Materials and Methods

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1 Supporting Information for Interaction of nanoparticles with lipid membrane by Y. Roiter, M. Ornatska, A. R. Rammohan, J. Balakrishnan, D. R. Heine, and S. Minko*, Department of Chemistry and Biomolecular Science, Clarkson University, Potsdam, New York , Corning Inc., Corning, New York 14831, * To whom correspondence should be addressed. E- mail: sminko@clarkson.edu. Materials and Methods Materials Silica nanoparticles suspensions were acquired from several providers. (1) Alfa Aesar (USA, MA), average diameter (d a ) 4 nm, 10 nm, 14 nm, and 20 nm; (2) Sigma-Aldrich (MO, USA), d a =15 nm; (3) Nyacol Nano Technologies (MA, USA), d a = 20 nm and 35 nm; (4) Nissan Chemical America (TX, USA), d a =25 nm and 45 nm; (5) Polysciences (PA, SA): d a = 50 nm and 100 nm; (6) Microparticles GmbH (Germany), d a = 200 nm. Particles from different manufacturers were tested in order to verify the reproducibility of the results. Lipids, L-α-dimyristoyl phosphatidylcholine (DMPC, phase transition temperature 24 o C), and polar lipid extract from bovine liver were purchased from Avanti Polar Lipids (AL, USA) with the composition: cholesterol - 5 %wt.; phosphatidylethanolamine 26 %wt.; phosphatidylinositol - 9 % wt.; phosphatidylcholine - 42 %wt.; lyso phosphatidylinositol - 1 %wt.; others, including neutral lipids - 17%wt. Silicon wafers were purchased from Silicon Quest International (CA, USA). Millipore water (18.3 MΩ cm) was used in all preparations. All other chemicals were acquired from Sigma-Aldrich (MO, USA). Preparation of unilamellar phospholipid vesicles DMPC (25 mg) was dissolved in 1 ml of chloroform. Chloroform was evaporated under the stream of dry nitrogen, and further dried overnight under vacuum. Vesicles were obtained by sonicating the DMPC suspensions for 2 hours (Branson 2510, Branson Ultrasonics, CT, USA) at 40 o C in 10 ml of Tris-HCl buffer (10mM Tris-HCl, 150mM NaCl, 2mM CaCl 2 2H 2 O, ph 7.4 adjusted by NaOH). Vesicle suspensions were stored in refrigerator. Working lipid suspension was diluted to concentration 0.5 mg/ml in Tris-HCl buffer before use as follows: 2.5 mg/ml lipid suspension was heated to 40 o C and extruded through 0.2 µm Teflon filter heated as well to 40 o C. According to Dynamic Light Scattering analysis (Particle Size Analyzer 90 Plus, Brookhaven Instruments, NY, USA), typical vesicle diameter was of 184 nm, PDI 1.2. Preparation of silicon wafers Cut silicon chips (11 11 mm 2 ) were, successively, sonicated in absolute ethanol for 15 min; sonicated in methylene chloride for 15 min twice; cleaned in water : 30 % ammonia : 33 % hydrogen peroxide solution 1:1:1 for 1 hour at 60 o C; thoroughly rinsed with water; kept under water for 24 hours; and dried by nitrogen blowing before use. Nanoparticle deposition Cast deposition was used to apply particles 40 nm and larger onto the surface. Water dispersions of different concentrations for different sizes of particles were used: 0.25 g/l for nm particles; 0.7 g/l for 100 nm particles; 1.8 g/l for 200 nm particles. Dispersions were sonicated for 1 hour before the 1

