pubs.acs.org/Langmuir © 2009 American Chemical Society
Interaction of Lipid Membrane with Nanostructured Surfaces Yuri Roiter,† Maryna Ornatska,† Aravind R. Rammohan,‡ Jitendra Balakrishnan,‡ David R. Heine,‡ and Sergiy Minko*,† †
Department of Chemistry and Biomolecular Science, NanoBio Laboratory (NABLAB), Clarkson University, Potsdam, New York 13699-5810, and ‡Corning Incorporated, Corning, New York 14831 Received January 11, 2009. Revised Manuscript Received February 27, 2009
Tiny details of the phospholipid (DMPC) membrane morphology in close vicinity to nanostructured silica surfaces have been discovered in the atomic force microscopy experiments. The structural features of the silica surface were varied in the experiments by the deposition of silica nanoparticles of different diameter on plane and smooth silica substrates. It was found that, due to the barrier function of the lipid membrane, only particles larger than 22 nm in diameter with a smooth surface were completely enveloped by the lipid membrane. However, nanoparticles with bumpy surfaces (curvature diameter of bumps as that of particles 5, the lipid membrane envelops particles larger than 22 nm. The interaction of the lipid membrane with particles larger than 22 nm and with bumps on the surface where the size of the bumps fits into the range 1/5 < κ < 5 will be considered separately. All measurements were performed at 28 C, above gel-liquid crystal phase transition temperature of studied DMPC (24 C). Lipid Membrane on Smooth Substrate, K , 1. This reference experiment demonstrates the formation of a lipid bilayer about 5 nm thick on a flat and smooth silica surface (Figure 1a and 1b.1). AFM allows observation of many details of the SLB formation in situ. For instance, observation may reveal lipid vesicles adsorbed onto the surface, which are flattened to a thickness of about two bilayers (Figure 1a [blue arrows] and 1b.2). Sometimes such vesicles flattened directly under the AFM tip, and then, the immediate formation of lipid bilayer patches is observed. Such a case is shown in Figure 1a with green arrows and the profile in Figure 1b.3. The image reveals the lipid vesicle as it flattens over the substrate. Both the flattened fraction and the residual parts of the vesicle can be clearly seen on the image and its cross section. With time, lipid bilayer patches grow in size and merge until, eventually, the surface is completely covered with uniform lipid bilayer. Lipid Membrane on Particles Smaller than 1.2 nm, K < 1/5. Analysis of the substrate with particles under ∼1.2 nm in diameter found that the lipid membrane follows the topographical features of the underlying substrate (Figure 2a-c). In other words, the topography of the membrane repeats the topography Langmuir 2009, 25(11), 6287–6299
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Figure 1. AFM image recorded in situ for the DMPC bilayer on the smooth silica surface: (a) blue arrows denote vesicles adsorbed on the silica substrate; green arrows show the vesicles unfolding under the AFM tip (the location of the cross sections marked with straight lines); and cross-sectional profiles represent various structural features formed by the deposited lipid on the surface. (b) 5.3 nm thick lipid bilayer (1); flattened 11.5 nm lipid vesicle (2); lipid vesicle under the AFM tip fusing onto the substrate and forming the lipid bilayer (scanning was performed upward) (3).
