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Atomic Force Microscopy of Cationic Liposomes Neil H. Thomson,† Ian Collin,‡ Martyn C. Davies,† Karen Palin,‡ David Parkins,‡ Clive J. Roberts,*,† Saul J. B. Tendler,† and Philip M. Williams† Laboratory of Biophysics and Surface Analysis, School of Pharmaceutical Sciences, University of Nottingham, University Park, Nottingham NG7 2RD, U.K., and Glaxo Wellcome Research and Development, Park Road, Ware, Hertfordshire SG12 0DP, U.K. Received September 21, 1999. In Final Form: February 14, 2000 Cationic liposomes, formed from a 1:1 molar mixture of p-ethyldimyristoylphosphatidylcholine (EDMPC) and cholesterol by extrusion through filters, have been investigated using atomic force microscopy (AFM). The fluid nature of liposomes makes them a challenging prospect for imaging by AFM. Tapping-mode imaging in air and under fluid was carried out with the liposomes deposited on either mica or mica treated with aminopropylsilane (AP-mica), surfaces with negative and positive potentials, respectively. The interaction of the liposomes with both surfaces was investigated, and the observed liposome structure was found to be dependent on the nature of the surfaces. Structures close to those expected in solution from light scattering measurements were observed on the AP-mica surfaces. Tip-induced fusion of individual liposomes could also be observed. This study illustrates that AFM can be used as a tool for studying dynamic processes involving lipid vesicles.
Introduction Atomic force microscopy (AFM)1 imaging of biological entities and molecules2,3 relies on successfully immobilizing them to a surface on which they can be distinguished (i.e., the surface must have a roughness that is small compared to the size of the entity). The interaction between the entity and surface needs to be greater than that between the tip and the entity for successful and reproducible imaging. Usually one is interested in minimizing the interaction between the entity and the surface so that its solution structure is not compromised. AFM is a technique that can image surfaces at very high resolution under aqueous environments.4 If the entity is not significantly disturbed by immobilization, then one can be confident that an observed structure is close to its structure when it is “free” in solution. The aim of this work was to determine the solution structure of cationic liposomes through imaging in air and liquid on two surfaces with different surface potentials, namely, mica and mica treated with aminopropylsilane (AP-mica). Mica is negatively charged in solution with a surface charge density of -0.009 C/m2,5 and AP-mica is positively charged with an estimated surface potential of about 7mV in 0.01 M salt.6 By use of the Grahame equation for low surface potentials,7 the potential on AP-mica converts to a surface charge density of 0.0016 C/m2. Liposomes (also known as lipid vesicles) * To whom correspondence should be addressed. E-mail:
[email protected]. † University of Nottingham. ‡ Glaxo Wellcome Research and Development. (1) Binnig, G.; Quate, C. F.; Gerber, Ch. Phys. Rev. Lett. 1986, 56, 930. (2) Shao, Z.; Yang, J. Q. Rev. Biophys. 1995, 28, 195. (3) Hansma, H. G.; Hoh, J. H. Annu. Rev. Biophys. Biomol. Struct. 1994, 23, 115. (4) Drake, B.; Prater, C. B.; Weisenhorn, A. L.; Gould, S. A. C.; Albrecht, T. R.; Quate, C. F.; Cannell, D. S.; Hansma, H. G.; Hansma, P. K. Science 1989, 243, 1586. (5) Pashley, R. M. J. Colloid Interface Sci. 1981, 80, 153. (6) Lyubchenko, Y. L.; Shlyakhtenko, L. S. Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 496. (7) Israelachvili, J. N. In Intermolecular and surface forces: with applications to colloidal and biological systems; Academic Press: London, 1985.