2 deposition. 20µl drop of nanoparticle suspension evenly spread on dried silicon chip and allowed to slowly dry at room temperature. Spin-assisted deposition was used to apply particles smaller than 40 nm. Concentrations of used solutions were: 0.10 g/l for 4 nm particles; 0.25 g/l for nm particles; 0.75 g/l for 35 nm particles. 100µl drop of nanoparticle suspension was deposited on dry silicon chip and allowed to stand for 1 minute and then spin-coated for 9 minutes at 1000 rpm. The combined procedure was applied for the mixtures of smaller and larger particles. First, smaller particles were deposited by spin-coating. 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 (LB) formation on the surface in comparison with the untreated samples. However, the procedure did ensure similar surface composition over the all range of samples, and provided the fixation of the particles on the surface. To perform the treatment, paper filter was placed inside of the Petri dish cover and was soaked with 20 µl of TEOS. Silicon wafers with deposited silica nanoparticles were placed in the Petri dish, covered, and incubated for 2 hours at 50 o C. After incubation, samples were treated with plasma for 2 minutes (Expanded Plasma Cleaner, W, Harrick Plasma, NY, USA), and kept for 1 hour under Tris-HCl buffer. 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 custom-built temperature stage, and operated in tapping mode in liquid. Samples were scanned using NPS silicon nitride probes (Veeco Instruments, NY, USA) with spring constant of 0.32 N/m, and resonance frequency in aqueous media of ~9 khz. Radius of the tip curvature was determined internally for each experiment series accounting for the overall spherical shape of the recorded particles, and was most often found to be from 25 to 50 nm. Scanning was performed with an amplitude set point in the range of 0.9 to 2.4 V, tapping force of 95-98% from the set point, integral gain , proportional gain 2-6, speed of scanning Hz, and temperature 28.0±0.5 o C (temperature in the fluid cell of 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 silicone rubber O-ring. 50 µl of Tris-HCl buffer was introduced into the fluid cell. A microscope was then equilibrated for about hours to minimize thermal drift of scanner, where 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 of 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 nm, and was easily compensated by the lateral offset command. Images were taken continuously (providing the details of lipid bilayer formation) or periodically (providing the picture of lipid bilayer formed without the disturbance by AFM tip). The best results were obtained when 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 in 20 min; and in the second step 20 µl of the suspension was added, and the image of the lipid bilayer was recorded in 20 minutes. After each addition of the lipid, the tip was retracted by 10V set point. The tip was slowly oscillated with 20 µm amplitude. In this regime the tip did not hinder deposition of the lipid onto the studied surface. The study with bovine liver polar extract was performed at 37±0.5 o C. 2

3 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 suspensions used for the wafer preparation on 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 20x objective lens and 488 nm laser. FITC-insulin ( 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 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. Lipid suspension (0.5 g/l, µl) was deposited on the silicon wafer decorated with nanoparticles, sealed in a vapour-tight Petri dish with filter paper placed inside wetted with Tris-HCl buffer, and incubated for 30 min at 30 o C temperature. Then, the Petri dish with sample was placed into refrigerator (5 o C) for 10 min to provide stable gel-phase DMPC bilayer over the surface. 50 µl of g/l solution of FITC-insulin in Tris-HCl buffer cooled to 5 o C was then added to the drop of lipid suspension and kept at 5 o C for another 30 min (except for the case when the edge of lipid bilayer was taken, where lipid suspension was removed from the sample prior to FITC-insulin addition). Afterwards, the sample was carefully washed with buffer cooled to 5 o C Tris-HCl and placed on microscopic cover glass, facedown without drying, and viewed with 200x magnification. Placement of all the samples of particles on the same wafer ensured the identical conditions of their examination. Data processing AFM images were processed using a WSxM software [Horcas, I. et al. WSXM: A software for scanning probe microscopy and a tool for nanotechnology. Rev. Sci. Instrum. 78, (2007)]. 3D Images were prepared from real AFM topography images using an Autodesk 3ds Max 8 (Autodesk, CA, USA). Subtraction of AFM Images We 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 alignments procedure was performed accounting for thermal or mechanical drift, along slow and fast scan axes. AFM data matrices were loaded from WSxM software format and presented on a screen as raster images (for convenience of operator). 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 by bilinear averaging algorithm; skewing was performed by linear averaging; downscaling was made by the fractional averaging. These algorithms provide average height values based on either averaging of 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. 3