of the silica substrate. A further increase of the particle diameter results in the formation of pores in the membrane as discussed below. The transition point ∼1.2 nm is an estimated value obtained from the analysis of about 100 particles with an error of at least 0.3 nm that corresponds to the inherent rms roughness of the SiO2 wafer. For example, the particle marked as 1 in Figure 2d is 1.2 nm in diameter as shown in the corresponding cross section (i) (Figure 2h.1, the cross section locus is marked with the line 1 in the second copy of the topography image, Figure 2d [middle column]). The left and middle columns of the images in Figure 2 are the same sequence of AFM images duplicated for the convenience of their analysis. The AFM images are aligned with respect to the initial image (Figure 2d). Deposition of the lipid bilayer results in a change in the profile Figure 2h.1 (ii). The surface became smoother after deposition; however, the cross section indicates the nanoparticles located underneath the lipid bilayer. The subsequent exposure of the sample to an insulin solution for different period of times did not bring about any changes in the profiles (Figure 2h.1 (iii) and Figure 2h.1 (iv)). Insulin is a 3.5 nm diameter globular protein, which adsorbs on silica surfaces and does not adsorb on the lipid bilayer. No changes were detected in the profiles of the samples exposed to the insulin solution; thus, these particles caused no rupture or permeable (for insulin) pores in the lipid membrane. These observations show that the membrane closely follows the topography of the silica surface and seals the substrate. Lipid Membrane on 1.2-5 nm Particles, 1/5 < K < 1. Three examples of single nanoparticles 3.4 nm (particle 2), 2 nm (particle 3), and 5 nm (particle 4) in diameter are shown in Figure 2e,f,g, respectively, with corresponding cross-sectional profiles in Figure 2h (2-4). The addition of DMPC results in a dramatic change in profiles (Figure 2h (ii) and (iii)). The AFM images reveal that the lipid bilayer “covers” the nanoparticles with no elevations but, rather, with a small decrease of the bilayer thickness over the particle. This result suggests that the bilayer does not follow the structures formed by the deposited particles on surface. The presence of hydrophilic particles in the continuous lipid bilayer is thermodynamically unfavorable, since the DOI: 10.1021/la900119a
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Figure 2. Formation of the lipid bilayer and the dynamics of insulin adsorption via AFM images representing the effects of 5. A series of adapted 3D AFM images (Figure 8a-d) demonstrate a typical picture of the lipid membrane formation on a silica surface decorated with 5-140 nm nanoparticles. Even in the early stages of the SLB formation, bilayer fragments can be seen on the large particles (Figure 8b). In the final stage, the bilayer almost completely covers the large particles, whereas the holes remain only around the smaller nanoparticles (Figure 8c). This difference in coverage of small and large particles can be clearly seen in the 3D image of the lipid bilayer (Figure 8d). Figure 8f-k show the original AFM images of the lipid bilayer on the silica substrate decorated by a mixture of several fractions of the particles ranging in size from 5 to 140 nm. The AFM images shown in Figure 8f-h are aligned. Figure 8g, h shows the topography of the sample partially covered by lipid bilayer and the lipid coating in the final stage, respectively. The image subtraction assists in the visualization of the deposited lipid bilayer and in the building of adapted 3D images (Figure 8a-d) based on the original and subtracted AFM data matrices. Figure 8i shows “free standing” lipid bilayer islands (marked with blue arrows). Figure 8j demonstrates pores in the lipid bilayer around smaller nanoparticles and on larger particles elevated above the layer (denoted by green arrows). Free Energy Model. To more closely examine the particlecoating behavior of the lipid membrane, we constructed a free energy model of the membrane that describes the extent of coverage as a function of the particle size. The model is shown schematically in Figure 9. The membrane, depicted as the thick solid line, coats a silica particle of radius r on a silica substrate. The membrane itself has a radius of rmem and a constant surface area of Amem. The height at which the membrane separates from the particle is denoted by h. This height also determines the arc length of membrane-particle contact denoted by θ and the membrane-particle contact area, Acap. 6294
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Figure 7. Lipid bilayer formation in the presence of 5-20 nm silica particles. Adapted 3D AFM images: (a) substrate with particles and no lipid; (b) surface partially covered by lipid bilayer (shown in silver color); (c) lipid bilayer formed on the substrate; (d) image of the lipid bilayer after “subtraction” of the particles and the substrate. Schematics: (e) illustrates how the lipid bilayer forms a pore around 5-22 nm particles. Original AFM images: (f) sample at the initial stage without a lipid bilayer; (g) partial coverage of the surface with lipid; (h) the sample in the final stage covered by a lipid bilayer; (i) islands of lipid bilayer obtained by subtraction of image (f) from image (g); (j) lipid bilayer with holes around nanoparticles obtained by subtraction of image (f) from image (h); (k) lipid bilayer obtained by subtraction of image (g) from image (h).