are a very challenging sample for current AFM technology since they are fluid and dynamic systems. Cationic liposomes are important as potential delivery vehicles for DNA in gene therapy techniques. They avoid immunogenic problems associated with viral delivery, are simple to prepare, and can carry DNA with practically no size limit.8 A limited AFM study has been carried out in air, investigating the effect of different cholesterol derivatives on liposome size for gene delivery.9 A correlation was made between liposome size and transfection efficiency in vitro. Recently, a novel cationic lipid, p-ethyldimyristoylphosphatidylcholine (EDMPC), when mixed with cholesterol (1:1 molar ratio) has been shown to be efficient at in vivo transfection provided that the liposomes are less than about 200 nm in diameter.10 There was a lack of positive correlation between in vivo and in vitro transfection, suggesting that the transfection mechanisms are different in the two cases. To gain an understanding of the structure/ function relationship of these liposomes complexed with DNA, it is desirable to directly image their structure. This paper outlines AFM techniques for investigating the structure of these EDMPC/cholesterol liposomes and their interaction with mica and AP-mica surfaces. Only a handful of papers have been published previously on imaging either liposomes or lipid vesicles isolated from biological material. Liposomes made of a mixture of dipalmitoyl phosphatidylcholine (DPPC) and cholesterol in fluid have been imaged using contact mode AFM.11 The liposomes were bound to gold surfaces using an immunochemical technique, to prevent the liposomes from being swept away by the tip. The image contrast was force dependent, and the liposomes were swept away above a threshold load force of 4.5 nN. Singh and Keller12 (8) Singhal, A.; Huang, L. In Gene Therapy: from laboratory to the clinic; Hui, K. M., Ed.; World Scientific: Singapore, New Jersey, London, New York, 1994; Chapter 6. (9) Kawaura, C.; Noguchi, A.; Furuno, T.; Nakanishi, M. FEBS Lett. 1998, 421, 69. (10) Gorman, C. M.; Aikawa, M.; Fox, B.; Fox, E.; Lapuz, C.; Michaud, B.; Nguyen, H.; Roche, E.; Sawa, T.; Wiener-Kronish, J. P. Gene Ther. 1997, 4, 983. (11) Shibata-Seki, T.; Masai, J.; Tagawa, T.; Sorin, T.; Kondo, S. Thin Solid Films 1996, 273, 297. (12) Singh, S.; Keller, D. J. Biophys. J. 1995, 60, 1401.
10.1021/la991256p CCC: $19.00 © 2000 American Chemical Society Published on Web 04/27/2000
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successfully imaged dipalmitoyl phosphatidylethanolamine (DPPE) vesicles adsorbed on mica in air. They also found that DPPC vesicles collapsed in air to form layers on the mica. DPPE vesicles have now been imaged under liquid in contact mode without collapse, but the imaging was not very clear.13 With an increase of the contact force, the liposomes collapsed into patches. A number of groups have imaged supported lipid layers using AFM.14-23 Langmuir-Blodgett (LB) films can be imaged with molecular or submolecular resolution.14-16,24 Only one study using tapping-mode in fluid has enabled the imaging of intact liposomes; in this case, DPPC physisorbed on mica, formed bilayers and areas of individual liposomes.25 Concerning biological rather than “synthetic” samples, synaptic vesicles have been successfully imaged26,27 and their elastic properties have been probed using the forcemapping technique.28 Under isoosmotic buffers these vesicles could be imaged intact, reproducibly on mica, but collapsed on introduction of hypoosmotic buffers to the AFM fluid cell. Most previous studies have been motivated by the possibility of using supported membranes for studying membrane proteins by AFM. Our goal was to achieve reproducible imaging of individual liposomes, which would enable the structures of gene therapy vectors formed using liposomes to be studied using AFM. In this paper, we demonstrate that using tapping-mode imaging, crosslinking of the liposomes to the surface is not essential, but control of the chemistry of the supporting surface is paramount. Importantly, we observed fusion of individual liposomes occurring, which demonstrates that AFM can be used as a tool for monitoring vesicle fusion events. Materials and Methods Liposomes. Cationic liposomes were formed using a 1:1 molar ratio of p-ethyldimyristoyl phosphatidylcholine (EDMPC) and cholesterol. Liposomes of very small dimensions were obtained by extrusion through a series of filters with smaller and smaller pores.29 The smallest pore size used was 50 nm in diameter, and (13) Mu¨ller, D. J.; Amrein, M.; Engel, A. J. Struct. Biol. 1997, 119, 172. (14) Egger, M.; Ohnesorge, F.; Weisenhorn, A. L.; Heyn, S.