4 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. Alignment in XY-plane was followed by alignment in Z-direction. Regions of the 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 point-by-point basis. For each marked region (j) an average over the region offset values (< Z xyj >) were evaluated. The subtracted data matrix with Z coordinated was corrected by subtraction of z xy values evaluated as z xy = < Z xyj > for uncovered regions and z xy = < Z xyj > 5 nm for regions of the substrate coated by lipid bilayer. Evaluation of particle coverage with lipid bilayer Diameter of particles deposited on Si-wafer was evaluated from the profiles (height) in 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 Fig. S5) was plotted for several groups of particles accounted 24 particles in each group. The particles were assigned to different groups by diameter with 2 nm and 10 nm increments for 6-40 nm and nm ranges of diameters, respectively. We evaluate the coverage of the particles by lipid bilayer based on the evaluation of 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 lipid membrane, corresponding 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 (A P, image pixels) and areas covered by lipid bilayer (A L, image pixels). Simple relation of the pixel quantities 100% A L /A P provided the estimation of partial coverage for each corresponding particle. Large standard deviation bars in the transition zone (see Fig. S5) are conditioned not by the accuracy of the measurements, but by the distribution of membrane coverage over particles of certain diameter. For example, in this range of particle sizes (20-40 nm) one particle could be covered completely, and another one of the same diameter could be not covered at all, thus indicating the transitional region of surface curvature for the lipid membrane deposition. Prolonged time of bilayer preparation and increase of lipid concentration, after the formation of final membrane, did not change this situation. Richards function was applied to fit our experimental data (SRichards2 in Origin 7.5 SR4, OriginLab, USA, MA). This generalized logistic function provided a good fit of the experimental data (coefficient of determination is 0.993). Supporting Discussion This report summarizes the results of the investigation of effects of nanometer-sized roughness on structure of lipid bilayer membrane obtained by fusion of DMPC unilamellar vesicles to the surfaces of different curvature. Flat silica surfaces decorated with silica nanoparticle in a wide range of sizes, from 1 to 200 nm, were used for the model surfaces. Results indicate that studied lipid bilayer forms pores around the nanoparticles larger than 1.2 nm and smaller than 22 nm (Fig. S1a,b). At the same time, lipid membrane is not disrupted by nanoparticles placed on a surface or other surface features below approximately 1.2 nm in diameter and follows over such features (Fig. S1c). The large nanoparticles with diameters over 22 nm are enveloped by lipid 4

5 bilayer (Fig. S1d), unless the surface of the particle is rough when lipid bilayer covering of spherical particles will be incomplete (Fig. S1e). Fig. S1. Schematics of lipid bilayer membrane fusion on rough surfaces: (a) lipid bilayer forms a pore around particles larger than 1.2 nm and (b) smaller than 22 nm whether the nanoparticles are smaller or larger than the thickness of the bilayer; lipid membrane follows the surface topography with nanoparticles below 1.2 nm (c) and envelopes large nanoparticles of more than 22 nm in diameter; (d) there is no information about the exact structure of the lipid bilayer located in the lower hemisphere of the particle in close vicinity to the silica substrate and, so, the schematic drawing of the bilayer for this area in image (d) is speculative. The coverage of bigger particles can be incomplete due to the surface bumps (e). The structure of bilayer in the area marked with circle in the cartoon (d) can not be resolved in the AFM experiments. The findings are based on in situ AFM monitoring of lipid fusion onto silica surfaces with controlled nanometer roughness. The information given here represents raw or minimally processed AFM images, TEM, and confocal microscopy images, as well as other necessary information used to justify the findings of this paper. AFM Studies Figure S2 shows an example of lipid bilayer formation on the Si-wafer decorated with nanoparticles of a 5-20 nm diameter. AFM images (Fig. S2a-c) were aligned. Figures S2b and S2c show topography of the sample partially covered by lipid bilayer, and the surface in the final stage covered by lipid, respectively. The main difference between the topography images in the final and initial stages is a reduced height of the nanoparticles. We subtract the aligned images in order to find this difference and, thus, to determine the location of the deposited lipid bilayer. The subtracted images are shown in the right column (Fig. S2d-f). Figure S2d represents free standing lipid bilayer isles obtained by subtraction of the image (a) from the image (b), and Fig. S2e, obtained by subtraction of the image (a) from the image (c), clearly demonstrates the existence of pores in the lipid bilayer, surrounding 5-20 nm nanoparticles. 5