Below h, we assume that the membrane adopts a radius of curvature r2 down to the point where it contacts the substrate at a distance r1 from the center of the silica particle. The area of the membrane between the particle and substrate, Awater, is defined by the torus with radius r1 and tube radius r2. The remaining membrane area in contact with the substrate is Asub. The expression for the free energy of the membrane is obtained by borrowing from the vesicle spreading model of Efremov et al.48 and adding terms to account for the silica particle. The free energy consists of two components: the interfacial energy due to the surface tension between the membrane, water, and silica surfaces and the bending energy due to coating the silica particle. The interfacial free energy considering a total substrate area of Amem is given by the interfacial area and surface tension as Gint ¼ Amem γlw þ Acap γls þ Awater γlw þ Asub γls þ ðAmem þ Aparticle -Asub -Acap Þγsw ð1Þ where γlw, γls, and γsw are the lipid-water, lipid-silica, and silica-water surface tensions, respectively. The bending free energy is expressed as Gbend ¼ Acap
1 1 1 2 1 1 1 2 þ Awater Kc þ Kc þ 2 r r 2 r1 r2
ð2Þ
where Kc is the membrane bending modulus. To eliminate the dependence on the total membrane size, we solve for the free energy difference between the particle coating membrane and the Langmuir 2009, 25(11), 6287–6299
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Figure 8. Lipid bilayer formation in the presence of 5-140 nm silica particles. Adapted 3D AFM images: (a) substrate with particles and no lipid; (b) surface partially covered by lipid bilayer (shown in silver color); (c) lipid bilayer formed on the substrate; (d) image of the lipid bilayer after “subtraction” of the particles and the substrate. Schematics: (e) illustration of how the lipid bilayer envelopes particles larger than 22 nm particles. The structure of the bilayer area encircled in (e) is speculative, since it cannot be resolved or assumed from AFM experiments. Original AFM images: (f) the sample at the initial stage without the lipid bilayer; (g) partial coverage of the surface with lipid; (h) the sample in the final stage covered by the lipid bilayer; (i) islands of lipid bilayer obtained by subtraction of image (f) from image (g); (j) lipid bilayer with holes around nanoparticles obtained by subtraction of image (f) from image (h); (k) lipid bilayer obtained by subtraction of image (g) from image (h). Blue arrows denote examples of areas covered by the lipid at the partial coverage. Green arrows denote the examples of areas (dark spots) on large particles that were not covered by lipid.
Figure 9. Schematic of the membrane free energy model showing the lipid membrane as the thick solid line partially coating the spherical silica particle of radius r and the flat silica substrate.
membrane coating the flat substrate with the particle exposed where the free energy is Gflat ¼ Amem ðγlw þ γls Þ þ Aparticle γsw
ð3Þ
Combining eqs 1-3 gives the expression for the change in free energy due to the presence of the particle " # 1 1 1 2 ΔG ¼ Awater γlw þ γsw -γls þ Kc þ þ 2 r1 r2 " # 1 1 1 2 þ ð4Þ Acap Kc 2 r r The unknown variables in eq 4 are expressed in terms of the dimensionless variable h* = h/r yielding r1 = 2r/h*(2h* - h*2)1/2 Langmuir 2009, 25(11), 6287–6299
and r2 = 2r/h* - r. To obtain Awater, we apply Pappus’s centroid theorem for arc length s = θr2 and geometric centroid at x = r1 - r2 (1 - cos θ)/θ, which results in Awater = 2π(θr1r2 r22(1 - cos θ)). The area of the particle coated by the membrane is Acap = 2πrh. The free energy defined in eq 4 is solved for multiple values of the particle radius ranging from 1 to 50 nm and h* ranging from 0 to 1. The surface tension and bending modulus values are γlw = 0.02 J/m2, γls = 0.01 J/m2, γsw = 0.006 J/m2, and Kc = 1.2 10-19 J.48 With these values, the dependence of ΔG on h* and r is as shown in Figure 10. For each particle size, the free energy shows a minimum that balances the decrease in interfacial energy due to the membrane-substrate interaction with the increase in bending energy due to the diminishing radius of curvature as h approaches 2r. The energy minimum is quite broad for small particle sizes due to the small amount of surface area involved, whereas the energy becomes increasingly sensitive to h* for larger particle sizes. Extracting the energy minimum and corresponding height h* from Figure 10 results in the energy dependence on particle size shown in Figure 11. Here, we see that the membrane will coat most of the particle if the particle radius is greater than about 15 nm, although at least half of the particle will be coated for even the smallest nanoparticles. The change in free energy of the coating membrane is compared to the energy required for the equivalently sized membrane to form a pore. This energy is calculated as the bending energy required to form endcaps around a circle of radius r as detailed in Efremov et al.48 The final expression is ΔGpore ¼ 2πKc
