and we recently extended this work to the controlled release of plasmid DNA.31 These erodible assemblies could present structural frameworks for the local or noninvasive delivery of DNA or other therapeutic macromolecules43 from the surfaces of implantable materials and biomedical devices. In many of the examples above, film disintegration is triggered by an event such as a change in pH or the addition of a small molecule that disrupts the bulk structure of a film. In these cases, film disruption generally occurs over relatively short time periods (i.e., either instantaneously or within several minutes or hours) that provide an obstacle to the detailed characterization of physical erosion mechanisms.29,32,34,36,37,39,41 By contrast, films fabricated from SPS and polymer 1 erode and release material over a period of ∼2 days under physiological conditions.30,31 In addition to providing for more extended periods of release, therefore, these degradable materials also offer an opportunity to characterize physically the nature of the erosion process. Here, we report the characterization of the physical morphology and nanometer-scale structure of 80-100 nm thick multilayered films (33) Dubas, S. T.; Farhat, T. R.; Schlenoff, J. B. J. Am. Chem. Soc. 2001, 123, 5368-5369. (34) Dubas, S. T.; Schlenoff, J. B. Macromolecules 2001, 34, 37363740. (35) Etienne, O.; Schneider, A.; Taddei, C.; Richert, L.; Schaaf, P.; Voegel, J. C.; Egles, C.; Picart, C. Biomacromolecules 2005, 6, 726-733. (36) Inoue, H.; Sato, K.; Anzai, J. Biomacromolecules 2005, 6, 2729. (37) Sato, K.; Imoto, Y.; Sugama, J.; Seki, S.; Inoue, H.; Odagiri, T.; Hoshi, T.; Anzai, J. Langmuir 2005, 21, 797-799. (38) Schuler, C.; Caruso, F. Biomacromolecules 2001, 2, 921-926. (39) Serizawa, T.; Yamaguchi, M.; Akashi, M. Macromolecules 2002, 35, 8656-8658. (40) Serizawa, T.; Yamaguchi, M.; Akashi, M. Angew. Chem., Int. Ed. 2003, 42, 1115-1118. (41) Sukhishvili, S. A.; Granick, S. J. Am. Chem. Soc. 2000, 122, 9550-9551. (42) Kharlampieva, E.; Sukhishvili, S. A. Langmuir 2003, 19, 12351243. (43) Wood, K. C.; Boedicker, J. Q.; Lynn, D. M.; Hammond, P. T. Langmuir 2005, 21, 1603-1609.
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constructed from polymer 1 and either SPS or plasmid DNA as a function of full or partial erosion under physiological conditions. We find striking differences in the surface morphologies and erosion behaviors of films fabricated from these two different polyanions. For films fabricated using SPS as a polyanion, ellipsometry and atomic force microscopy (AFM) data are consistent with a gradual erosion process that occurs in a uniform, topdown manner without the generation of significant holes, pits, or cracks on the nanometer or micrometer length scales. This erosion profile suggests the potential application of these materials to the design of films that could provide control over the sequential release of multiple polyanionic species. By contrast, films fabricated from polymer 1 and plasmid DNA undergo striking changes in surface morphology under physiologically relevant conditions that may influence significantly the range of DNA release profiles or applications for which these films are suited. Materials and Methods General Considerations. Silicon substrates (e.g., 0.5 × 2.0 cm) were cleaned with acetone and water and then dried under a stream of compressed air passed through a 0.4-µm filter. Surfaces were then activated by etching in an oxygen plasma for 5 min (Plasma Etch, Carson City, NV). Ellipsometric thicknesses of films deposited on silicon substrates were determined using a Gaertner LSE Stokes ellipsometer (632.8 nm, incident angle ) 70°). Data were processed using the Gaertner Ellipsometer Measurement Program software package. Relative thicknesses were calculated by assuming an average refractive index of 1.55 for the multilayer films. Thicknesses were determined in at least four different standardized locations on each substrate and are presented as an average (with standard deviation) for each substrate. All films were dried under a stream of filtered compressed air prior to measurement. For the characterization of surface morphology by scanning electron microscopy (SEM), an accelerating voltage of 3 kV was used to obtain images on a LEO DSM 1530 scanning electron microscope. Samples were coated with a thin layer of gold using a sputterer (30 s at 45 mA, 50 mTorr) prior to analysis. Materials. Test grade n-type silicon wafers were purchased from Si-Tech, Inc. (Topsfield, MA). Linear poly(ethylene imine) (LPEI, MW ) 25 000) was obtained from Polysciences, Inc. (Warrington, PA). SPS (MW ) 70 000) and sodium acetate buffer were purchased from Aldrich Chemical Co. (Milwaukee, WI). Polymer 1 (Mn ≈ 20 000) was synthesized as previously described.44 Plasmid DNA [pEGFP-N1 or pDsRed2-N1 (4.7 kb), >95% supercoiled] was purchased from Elim Biopharmaceuticals, Inc. (San Francisco, CA). Sodium phosphate monobasic and sodium phosphate dibasic heptahydrate were purchased from Fisher, and sodium chloride was purchased