6 Fig. S2f provides additional information for the analysis of lipid bilayer structure and verifies the accuracy of the subtraction procedure. Fig. S2. Lipid bilayer formation on the sample with 5-20 nm silica nanoparticles: (a) the sample in the initial stage without lipid bilayer; (b) partial coverage of the surface with lipid; (c) the sample in the final stage covered by lipid bilayer; (d) islands of lipid bilayer obtained by subtraction of the image (a) from the image (b); (e) lipid bilayer with holes around nanoparticles obtained by subtraction of the image (a) from the image (c); (f) lipid bilayer obtained by subtraction of the image (b) from the image (c). Figure S3 shows AFM images of lipid bilayer on the Si-substrate decorated by the mixture of several fractions of the particles in size range nm. The AFM images in Fig. S3a-c were aligned. Figures S3b,c show topography of the sample partially covered by lipid bilayer, and the surface in the final stage covered by lipid, respectively. It is difficult to discern the lipid bilayer on the images visually, because much larger particles are present as compared to the experiment shown in Fig. S2. In this instance it was very important to perform image subtraction to visualize deposited lipid bilayer. Fig. S3d shows free standing lipid bilayer isles (marked with blue arrows). Figure S3e demonstrates pores in the lipid bilayer around smaller nanoparticles and on larger particles elevated above the layer (denoted by green arrows). Figure S3f provides the information for the detailed analysis of the lipid bilayer location and gives an additional proof of the coverage of larger particles. Therefore, if the presence of 6

7 bilayer on the particles in Figure S3d was found mistakenly, because of imaging errors, the shape of the spots (denoted by blue arrows) would be different in images Fig. S3d and S3f. Fig. S3. AFM images of lipid bilayer on Si-wafer decorated with a mixture of nm particles: (a) the sample in the initial stage without lipid bilayer; (b) partial coverage of the surface with lipid; (c) the sample in the final stage covered by lipid bilayer; (d) subtracted image (b minus a); (e) subtracted image (c minus a); (f) subtracted image (c minus b). Blue arrows denote examples of areas covered by lipid at the partial coverage. Green arrows denote the examples of areas (dark spots) on large particles, which were not covered by lipid. Experiments presented in Figs. S2, S3, as well as experiments using a mixture of 5-65 nm in size particles as shown in Fig. S4, allowed for the evaluation of the critical diameter of particles where lipid bilayer can be formed. Profiles shown in Fig. S4g demonstrate the particles in the initial stage, which were later covered by lipid bilayer. Detailed analysis of the coverage of particles of different diameters (Fig. S5) have shown that transition region from the absence of coverage to nearly full coverage of the particles spans from about 14 nm to 35 nm. Half of the particles surface area is covered by lipid bilayer for ~22 nm particles. Nanoparticles with a diameter larger than 22 nm are enveloped by lipid bilayer, while smaller 7

8 nanoparticles remain mostly uncovered. Nevertheless, it can be seen that the transition in the coverage vs. particle diameter plot is relatively smooth. Since further addition of lipid suspension and prolonged observation did not lead to the change of coverage, we attribute this change to the specifics of the topography of each individual particle (e.g. irregular shapes, which nanoparticles acquired during their synthesis). Fig. S4. AFM images recorded for the evaluation of the critical diameter of silica particles above which particles are enveloped by lipid bilayer: (a) the sample in the initial stage without lipid bilayer; (b) partial coverage of the surface with lipid; (c) the sample in the final stage covered by lipid bilayer; (d) result of the subtraction of images (b) and (a); (e) result of the subtraction of images (c) and (a); (f) result of the subtraction of images (c) and (b). Blue arrows denote examples of lipid bilayer islands appearing on particles after the partial coverage. Green arrows point to the lipid bilayer which formed 8

9 on particles later. (g) Profiles of particles that later will be covered by lipid bilayer. Numbers in circles correspond to the denoted locations of the profiles in (a). Fig. S5. Particle coverage by lipid bilayer vs. diameter of the particles. Standard deviation bars in the transition zone represent the distribution of coverage over particles of certain diameter. Dotted lines show 50 % coverage. The solid line is a fitting curve (Richards function). Figure S6 presents analysis of lipid membrane on the Si-wafer surface decorated with silica nanoparticles which are smaller than the thickness of the lipid bilayer. Particles smaller than the lipid bilayer are not seen in topography images after the lipid bilayer formation, as it can be concluded when comparing the images before (Fig. S6a) and after (Fig. S6b) lipid deposition. However, AFM tip interaction with different materials provides different phase offsets for tapping mode of the AFM imaging. That allowed us to draw a conclusion about the formation of pores around these particles (Fig. S6c). Representative examples of the profiles obtained for the particles in the initial state with no lipid (denoted with i), and for the lipid bilayer deposited over the particles (denoted with ii) for different particle diameters (in the range nm) show no changes in the bilayer thickness in the locus of particles on the surface (Fig. S7). The topography over many of those particles revealed a small dimple that could be an AFM representation of a pore that is small in diameter, over the particles often hidden by the instrumental noise. Averaging of the 8 profiles over particles smaller than 5 nm in Fig. S7 actually recovered about 0.25 nm deepening in the topography (Fig. S8). Indeed, from the geometrical consideration (Fig. S9b), 30 nm tip, such as used in the experiment presented in Figs. S6-S8, should penetrate the hole over 3 nm in diameter particle by 0.30 nm. That level of penetration is very close to the obtained 0.25 nm value. These results, acquired from phase image and penetration of tip over the particles, mean that the lipid bilayer has formed pores around the particles below 5 nm in diameter. 9