ðt þ rÞ2 ðπr þ tÞ ðr þ t=2Þ2 t
ð5Þ
where t = 3 nm (applied in Efremov’s model) for the thickness of hydrophobic part of the membrane. Thus, the membrane coating model predicts a transition between particle coating and pore forming with pore formation being energetically favorable for particle radii below 13 nm (or diameter below 26 nm) in accord with the experiment. Lipid Bilayer on Bumpy Particles. It was found that some samples of larger particles (κ > 5) were only partially covered by the lipid bilayer (Figure 8c,d,j). Using transmission electron microscopy (TEM) (Figure 5j), it was discovered that some samples of nanoparticles have a bumpy structure (Figure 5h,j). The curvature diameter of the bumps on the surface was around 10 nm. It is likely that nanoparticles or nanostructures that affect the formation of pores in a lipid membrane will show the same behavior on planar and curved surfaces. Thus, it may be speculated that, if the bump curvature is in the range 1/5 < κ < 5, they are able to pierce the lipid bilayer. This hypothesis was supported in the experiment with fluorescent dye-labeled insulin. Two different samples of nanoparticles were used. The sample of 100 nm diameter bumpy nanoparticles, from Polysciences (Figure 5h,j), and the sample of 200 nm nanoparticles with smooth surfaces (Figure 5i,k), from Microparticles GmbH, were deposited on SiO2 wafers and coated by the lipid bilayer. Afterward, the samples were exposed to the solution of labeled insulin. Optical images of the samples obtained with confocal microscope are shown in Figure 5d,e. The sample prepared from bumpy particles (Figure 5d) is colored more intensely compared to the sample with smooth particles (Figure 5e). Thus, the labeled insulin adsorbed on the bumpy nanoparticles through the holes created in the lipid bilayer around the surface features on the bumpy particles. DOI: 10.1021/la900119a
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Figure 10. Free energy of the membrane system as a function of the amount of silica particle coverage for particle radii ranging from 1 to 50 nm. γlw = 0.02 J/m2, γls = 0.01 J/m2, γsw = 0.006 J/m2, and Kc = 1.2 10-19 J.
Figure 11. Free energy of the membrane system as a function of the particle radius. The model predicts that a pore-forming structure is more energetically favorable than a particle-coating structure for particles below 13 nm.
Discussion Fusion of lipid vesicles to the surface of a polar solid substrate evolves over several steps: approaching the surface, interaction with the substrate, and spreading of the bilayer over the solid surface. Here, the final stage of the formation of the lipid bilayer is discussed. The reported experiments revealed that the spreading lipid membrane over the nanostructured surface results in the formation of a complex morphology of the lipid membrane where the local structure of the membrane is affected by the dimensions and shapes of the nanostructures. This paper addresses several important concerns regarding the process of the SLB formation. Since spreading of the lipid bilayer over the surface by fusion of vesicles is a rapid process, it may be argued that many metastable structures could appear in the SLB. Although analysis of the complex mechanism of lipid vesicle fusion is beyond the scope of this paper, it should be mentioned that the current author’s study of the fusion kinetics revealed that the fusion passes through multiple cycles of the attachment of vesicles to the substrate, spreading of the bilayer, and then the shrinking of the bilayer and detachment of lipid from the surface. The spreading of the vesicle is a rapid process; however, due to the multiple deposition and detachment steps, the final “equilibrium” structure was usually approached in half an hour or more. As soon as the final structure was approached, no further changes were observed in the SLB structure over time and after further addition of lipid vesicles to the solution. Thus, it is very unlikely that metastable structures dominate in the SLB morphology in 6296
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these experiments. The second important question is whether the AFM tip may create artifacts in the images. Multiple repetitions of the experiments under the same conditions and with changes to the number and frequency of scans in time helped to identify any artifacts caused by the probe. No evidence was found (besides some rare examples) for the damaging effects of the AFM probes in these experiments. Only reproducible structures were included in the experiments. The analysis of the SLB structures on the nanostructured surfaces clearly demonstrates the barrier properties of the lipid membrane. The barrier function is attributed to two major components: surface tension and bending (curvature) energy, both, to a large extent, due to the hydrophobic interactions. These