from EM Science. All commercial polyelectrolytes were used as received without further purification. Deionized water (18 MΩ) was used for washing steps and to prepare all polymer solutions. PBS buffer was prepared either (1) by diluting commercially available concentrate (EM Science) and adjusting the pH to 7.4 with 1.0 M HCl or NaOH or (2) as a 10 mM stock at the desired pH by adjusting the ratio of dibasic to monobasic phosphate salt solutions; ionic strength was varied by adding sodium chloride at the desired concentration. All buffers and polymer solutions were filtered through a 0.2-µm membrane syringe filter prior to use. Preparation of Polyelectrolyte Solutions. Solutions of polymer 1 used for dipping (5 mM with respect to the molecular weight of the polymer repeat unit) were prepared in sodium acetate buffer (100 mM, pH 5.0) and filtered through a 0.2-µm membrane syringe filter prior to use. Solutions of LPEI and SPS used for the fabrication of LPEI/SPS precursor layers (20 mM with respect to the molecular weight of the polymer repeat unit) were prepared using a 50 mM NaCl solution in 18 MΩ water. (44) Lynn, D. M.; Langer, R. J. Am. Chem. Soc. 2000, 122, 1076110768.
Erodible Multilayered Polyelectrolyte Films LPEI solutions contained 5 mM HCl to aid polymer solubility. SPS solutions used for the deposition of polymer 1/SPS layers (20 mM with respect to repeat unit) were prepared in Milli-Q water containing 0.067 mM HCl. Solutions of plasmid DNA were prepared at 1 mg/mL in sodium acetate buffer and were not filtered prior to use. Fabrication of Multilayered Films. Films were deposited on planar silicon substrates precoated with 10 bilayers of LPEI/ SPS (terminated with a layer of SPS) to ensure a suitably charged surface for the adsorption of polymer 1 as previously described.30,31 Fabrication of multilayered films was conducted using an alternate dipping method according to the following general protocol: (1) Substrates were submerged in a solution of polycation for 5 min. (2) Substrates were removed and immersed in an initial water bath for 1 min followed by a second water bath for 1 min. (3) Substrates were submerged in a solution of polyanion for 5 min. (4) Substrates were rinsed in the manner described above. This cycle was repeated until the desired number of polycation/polyanion bilayers (typically 10 each) had been deposited. This process was fully automated using an automated dipping robot (model DR-3 dipping robot, Riegler & Kirstein GmbH, Berlin, Germany) with the exception of (a) the polymer 1/SPS layers used to observe film morphology during the deposition process (see text) and (b) the polymer 1/plasmid layers in DNA-containing films; in these cases, dipping was performed manually. Following the deposition process, films were allowed to dry in ambient air or were dried under a stream of filtered compressed air. Erosion of Multilayered Films. Experiments designed to investigate the erosion profiles of multilayered polymer 1/SPS or polymer 1/DNA films were performed in the following general manner: Film-coated substrates were placed in a vial containing a sufficient amount of the desired buffer to completely cover the film-coated portion of the substrate. Typically, the samples were incubated in PBS buffer at 37 °C unless otherwise indicated. Samples were removed for analysis at predetermined intervals, rinsed in two 1-min water baths, and dried under filtered compressed air. Film thickness and surface topography were analyzed by ellipsometry and/or AFM, after which the films were placed in a vial containing fresh PBS buffer and returned to the 37 °C incubator. Control experiments established that allowing films to stand dry for several hours between periods of incubation (e.g., for analysis by AFM) did not yield an observable change in erosion rates once they were returned to the incubation medium. Characterization of Surface Topography by AFM. Film topography and surface roughness were obtained from height data imaged in tapping mode on a Nanoscope Multimode atomic force microscope (Digital Instruments, Santa Barbara, CA), using scan rates of 10-20 µm/s to obtain 256 × 256 pixel images. For imaging in air, silicon cantilevers with a spring constant of 40 N/m and a radius of curvature of 9 days to reach the initial thickness of the nonerodible LPEI/SPS foundation layers (Figure 3A, thickness determined by ellipsometry). The surface of an identical and continuously submerged film remained smooth and free of pits and holes over a 2500 µm2 area over this 9-day period (Figure 3B), and the RMS roughness of the film remained less than the average thickness of a polymer 1/SPS bilayer for the duration of the experiment. These results are consistent with the results obtained for dried films above and provide additional evidence that a smooth and well-defined erosion process occurs over relatively large micrometer-scale expanses of the film. It is worth noting that any topographic features of these films smaller than ≈10 nm would be obscured by AFM tip convolution effects.