10 Fig. S6. AFM characterization of lipid bilayer coverage of small 1 8 nm silica nanoparticles. Topography of the initial substrate; (a) topography of lipid bilayer covering the substrate; (b) corresponding phase image of the lipid bilayer. Fig. S7. Ten multiple profiles from Fig. S6, numbers in circles correspond to the denoted profiles in Figs. S6a,b. (i) initial surface profile; (ii) profile after the LB formation. 10

11 Fig. S8. Average profile from 8 different scans centered over particles of <5 nm diameter from Fig. S7 (dotted line shows average value of recorded lipid bilayer height). Thus, it was very important to carefully consider the effects of AFM probe in this research. In our experiments, the probes used had nm tip radius after instrument equilibration, initial image recording, and addition of lipid. Fig. S9 shows effects of tip radius on resulting image, considering both small particle and small hole topography. Apparent lateral diameter of the nanoparticles will be significantly larger due to dilation effect. However, height measurements are highly accurate and correspond precisely to the true diameter of nanoparticles. In contrast, the visualization of small holes is limited by the size of AFM tip. Fig. S9 demonstrates the apparent profiles that could be obtained for the porous structure around 3 nm in diameter particle if recorded without instrumental noise for non-elastic sample. Fig. S9. Theoretical curves (red lines) of lateral dilation of 3nm particle and penetration of tip into the pore formed by lipid bilayer around 3 nm particle for the AFM tip with radius of curvature: (a) 50 nm; (b) 30 nm; (c) 10 nm. 11

12 To prove the existence of pores around particles smaller than 5 nm in diameter, we have performed experiments utilizing insulin probes. The selective adsorption of the protein on silica surface (but not on lipid bilayer) was used in this study. Fig. S10 shows an example of the analysis of raw AFM images representing the experiment with insulin probe adsorption on small nanoparticles immersed in lipid bilayer. AFM images in the left and right column are the same sequence of images duplicated for convenience of their analysis. The AFM images are aligned with respect to the initial image (Fig. S10a). It can be seen (Fig. S10b) that small particles are immersed or hidden by the lipid bilayer (for example particles marked with numbers 1-4). At the same time, the lipid layer closely follows smaller surface features on the Si-wafer (denoted by blue arrows). We observed no noticeable changes (Fig. S9c) immediately after the addition of insulin solution into the fluid cell. However, new topographical features appeared over the locations of hidden particles in a timeframe of min, as denoted by red arrows (Fig. S10d,e). As such changes were never noticed without the addition of insulin, these new features are representing insulin molecules, which found and labelled the pores in lipid membrane due to the adsorption onto the nanoparticles exposed to the solution. The same information is depicted in Fig. S10f.1, in terms of the profile plots showing the consequent profiles (marked in the right column of the images). The plots demonstrate the appearance of ~2 nm surface feature over the lipid bilayer. A simple calculation shows that considering the thickness of lipid bilayer and the height of silica nanoparticles, the size of the adsorbed object is about 3.6 nm, which corresponds to the size of the insulin molecule (~3.5 nm). Profiles in Fig. S10f.2-4 demonstrate different situations representing the interaction of particles with lipid membrane. Fig. S10f.2 illustrates, through penetration of the AFM tip, the formation of the pore around a 2 nm particle. The pore formed around the bigger aggregate of small nanoparticles, where the AFM tip revealed the hole, is shown in Fig. S10f.4. It is, therefore, apparent that this big pore was gradually filled up with insulin molecules and its bottom was elevated. In Fig. S10f.3 one can observe an example of 1.2 nm particle which has not pierced the lipid bilayer, but was followed by the membrane, as it is demonstrated by four consequent profiles of lipid bilayer recorded over that particle. 12