contributions to the barrier function of the lipid membrane is nicely analyzed in the model of Deserno and Gelbart61,63 for interactions of nanoparticles and lipid vesicles. The major differences between the SLB and lipid vesicles is that (1) the SLB is considered as an infinite 2D structure, while the lipid vesicle has given dimensions; and (2) in the case of the interaction of the vesicle with a nanoparticle, the surface tension and bending energy is balanced by adhesion energy between the nanoparticle and the lipid membrane. For the SLB, in addition to the adhesion between the lipid bilayer and the nanoparticle, adhesion between the lipid bilayer and the solid substrate supporting the nanoparticle must also be taken into account. Due to this difference, an upper limit of the nanoparticle diameter for enveloping nanoparticles with the lipid membrane is expected. To address the specific SLB structure on surfaces decorated with nanoparticles, we applied the develop free energy model of the membrane that describes the coverage as a function of the particle size and obtained the upper limit to be 26 nm (Figure 11). From the statistical analysis of about 700 particles from at least 10 images of lipid membrane around nanoparticles ranging in diameter from 5 to 140 nm with an emphasis on 5-65 nm range (see Supporting Information, Figure S2) the experimental value of d0 was found to be 22 nm. This value was evaluated as corresponding to 50% coverage by the lipid bilayer, and it was obtained from the plot surface coverage versus the particle diameter (Figure 12). Nanoparticles with a diameter greater than 22 nm are mostly enveloped by lipid bilayer (Figure 8e), while smaller nanoparticles remain mostly uncovered (Figure 7e). The upper limit for the particle diameter 22 nm is smaller than that obtained in the free energy model 26 nm but still reasonable given the uncertainty in the surface tension parameters and the simplicity of the model. Further, the model shows that, for particle sizes large enough to avoid pore formation, most of the particle is coated by the membrane as opposed to having just the upper portion coated and the rest of the particle surface exposed to water. Hence, this model provides some additional insight on membrane conformation around nanoparticles that is inaccessible from AFM measurements alone. Further decreases in the particle diameter result in the decrease of pore size, and finally, the nanobumps become capped by the lipid membrane. Thus, lipid membrane follows the substrate surface, leaving small grooves and cavities (with curvature diameter below 1.2 nm) filled with water. The experimental results are in accord with dynamic simulations of a continuum SLB on silica substrates in an implicit solvent. The computational results predict that the bilayer conforms to the substrate topography for feature sizes 22 nm are mostly coated by the lipid bilayer (Figure 13d) unless the surface of the particle is rough, in which case coating around bumpy spherical nanoparticles is incomplete (Figure 13e). The results obtained in this study are useful for the selection of dimensions and shapes for drug-delivery cargo and for the substrate for supported lipid bilayers. They also help in understanding the role of length scales involved in the mechanisms of endocytosis and cytotoxicity of nanoparticles. Our finding could also be used for the development of novel strategy for nanopatterning of substrates with biomolecules and other species precisely placed into the pores formed in the lipid bilayer. The reported system can be seen as an example of the self-assembly approach to the fabrication nanostructured materials.64,65 The proposed here approach extends a range of recently reported self-assembly based fabrication methods for porous materials with a gel matrix with well-controlled pore size and pore size distribution.66-68
Experimental Section
Figure 13. Schematics of lipid bilayer membrane fusion on rough surfaces: lipid bilayer forms a pore around particles (a) larger than 1.2 nm and (b) smaller than 22 nm, whether the nanoparticles are smaller or larger than the thickness of the bilayer; the lipid membrane (c) follows the surface topography with nanoparticles below 1.2 nm and (d) envelops large nanoparticles of more than 22 nm; (e) incomplete coverage of bigger particles due to the presence of surface bumps. The structures of bilayer areas such as the one circled in (d) are speculative, since they cannot be resolved or assumed from AFM experiments.
The discrepancy may be due to the difference in formulation of the model membrane when compared to the adsorbed bilayers described above. The model suggests that the inability to conform to larger surface curvatures is due to the unfavorable bending energy of the membrane, compared to the attractive force pulling the membrane to the surface. In the case of the model, the results show considerable deviations of the membrane from the substrate. In the experimental study described here, pure DMPC bilayers lack natural mechanisms to sustain their lateral integrity (e.g., transmembrane proteins, sugars, collagen, etc.). The adverse bending energy needed to coat large features would thus result in a local pore formation. Langmuir 2009, 25(11), 6287–6299