Influence of Solution pH and Ionic Strength on the Erosion of Polymer 1/SPS Films. Polymer 1 is a cationic polyester that has previously been demonstrated to degrade via ester hydrolysis under physiological conditions (e.g., pH 7.4, 37 °C).44,47-49 The gradual decreases in film thickness described above are consistent with a mechanism of film erosion that is at least partially hydrolytic. However, these experiments do not conclusively rule out contributions from other environmental factors, such as the ionic strength of the incubation medium, that could also contribute to film disruption.33,34,50 Although it is possible to propose erosion mechanisms based exclu(47) Akinc, A.; Lynn, D. M.; Anderson, D. G.; Langer, R. J. Am. Chem. Soc. 2003, 125, 5316-5323. (48) Lynn, D. M.; Anderson, D. G.; Putnam, D.; Langer, R. J. Am. Chem. Soc. 2001, 123, 8155-8156. (49) Lynn, D. M.; Amiji, M. M.; Langer, R. Angew. Chem., Int. Ed. 2001, 40, 1707-1710. (50) Kovacevic, D.; van der Burgh, S.; de Keizer, A.; Stuart, M. A. C. Langmuir 2002, 18, 5607-5612.
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Figure 3. (A) Plot of ellipsometric thickness and RMS surface roughness for a polymer 1/SPS multilayered film incubated and imaged under continuous submersion in PBS buffer at ambient room temperature (see text): (]) optical thickness values determined by ellipsometry; (×) RMS surface roughness of the film determined by AFM. Substrates were precoated with 10 bilayers (∼20 nm) of an LPEI/SPS film prior to experiment. (B) Tapping mode in situ AFM images (50 µm × 50 µm) of the film at various time points during erosion. The number in the upper left-hand corner of each image indicates hours and corresponds to the time points shown in part A. Heights h are approximate and were determined by ellipsometry using a separate, identical film; scale in z direction is 30 nm.
sively on chemical hydrolysis, separate physical erosion mechanisms could be driven simply by polymer desorption as polycation-SPS interactions are broken and replaced by interactions with components of the buffer solution.33,34,50 To examine the influence of changes in ionic strength on the erosion profiles of polymer 1/SPS films, we conducted experiments using phosphate buffer (pH 7.2) adjusted to four different ionic strengths using different concentrations of NaCl (0, 50, 100, and 500 mM). We observed no significant differences in the erosion profiles of films incubated and eroded over this range of conditions (Figure 4A). All four films converged on the same terminal thickness, suggesting that the LPEI/SPS foundation layers remained stable over this range of ionic strengths. We conclude from these experiments that ionic strength does not contribute significantly to the erosion of polymer 1/SPS films in the physiological range.
We note that erosion occurred more slowly in the custom PBS buffers above prepared at pH 7.2 (e.g., 100-150 h, with a rate ranging from 0.9 to 1.4 nm/h for the first 47 h) as compared to the results obtained in preparations of commercial PBS used in earlier experiments prepared at pH 7.4 (e.g., Figure 2; ∼2 nm/h). These results suggest that small differences in pH might profoundly influence erosion rates, and they provide evidence in support of an erosion mechanism based on polymer hydrolysis. To examine further the influence of pH on the erosion of films fabricated from polymer 1, we characterized the erosion profiles of three additional films incubated in PBS prepared at pH values of 5.1, 7.2, and 8.9. Film thickness decreased very rapidly under alkaline conditions, with the thickness of this film decreasing by greater than half during the first hour of incubation (Figure 4B). Erosion occurred far more slowly at acidic pH, requiring >900 h
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Figure 4. (A) Plot of ellipsometric thickness versus time for identical polymer 1/SPS multilayered films incubated at 37 °C in phosphate buffer (pH 7.2) adjusted to 0 mM (×), 50 mM ([), 100 mM (9), and 500 mM (2) NaCl. Substrates were precoated with 10 bilayers of an LPEI/SPS film prior to experiment. (B) Plot of ellipsometric thickness versus time for three identical polymer 1/SPS multilayered films incubated at 37 °C in phosphate buffer adjusted to pH 5.1 ([), 7.2 (9), and 8.9 (2). Substrates were precoated with 10 bilayers (∼20 nm) of an LPEI/SPS film prior to experiment.