-P.; Drake, B.; Prater, C. B.; Gould, S. A. C.; Hansma, P. K.; Gaub, H. E. J. Struct. Biol. 1990, 103, 89. (15) Zasadzinski, J. A. N.; Helm, C. A.; Longo, M. L.; Weisenhorn, A. L.; Gould, S. A. C.; Hansma, P. K. Biophys. J. 1991, 59, 755. (16) Weisenhorn, A. L.; Egger, M.; Ohnesorge, F.; Gould, S. A. C.; Heyn, S.-P.; Hansma, H. G.; Sinsheimer, R. L.; Gaub, H. E.; Hansma, P. K. Langmuir 1991, 7, 8. (17) Hansma, H. G.; Weisenhorn, A. L.; Gould, S. A. C.; Sinsheimer, R. L.; Gaub, H. E.; Stucky, G. D.; Zaremba, C.; Hansma, P. K. J. Vac. Sci. Technol., B 1991, 9, 1282. (18) Brandow, S. L.; Turner, D. C.; Ratna, B. R.; Gaber, B. P. Biophys. J. 1993, 64, 898. (19) Birdi, K. S.; Vu, K. T. Langmuir 1994, 10, 623. (20) Tamm, L. K.; Bo¨hm, C.; Yang, J.; Shao, Z.; Hwang, J.; Edidin, M.; Betzig, E. Thin Solid Films 1996, 284-285, 813. (21) Fang, Y.; Yang, J. J. Phys. Chem. 1996, 100, 15614. (22) Dufreˆne, Y. F.; Barger, W. R.; Green, J.-B. D.; Lee, G. U. Langmuir 1997, 13, 4779. (23) Fang, Y.; Yang, J. Biochim. Biophys. Acta 1997, 1324, 309. (24) Muscatello, U.; Valdre, G.; Valdre, U. J.f Microsc. 1996, 182 (3), 200. (25) Solletti, J. M.; Botreau, M.; Sommer, F.; Brunat, W. L.; Kasas, S.; Tran Minh Duc; Celio, M. R. Langmuir 1996, 12, 5379. (26) Parpura, V.; Doyle, R. T.; Basarsky, T. A.; Henderson, E.; Haydon, P. G. Neuroimage 1995, 2, 3. (27) Garcia, R. A.; Laney, D. E.; Parsons, S. M.; Hansma, H. G. J. Neurosci. Res. 1998, 52, 350. (28) Laney, D. E.; Garcia, R. A.; Parsons, S. M.; Hansma, H. G. Biophys. J. 1997, 72, 806. (29) Hope, M. J.; Bally, M. B.; Webb, G.; Cullis, P. R. Biochim. Biophys. Acta 1985, 812, 55.
Thomson et al. Table 1. Contact Angles of Water on Mica Taken after the Mica Had Been Incubated with Liposome Dispersions of Various Concentrations, Rinsed, and Dried (Average of Five Measurements) concn of EDMPC or cholesterol in the liposome suspension (mM)
static contact angle on dried surfaces measured with deionized water (deg)
20 2 0.2 0.02
81.8 ( 3.3 84.2 ( 5.8 77.2 ( 10.8 80.7 ( 7.2
this yielded liposomes with a narrow distribution around 80 nm in diameter as measured by laser light scattering (Brookhaven Zeta Plus analyzer). Mica. Mica (Agar Scientific, Stansted, Essex, U.K.) was freshly cleaved using adhesive tape, and dispersions of the liposomes were deposited in precise volumes using GILSON pipets at various concentrations, between 20 µM and 20 mM of EDMPC content. The solution was incubated on the surface for a given time. For imaging in air, the surfaces were rinsed with excess 10 mM Tris buffer (pH ) 8) and dried gently under a stream of dry nitrogen gas. For imaging in solution the same rinsing procedure was followed, but the surfaces were kept wet and introduced as such, into the AFM fluid cell. AP-Mica. AP-Mica was prepared by incubating freshly cleaved mica with 2 µL of pure aminopropylsilane (SIGMA, St. Louis, MO) in a plastic Petri dish under a dry nitrogen environment for 2 h. This produces vapor-deposited silane surfaces as previously described.30 The solutions for imaging were applied to these surfaces as described for the mica above. AFM Imaging. All imaging was performed using a Digital Instruments Nanoscope III Multimode system (Santa Barbara, CA) with a vertical engage E scanner (maximum range 10 × 10 × 2.5 µm). A commercial glass fluid cell was used without an O-ring for imaging under buffer. Oxide-sharpened silicon nitride tips with integrated cantilevers with nominal spring constants of 0.38 N/m were used for imaging in