Materials. Silica nanoparticle suspensions were acquired from several suppliers: (1) Alfa Aesar (USA, MA), samples with average diameters of (da) 4 nm, 10 nm, 14 nm, and 20 nm; (2) Sigma-Aldrich (MO, USA), da = 15 nm; (3) Nyacol Nano Technologies (MA, USA), da = 20 and 35 nm; (4) Nissan Chemical America (TX, USA), da = 25 and 45 nm; (5) Polysciences (PA, USA): da = 50 and 100 nm; (6) Microparticles GmbH (Germany), da = 200 nm. Lipids, L-R-dimyristoyl phosphatidylcholine (DMPC, phase transition temperature 24 C) were purchased from Avanti Polar Lipids (AL, USA). Silicon wafers were purchased from Silicon Quest International (CA, USA). Millipore water (18.3 MΩ 3 cm) was used in all preparations. All other chemicals were acquired from Sigma-Aldrich (MO, USA) and used as received. Preparation of Unilamellar Phospholipid Vesicles. DMPC (25 mg) was dissolved in 1 mL of chloroform. The chloroform was evaporated under the steam of dry nitrogen and further dried overnight in a vacuum. Vesicles were obtained by sonication of the DMPC suspensions for 2 h (Branson 2510, Branson Ultrasonics, CT, USA) at 40 C in 10 mL of Tris-HCl buffer (10 mM Tris-HCl, 150 mM NaCl, 2 mM CaCl2 3 2H2O, pH 7.4 adjusted by NaOH). Vesicle suspensions were stored in a refrigerator. The lipid suspension was diluted to a concentration of 0.5 mg/mL in Tris-HCl buffer before use, as follows: 2.5 mg/mL lipid suspension was heated to 40 C and extruded through 0.2 μm Teflon filter, also heated to 40 C. According to dynamic light scattering analysis (Particle Size Analyzer 90 Plus, Brookhaven Instruments, NY, USA), the typical vesicle diameter was 184 nm with a polydispersity index of 1.2. (64) Nie, Z. H; Kumacheva, E Nat. Mater. 2008, 7, 277–290. (65) Palmer, L. C.; Stupp, S. I. Acc. Chem. Res. 2008, 41, 1674–1684. (66) (a) Sidorenko, A.; Tokarev, I.; Minko, S.; Stamm, M. J. Am. Chem. Soc. 2003, 125(40), 12211–12216. (b) Tokarev, I.; Krenek, R.; Burkov, Y.; Schmeisser, D.; Sidorenko, A.; Minko, S.; Stamm, M. Macromolecules 2005, 38, 507–516. (67) (a) Tokarev, I.; Orlov, M.; Minko, S. Adv. Mater. 2006, 18, 2458–2460. (b) Orlov, M.; Tokarev, I.; Scholl, A.; Doran, A.; Minko, S. Macromolecules 2007, 40, 2086–2091. (68) (a) Gopishetty, V.; Roiter, Y.; Tokarev, I.; Minko, S. Adv. Mater. 2008, 20, 4588–4593. (b) Tokarev, I.; Minko, S. Adv. Mater. 2009, 21, 241–247. (c) Tokarev, I.; Gopishetty, V.; Zhou, J.; Pita, M; Motornov, M. Katz, E.; Minko, S. ACS Appl. Mater. Interfaces 2009, 3, 532–536.
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Preparation of Silicon Wafers. Cut silicon chips (11 11 mm2) were successively sonicated in absolute ethanol for 15 min, sonicated in methylene chloride for 15 min twice, cleaned in a 1:1:1 solution of water, 30% ammonia, and 33% hydrogen peroxide for 1 h at 60 C, thoroughly rinsed with water, kept underwater for 24 h, and dried by nitrogen-blowing before use. Nanoparticle Deposition. Cast deposition was used to apply particles 40 nm and larger to the surface. Water dispersions of different concentrations for different sizes of particles were used: 0.25 g/L for 40-50 nm particles, 0.7 g/L for 100 nm particles, and 1.8 g/L for 200 nm particles. Dispersions were sonicated for 1 h before the deposition. A 20 μL drop of nanoparticle suspension was evenly spread onto a dried silicon chip and allowed to slowly dry at room temperature. Spin-assisted deposition was used to apply particles smaller than 40 nm. Solution concentrations were: 0.10 g/L for 4 nm particles, 0.25 g/L for 10-30 nm particles, and 0.75 g/L for 35 nm particles. A 100 μL drop of nanoparticle suspension was deposited on a dry silicon chip, allowed to stand for 1 min, and then spin-coated for 9 min at 1000 rpm. The combined procedure was applied for the mixtures of smaller and larger particles. First, smaller particles were deposited by spincoating. Next, larger particles were applied by cast deposition. All Samples Were Treated with Tetraethyl Orthosilicate (TEOS) Prior to Use. The procedure did not noticeably change the character of lipid bilayer formation on the surface in comparison with the untreated samples. However, the procedure did ensure similar surface composition over the range of samples and provided the fixation of the particles onto the surface. To perform the treatment, a paper filter was placed inside the Petri dish cover and soaked with 20 μL of TEOS. Prepared substrates were placed in the Petri dish, covered with a cap, and incubated for 2 h at 50 C. After incubation, samples were treated with plasma for 2 min (Expanded Plasma Cleaner, 10.15 W, Harrick Plasma, NY, USA), and kept for 1 h under Tris-HCl buffer. The typical thickness