under these acidic conditions. These pH/rate data are consistent with the relative rates of acid- and basecatalyzed ester hydrolysis and an erosion mechanism that is at least partially hydrolytic. We note, however, that these pH/rate data do not rule out a physical erosion mechanism involving the pH-dependent deprotonation of polymer 1 at alkaline pH and that conclusions drawn from these data should therefore be made with care. In the absence of direct spectroscopic observation of polymer hydrolysis products in these materials, we prefer to describe these films as “erodible” rather than “hydrolytically degradable” per se. Analysis of Assemblies Fabricated from Polymer 1 and Plasmid DNA. In contrast to films fabricated from SPS, assemblies fabricated from polymer 1 and plasmid DNA are considerably rougher and undergo striking nanometer-scale rearrangements upon incubation under physiologically relevant conditions. We selected monodisperse plasmid DNA constructs [pEGFP-N1 or pDsRed2N1 (4.7 kb), >95% supercoiled] for these studies on the basis of our previous investigations31 and the potential value of these reporter genes in gene delivery applications. The optical thickness profiles of plasmid-containing films increased as a linear function of the number of dipping cycles applied during fabrication, as reported previously.31 However, the optical thicknesses and surface morphologies of these polymer 1/plasmid films varied dramatically as a function of the specific fabrication procedures used. Figure 5 shows representative tapping mode AFM images of 10 µm × 10 µm areas of two plasmid-containing films fabricated from 10 polymer 1/DNA dipping cycles using wash baths containing either (A) deionized water or (B) sodium acetate buffer. Films washed intermittently with deionized water during fabrication (see Materials and Methods) were characterized by extreme surface roughnesses and the formation of irregular surface features on the micrometer scale (Figure 5A; RMS roughness ≈ 45 nm). The globular features observed for these films resemble the “islets” and “islands” recently described for the surface morphology of multilayered poly(lysine)/hyaluronic acid assemblies that grow by an exponential mechanism.51 In contrast to this earlier work, however, we did not observe the coalescence of these irregular features or exponential increases in optical thicknesses after the completion of as many as 20 dipping cycles (data not shown). (51) Picart, C.; Lavalle, P.; Hubert, P.; Cuisinier, F. J. G.; Decher, G.; Schaaf, P.; Voegel, J. C. Langmuir 2001, 17, 7414-7424.
Figure 5. Tapping mode AFM images (10 µm × 10 µm) of films constructed from 10 bilayers of polymer 1 and pEGFP plasmid using (A) deionized water rinse steps and (B) sodium acetate buffer rinse steps during fabrication. Heights h are approximate and were determined by ellipsometry; scales in z direction are 200 nm for each image.
Films fabricated using polymer, plasmid, and wash solutions all prepared using pH 5.0 acetate buffer were significantly smoother, having RMS roughness values on the order of 15 nm. As shown in Figure 5B for a 10 bilayer film, materials prepared using this washing procedure were generally devoid of pits, holes, and the uneven topographical features evident in Figure 5A. The differences in these film morphologies reflect the different pH and/or ionic strength cycles experienced by these assemblies during fabrication, and the data in Figure 5A underscore the need to supplement ellipsometric characterization of layer-by-layer film growth with analytical procedures such as AFM that provide topographic information at micrometer and nanometer length scales. We used smooth polymer 1/plasmid films fabricated using acetate buffer solutions (e.g., Figure 5B) to study the erosion profiles of DNA-containing assemblies by AFM and ellipsometry. In contrast to the smooth topographic profiles observed for polymer 1/SPS films (e.g., Figures 2 and 3), these DNA-containing materials underwent dramatic structural rearrangements upon incubation in PBS buffer. Figure 6 shows representative AFM images and ellipsometric thicknesses of a polymer 1/plasmid film ∼90 nm thick removed from buffer and imaged at 5-h intervals. We observed the formation of small round and oblate particles with diameters ranging from 50 to 400 nm and heights of up to 200 nm after as little as 1 h of incubation in PBS buffer. These features decreased in
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Figure 6. (A) Plot of ellipsometric thickness versus time for a polymer 1/DNA film incubated in PBS buffer at 37 °C. Silicon substrates were precoated with 10 bilayers (∼20 nm) of an LPEI/SPS film prior to experiment. (B) Tapping mode AFM images (10 µm × 10 µm) of a multilayered film fabricated from 10 bilayers of polymer 1 and pEGFP at various time points during erosion in PBS buffer. The number in the upper left-hand corner of each image indicates the number of hours of incubation. Height h was determined by ellipsometry; scale in z direction is 200 nm.