13 Fig. S10. Dynamics of insulin adsorption via AFM images representing effect of <5 nm nanoparticles on the structure of lipid bilayer: (a) Initial substrate topography prepared using 1-8 nm particles; (b) substrate coated by lipid bilayer; (c) the sample topography in 6-9 min after insulin ( g/l) addition; (d) in min after insulin addition; (e) in min after insulin addition. Numbers denote the selected particles from which the profiles were taken. Green arrows point to the exact position of numbered particles. Blue arrows denote examples of initial substrate features followed by lipid bilayer. Red arrows show the locus of the adsorption of single insulin molecules. (f) Four multiple profiles taken from (a-e). Numbers in circles correspond to the numbers in images (a-e). (i) Initial substrate topography, from (a); (ii) substrate coated with lipid bilayer, from (b); (iii) the sample topography in 6-9 min after insulin ( g/l) addition, from (c); (iv) the sample topography in min after insulin addition from (d); (v) the sample topography in min after insulin addition, from (e). iii, iv, and v profiles were artificially shifted with respect to ii profiles for convenience. 13

14 AFM experiment with polar lipid extract from bovine liver In order to check if the formation of pores is relevant to the mixtures of lipids in cell membranes, we have performed the study with a polar lipid extract from bovine liver representing a mixture of lipids. The results are shown in Fig. S11. All original AFM images and images-differences in this figure are aligned with respect to the initial image. It can be seen from Fig. S11e that in this case the formation of the lipid bilayer is similar to that observed for DMPC. Presence of smaller particles leads to the formation of holes in the lipid bilayer (white arrows), whereas lipid bilayer can envelope larger particles (e.g. 65 nm, blue arrows). Fig. S11. AFM images of the bilayers formed by polar lipid extract from bovine liver at 37 o C on Siwafer decorated with a mixture of 1-65 nm particles: (a) the sample in the initial stage without lipid bilayer; (b) partial coverage of the surface with lipid; (c) the sample in the final stage covered by lipid bilayer; (d) subtracted image (b minus a); (e) subtracted image (c minus a); (f) subtracted image (c minus b). Green arrows denote the rear case of observed vesicle on the surface before lipid bilayer formation. White arrows denote examples of pores in bovine liver lipid bilayer around 8-15 nm particles. Blue arrows denote the island of bovine liver lipid bilayer formed on 65 nm particle. Analysis of surface topography of the nanoparticles with TEM Transmission electron micrographs of the silica particles are shown in Fig. S12. Silica nanoparticles of 50 nm and 100 nm diameter (Fig. S12a) revealed rough surface with an average bump height of 3 nm, with the highest height of 5 nm. Particles of 200 nm diameter had a very smooth surface (Fig. S12b). Evidently, surface morphology of the nanoparticles can play a crucial role in the efficiency of lipid bilayer coatings on beads and nanoparticles. Rough surface of the nm particles can lead to the creation of pores in lipid bilayer that were observed on AFM images described above (Fig. S3e). Indeed, 14

15 the experiments with the fluorescent protein probe presented below indicate that large smooth particles caused less number of defects in lipid bilayer. Fig. S12. TEM image of silica nanoparticles: (a) 100 nm and (b) 200 nm in diameter demonstrating different surface roughness. Study of the lipid membranes structure with labelled proteins In order to obtain additional evidence of selective lipid bilayer formation on particles of different sizes, we have performed the confocal microscopy study with FITC-labelled insulin as molecular probe. Confocal microscopy images in Fig. S13 were obtained according to the procedure described above. Fig. S13a illustrates that FITC-insulin readily adsorbs on a silica surface free of lipid bilayer, but does not stain the area with lipid bilayer. Samples with small nanoparticles (1-20 nm in diameter) manifest intense fluorescence, due to the significant number of pores formed by those nanoparticles (Fig. S13b,c). Fluorescence of the sample with large nm nanoparticles (Fig. S13d) is only slightly less pronounced than for small nanoparticles. The particles with adsorbed labelled protein are clearly visible, most likely because of the pores formed in lipid bilayer around their surface bumps, as shown in Fig. S3e and Fig. S12. Those pores are available for the adsorption of FITC-insulin on exposed silica surfaces. The sample with 200 nm nanoparticles that has a very smooth surface, as shown with TEM (Fig. S12b), has only faint fluorescence in the areas with deposited nanoparticles (Fig. S13e), once again confirming that smooth nanoparticles would be fully covered with lipid bilayer. 15