of silica layers formed by hydrolyzed TEOS and measured ellipsometrically (Multiskop Ellipsometer, Optrel, Golm, Germany) on plain silicon wafers was 0.4 ( 0.05 nm. AFM Experiments. AFM images were recorded using a MultiMode scanning probe microscope (Veeco Instruments, NY, USA) equipped with a custom-built temperature stage, and operated, in tapping mode, in liquid. Samples were scanned using NPS silicon nitride probes (Veeco Instruments, NY, USA) with a spring constant of 0.32 N/m, and a resonance frequency in aqueous media of ∼9 kHz. The radius of the tip curvature was determined internally for each experiment series, accounting for the overall spherical shape of the recorded particles, and was found to be primarily in the range between 25 and 50 nm. Scanning was performed with an amplitude set point in the range 0.9-2.4 V, tapping force of 95-98% from the set point, integral gain 0.2-0.6, proportional gain 2-6, speed of scanning 0.4-2 Hz, and temperature 28.0 ( 0.5 C (the temperature in the fluid cell of the MultiMode microscope, was measured by a calibrated thermistor placed into the fluid cell). The sample was placed on the holder of the MultiMode, and carefully covered by the fluid cell (MTFML, Veeco Instruments, NY, USA) to minimize the mechanical drift effects caused by the silicone rubber O-ring. 50 μL of Tris-HCl buffer was introduced into the fluid cell. A microscope was then equilibrated for 0.5-2 h to minimize thermal drift of the scanner. The actual time of equilibration was estimated by the stabilization of the initial sample image. After that, the set point was increased to 5 V, 5-20 μL. The Tris-HCl buffer was carefully removed from the fluid cell and replaced with 5-20 μL of 0.5 g/L lipid suspension, and in the planned time, the set point was usually returned to the original value. A typical lateral sample displacement after this procedure was 100-1000 nm and was easily compensated by the lateral offset command. Images were taken continuously (providing the details of lipid bilayer formation) or periodically (providing a picture of lipid bilayer formed without
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being disturbed by the AFM tip). The best results were obtained if lipid suspensions were added in two steps. In the first step, 10 μL of the lipid suspension was added, and the image of a partial lipid bilayer was recorded after 20 min. In the second step, 20 μL of the suspension was added and the image of the lipid bilayer was recorded after 20 min. After each addition of the lipid, the tip was retracted to the 10 V set point. The tip was slowly oscillated with 20 μm amplitude. In this regime, the tip did not hinder the deposition of the lipid onto the studied surface. Transmission Electron Microscopy. Nanoparticles were examined by transmission electron microscopy (TEM, JEM 2010, Jeol, Japan). For TEM analysis, specimens were prepared by placing a 5 μL drop of the suspension used for the wafer preparation onto a copper grid with a transparent carbon film, followed by drying. Confocal Microscopy. Nikon C1 confocal with Inverted Eclipse TE 2000U light microscope (Nikon, NY, USA) was supplied with a 20 objective lens and a 488 nm laser. FITCinsulin (488-525 nm) was used as a fluorescent marker. Four 1 μL drops of the particle dispersions (1.8 g/L dispersion of 200 nm particles; 0.7 g/L of 60-140 nm particles; 0.02 g/L of 5-20 nm particles; and 0.01 g/L of 1-8 nm particles), which provided nearly the same surface coverage, were cast in different locations of the same silicon wafer and slowly dried. The lipid suspension (0.5 g/L, 20-100 μL) was deposited on the silicon wafer decorated with nanoparticles, sealed in a vapor-tight Petri dish containing filter paper wetted with Tris-HCl buffer, and incubated for 30 min at 30 C temperature. Then, the Petri dish with the sample was placed into a refrigerator (5 C) for 10 min to provide a stable gelphase DMPC bilayer over the surface. 50 μL of a 5 10-2 g/L solution of FITC-insulin in Tris-HCl buffer, cooled to 5 C, was then added to the drop of the lipid suspension and kept at 5 C for another 30 min. Afterward, the sample was carefully washed with cooled to 5 C Tris-HCl buffer and placed on a microscopic cover glass, face-down without drying, and viewed with 200 magnification. Placement of all the samples of particles on the same wafer ensured identical conditions during their examination. Data Processing. AFM images were processed using WSxM software.69 3D Images were prepared from real AFM topography images using Autodesk 3ds Max 8 (Autodesk, CA, USA). Subtraction of AFM Images. The present authors have written the software for subtraction of AFM images in Borland C++ Builder 6.0 environment (Borland, TX, USA). Subtracted AFM images were obtained as a difference between two AFM data matrices after alignment of the corresponding minuend and subtrahend images. The alignment procedure was performed, accounting for thermal or