both density and average size over the course of the 20-h incubation period, concomitant with a decrease in the average ellipsometric thickness and an increase in the concentration of DNA observed in solution (reported previously,31 data not shown). As shown in Figure 6A, the optical thickness of films fabricated from plasmid DNA decreased much more rapidly during the first several hours as compared to films fabricated from SPS (e.g., Figure 2), reflecting this dramatic change in the surface structure of these DNA-containing materials. The behavior of these DNA-containing materials is dramatically different from the erosion behavior of films fabricated from polymer 1 and SPS and could have a profound influence on the practical utilization of these materials. For example, we recently demonstrated that macroscopic objects (e.g., quartz slides) coated with polymer 1/pEGFP assemblies 100 nm thick can be used to direct the localized transfection of cells in culture in the absence of any additional transfection agents.52 Polycations are in general used widely as gene delivery agents because they self-assemble with and condense DNA into nanoparticles that can be internalized readily by cells.53 In addition to providing a mechanism for the localization of DNA at the surface of a coated object,54-58 (52) Jewell, C. M.; Zhang, J.; Fredin, N. J.; Lynn, D. M. J. Controlled Release 2005, in press. (53) Luo, D.; Saltzman, W. M. Nat. Biotechnol. 2000, 18, 33-37. (54) Klugherz, B. D.; Jones, P. L.; Cui, X. M.; Chen, W. L.; Meneveau, N. F.; DeFelice, S.; Connolly, J.; Wilensky, R. L.; Levy, R. J. Nat. Biotechnol. 2000, 18, 1181-1184. (55) Klugherz, B. D.; Song, C. X.; Defelice, S.; Cui, X. M.; Lu, Z. B.; Connolly, J.; Hinson, J. T.; Wilensky, R. L.; Levy, R. J. Hum. Gene Ther. 2002, 13, 443-454. (56) Shen, H.; Tan, J.; Saltzman, W. M. Nat. Mater. 2004, 3, 569574. (57) Segura, T.; Volk, M. J.; Shea, L. D. J. Controlled Release 2003, 93, 69-84. (58) Segura, T.; Shea, L. D. Bioconjugate Chem. 2002, 13, 621-629.
therefore, the images in Figure 6 suggest that the structural transformations that occur in these materials under physiologically relevant conditions could also influence and enhance further the internalization and processing of DNA by cells. The complete details of the cellbased transfection experiments described above have been reported in a separate contribution.52 In the context of the current study, however, we speculate that the morphological changes observed for these multilayered DNA-containing materials may prove to be advantageous with respect to the application of these materials to localized gene delivery. Figure 7 shows SEM images of a polymer 1/plasmid film (A) prior to incubation and (B-D) after 4 h of incubation in PBS buffer and provides additional evidence for the formation of nanoparticulate structures on the surface of the substrate. These images represent assemblies of polymer 1/DNA that have been incubated, dried, and subsequently coated with a thin layer of gold prior to imaging. As such, any interpretation of these images related to the behavior of these films in contact with aqueous buffer should be made with caution. However, the images shown in Figure 7B,C suggest that these particles may be fused physically to the surface of the substrate rather than simply being loosely associated with the surface. Figure 7D shows an image of a film that was incubated in PBS for 4 h and then scratched with a needle prior to coating with gold. This image is interesting for at least three reasons: (1) it shows clearly the presence of a residual thin, smooth film on the surface of the silicon substrate; (2) it shows the presence of nanoparticulate structures bound to and apparently fused directly with this residual film; and (3) it shows the presence of small craters or “footprints” in the residual film corresponding to the locations of particles that were presumably removed by force from the surface during scratching.
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Figure 7. SEM images of a multilayered film fabricated from eight bilayers of polymer 1 and pEGFP prior to (A) and after 4 h of incubation in PBS buffer (B-D). Image D shows an SEM image of a scratched film (see text). White scale bars are equal to 1 µm (A, B, D) and 500 nm (C).