16 Fig. S13. Confocal microscopy images of FITC-insulin adsorption on (a) uncoated silica wafer neighbouring with lipid bilayer, and silica nanoparticles of different sizes (lipid bilayer is formed over the samples observed in images (b-e)); (b) 1-8 nm particles, (c) 5-20 nm particles, (d) nm, (e) 200 nm particles. Corresponding AFM images (in the right column) visualize the particles of different diameters in the samples: (f) 1-8 nm, (g) 5-20 nm, (h) nm, (i) 200 nm. Images in Fig. S13a-e were intentionally taken near the border line formed by the dried drops of particle dispersions, in order to compare different samples of particles with SLB on the polished surface of the Si-wafer. Edges of the former drops are always stained brighter due to much higher concentration of nanoparticles in those areas. Fluorescent confocal microscopy additionally confirmed that lipid bilayer forms the pores around smaller particles, and high-curvature surface features of larger particles. 16

17 Computational model of bilayer behaviour on surfaces A three dimensional macroscopic model of supported lipid bilayers has been developed which uses a Brownian dynamic framework for looking at membrane behaviour as a function of surface topology. r dv r r r m = FDISSIPATIV E + FRANDOM + FCONSERVATI VE dt The dissipative force is handled by a Newtonian viscosity model, with a random force corresponding to the system temperature and a conservative force having contributions from intra-membrane interactions (like bending, elastic deformation, etc.) and inter membrane surface interactions. The membrane and the surface is treated as a three dimensional continuum as shown below (Fig. S14) where the surface can have different topologies (hemispherical, sinusoidal, random, etc.). Fig. S14. Sketch of bilayer and substrate representation. Each element of the membrane typically represents ~ 100 lipid molecules. The model was used to study dynamics of bilayers with a thickness of ~ 5 nm, and lateral dimensions of 100 µm x 100 µm. The membrane and substrate properties were assumed as follows: bending modulus of 5e-20 J, membrane elasticity of N/m, membrane substrate interaction was 0.8mN/m and membrane substrate equilibrium separation of 3.5 nm. The membrane properties were computed using coarse grained representation of the bilayer and the membrane substrate interaction parameters were obtained via fully atomistic simulations and Surface Force Apparatus (SFA) measurements. The membrane behaviour was studied over time periods of a few milliseconds. From the simulations the average minimum and maximum separation of the membrane from the silica surface was extracted for different surface roughnesses. 17

18 Fig. S15. Membrane conformation as a function of surface For a surface roughness of 3 nm or less, the range of separations of the membrane, plotted using the maximum and minimum values, is quite narrow. Such a range is expected due to the dynamical fluctuation of the membrane, induced by impinging solvent molecules (assumed here as a random noise). At these small values of roughness, the membrane conforms to the substrate at a distance equal to our assumed equilibrium value, with all deviations coming from fluctuations. For a surface roughness greater than 3 nm, the simulations show that membranes can no longer conform to the change in the surface curvature (Fig. S15). Our model suggests that this inability to conform to substrate features is due to the high bending penalty paid by the membranes, as opposed to the attractive force pulling the membrane to the surface. Due to computational reasons, we were unable to study membranes adsorbing on features much larger than the range of roughness shown. It must be emphasized that the model tries to mimic cell membrane fragments, which retain their integrity due in part to the presence of additional biomolecules. This model membrane is different from supported bilayers formed through the adsorption of individual lipids. In the latter case, lipids thermodynamically adsorb to the substrate to form supported bilayers, but probably do not form free bilayers in solution that adsorb onto the substrate. As a result, pore formation may be observed when lipids adsorb around features of certain sizes. This is the essential difference between the experimental and computational studies shown. In spite of the differences in their schematics, the physical effects seen in the membrane simulations can be compared to experimental observations of the supported bilayers. Primarily, both studies show that the bilayers will likely conform to substrate features that are very small (~1.2 nm in experiments, and ~3 nm in simulations), while a balance between bending energy and adhesion prevent such a conformation from occurring on particles or features larger than this value. The difference between the model and experimental results for the critical threshold of coating may possibly be due the difference in the integrity of the model and experimental bilayers. The agreement is reasonably close to be able to analyze the physical process involved. As the surface curvature increases the model membrane does not display pore formation. Instead, it simply no longer conforms to the surface topography. 18