mechanical drift, along slow and fast scan axes. AFM data matrices were loaded from the WSxM software format and presented on a screen as raster images (for operator convenience). Positioning, scaling, and skewing of the minuend image enabled its precise alignment with respect to the subtrahend image with resolution of a single data point. Algorithms commonly applied for the processing of raster images were used. Upscaling was calculated using a bilinear averaging algorithm. Skewing was performed by linear averaging, and downscaling was performed by fractional averaging. These algorithms provide average height values based on either averaging of the surrounding height data with respect to their lateral position relative to the new data point (upscaling and skewing), or averaging of all the data points falling into the new data point, weighed by the contribution of their fractions (downscaling). Every step of scaling or skewing is recalculated starting from the original AFM data matrix to avoid error accumulation. Interactive control of image alignment was provided by manual superposition of the semitransparent image-minuend (typically, samples with SLB) over the opaque image-subtrahend (typically, samples in the initial stage with no lipid) and simultaneous observation of the resulting image-difference. (69) Horcas, I.; Fernandez, R.; Gomez-Rodriguez, J. M.; Colchero, J.; GomezHerrero, J.; Baro, A. M. Rev. Sci. Instrum. 2007, 78, 013705.
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Roiter et al. Alignment in the XY-plane was followed by alignment in the Zdirection. Regions of equal offset (for uncoated substrate and the substrate coated by lipid bilayer) were marked in the superposed AFM images. Then, the subtraction was performed on a pointby-point basis. For each marked region (j) an average of the regions offset values (ÆΔZxyjæ) was evaluated. The Z coordinate was corrected by subtraction of Δzxy values evaluated as Δzxy = ÆΔZxyjæ for uncovered regions and Δzxy = ÆΔZxyjæ - 5 nm for regions of the substrate coated by lipid bilayer. Evaluation of Particle Coverage with Lipid Bilayer. The diameter of particles deposited on the silica wafer was evaluated from the height profiles of topographical AFM images of the samples in the initial stage (no lipid bilayer). The integral distribution function of surface coverage of the particles with lipid bilayer (see Figure 12) was plotted for several groups of particles, with 24 particles in each group. The particles were assigned to different groups by diameter, with 2 and 10 nm increments for 6-40 nm and 40-140 nm ranges of diameters, respectively. The coverage of the particles by the lipid bilayer was evaluated based on the quality of the lipid coating on top of the particles, since imaging of the lipid layer on the particle sides may be corrupted. If the top of the particle was found to have either no coverage or complete coverage by the lipid membrane, 0% or 100% coverage was assigned to that particle. If the coverage was partial, the masks were built over the top area to evaluate the surface area uncovered (AP, image pixels) and areas covered by the lipid bilayer (AL, image pixels). Simple relation of the pixel quantities 100% 3 AL/AP provided the estimation of partial
Langmuir 2009, 25(11), 6287–6299
Article coverage for each corresponding particle. Large standard deviation bars in the transition zone (see Figure 12) were caused not by the accuracy of the measurements, but by the distribution of membrane coverage over particles of a certain diameter. For example, in the range of particle sizes 20-40 nm, one particle could be covered completely and another of the same diameter could be not covered at all, thus indicating the transitional region of surface curvature for the lipid membrane deposition. Prolonging the time of bilayer preparation and increasing the lipid concentration after the formation of the final membrane did not change this situation. The Richards function was applied to fit the experimental data (SRichards2 in Origin 7.5 SR4, OriginLab, USA, MA). This generalized logistic function provided a good fit to the experimental data (coefficient of determination is 0.993).
Acknowledgment. This work was supported by Corning, Inc., and NY Center for Advanced Materials Processing at Clarkson University. We thank Dr. Sokolov and Mr. Bill Plunkett (Clarkson University) for their help with the fluorescent confocal microscopy and TEM analysis. We thank Dr. Akhremitchev (Duke University) for his help with the design of a custom-built temperature stage for AFM studies. Supporting Information Available: Theoretical curves of nanoparticle lateral dilation, penetration of a tip into the pore of a lipid bilayer, and AFM data of the lipid bilayer formation in samples with 5-65 nm particles. This material is available free of charge via the Internet at http://pubs.acs.org.
DOI: 10.1021/la900119a
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