The residual film observed in Figure 7D is consistent with the presence of the foundation of LPEI/SPS bilayers deposited on the bare silicon substrate prior to the deposition of the polymer 1/DNA layers (∼20 nm thick, see discussion above). These foundation layers were found to be stable upon incubation over this time period and do not undergo structural rearrangement, as determined above by AFM. We speculate that the interpenetration of adjacent polyelectrolyte layers that occurs necessarily1,3,59,60 at the LPEI/SPS-polymer 1/DNA interface during fabrication leads to the formation of nanoparticulate structures that are physically interpenetrating with or fused to these foundation layers upon rearrangement. The additional craters or indented particle “footprints” observed are consistent with the removal of particles fused to this residual film and further support this hypothesis. We note that the strength of the adhesive interactions between these particles and the surface could influence significantly the rates at which particles are released from the surface (or, conversely, actively removed from the surface by a cell). It may prove to be possible in further experiments, therefore, to exert control over the nature or extent of these interactions simply by engineering the structure or chemical identity of the foundation layers initially deposited on these substrates. Summary and Conclusions Multilayered polyelectrolyte films fabricated from polymer 1 erode gradually under physiological conditions and behave as thin film platforms for the controlled release of incorporated polyanions from film-coated surfaces. In this study, we used AFM and SEM coupled with ellipsometry to characterize the microscale and nanoscale topographies of polymer 1/SPS and polymer 1/plasmid DNA films as a function of erosion. Understanding the physical behavior of these erodible films at these length scales is important, because surface morphology could influence significantly the structural integrity or the range of controlled-release profiles accessible using these materials. Several conclusions can be drawn from this work. First, we find striking differences in the topography and erosion profiles of films fabricated from SPS and plasmid DNA. For films fabricated using SPS as a polyanion, AFM data are consistent with a gradual and well-controlled erosion process that occurs in a top-down, surface-oriented manner without the generation of significant roughness, holes, pits, or cracks over large, micrometer-scale areas of the film. By contrast, films fabricated from plasmid DNA undergo rearrangements on the nanometer scale to present polymer/DNA complexes ranging in size from 50 to 400 nm on the surface of coated silicon substrates. (59) Losche, M.; Schmitt, J.; Decher, G.; Bouwman, W. G.; Kjaer, K. Macromolecules 1998, 31, 8893-8906. (60) Castelnovo, M.; Joanny, J. F. Langmuir 2000, 16, 7524-7532.
Additional characterization of these structures by SEM suggested that they may be interpenetrated with or fused to smooth underlying polyelectrolyte layers on the silicon surface, providing a potential mechanism with which to manipulate adhesive forces and/or the rates at which these particles are released from the surface. In the current study, SPS serves as a model polyanion; films fabricated from transcriptionally active plasmid DNA are of both fundamental and practical interest in the context of gene delivery. The reasons for the dramatic differences in the erosion profiles for assemblies fabricated from these two different polyanions are not completely understood. However, these profiles represent two extremes of behavior (e.g., surface erosion and bulk structural rearrangement) that may influence significantly the range of controlled-release applications for which these films are suited. For example, the layer-by-layer fabrication procedure can be used, in principle, to fabricate multilayered films having stratified structures and spatially defined regions of chemically or functionally different polyelectrolytes. The well-defined nature of the erosion profile observed for polymer 1/SPS films suggests, therefore, that it may be possible to design assemblies that release two film components located in different regions of a film (e.g., the top and bottom regions) with well-defined release kinetics. The structural reorganization observed for DNA-containing films, however, suggests that control over the sequence with which two different plasmids are released from these materials may prove to be difficult using this polycation/DNA system. In the specific context of gene delivery and gene therapy, the presentation of condensed nanoparticles of DNA at these surfaces may be advantageous with respect to stimulating the internalization and intracellular processing of DNA by cells. A quantitative understanding of the factors influencing the fabrication, structure, and erosion profiles of these polyelectrolyte assemblies will be useful for the design of erodible assemblies for specific applications in which controlled film erosion and the release of therapeutic materials are desired. Acknowledgment. Financial support was provided by the National Institutes of Health (EB02746), the Arnold and Mabel Beckman Foundation, the National Science Foundation (through the University of Wisconsin Materials Research Science and Engineering Center), and the University of Wisconsin. N.J.F. gratefully acknowledges the NSF for a graduate research fellowship. We thank Christopher M. Jewell and Nicholas L. Abbott for numerous helpful discussions. LA050596+