both contact mode and fluid tapping-mode. For contact mode operation, the force was initially minimized after approach with zero scan size before imaging was commenced. For fluid tapping mode, the cantilevers were excited acoustically, through the cell and liquid, indirectly, at frequencies between 9 and 10 kHz (which is just below their resonant frequency in water). For tapping mode in air, silicon levers with nominal spring constants of about 30 N/m and resonant frequencies between 250 and 300 kHz were used. The lever was excited just below its resonant frequency. The tapping was light, as defined by Magonov et al. 31 Image Processing. Processing of images to determine liposome size and distribution was carried out using SXM Image, version 1.61 (Steve Barrett, 1998). Initially, images were flattened in the Nanoscope software using first-order fitting to remove sample tilt. The sizes of the liposomes were determined using freehand drawing around each liposome in SXM Image. This software can then calculate the number of pixels contained in the selection. The size was expressed as the diameter of a circle of equivalent area. Repeated measurements on individual liposomes indicated that this method gave about a 5% error in the determination of the projected area. Contact Angles. Contact angles were measured by placing a 2 µL drop of distilled water on the mica or AP-mica surfaces, with and without the addition of liposomes. The angle that the plane of the substrate surface made with the surface of the water droplet was determined using a horizontal microscope with a rotatable crosshair in the objective. Five measurements were made on different areas of the surface and averaged. Measurements were made as rapidly as possible after the addition of the water drop before relaxation of the drop under gravity occurred. (30) Lyubchenko, Y. L.; Gall, A. A.; Shlyakhtenko, L. S.; Harrington, R. E.; Jacobs, B. L.; Oden, P. I.; Lindsay, S. M. J. Biomol. Struct. Dyn. 1992, 10, 589. (31) Magonov, S. N.; Elings, V.; Whangbo, M.-H. Surf. Sci. 1997, 375, L385.
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Figure 1. Tapping-mode image in air of a mica surface after exposure to a liposomes dispersion of EDMPC concentration of 20 µM. The surface is essentially featureless, and liposome dispersions spread from higher concentrations have a similar appearance.
Results and Discussion Liposomes on Mica. The behavior of the liposome dispersion was investigated for 4 orders of magnitude of concentration, from the stock solution (20 mM) down to a 1000-fold dilution. Droplets of constant volume (20 µL) were deposited onto freshly cleaved mica, and their spreading behavior was observed. The droplets achieved equilibrium contact angles between 77° and 84° with the mica but took varying amounts of time to achieve equilibrium. At the highest concentration the droplet did not spread, and equilibrium was achieved almost instantaneously. A 10-fold dilution droplet partially spread before contracting within 2 s. A 100-fold dilution droplet spread over a wider area than the 10-fold dilution and then contracted to its equilibrium position within 23 s. The
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1000-fold dilution spread completely over the whole piece of mica before slowly contracting, taking approximately 5 min to achieve equilibrium. The varying time scales for equilibration reflect the kinetics of the liposome spreading. A representative AFM tapping-mode image in air of these surfaces after rinsing and drying is shown in Figure 1. It is smooth and devoid of features suggesting a homogeneous lipid layer. Table 1 summarizes measurements of the static contact angles with water taken on these dried surfaces. Within standard deviations, the contact angles are the same, suggesting that the final structure of the layer formed on the mica is very similar in each case. Note that the lower concentrations have larger standard deviations, suggesting less homogeneity in coverage. Comparing these values with the contact angle on mica, which is essentially zero, suggests that the hydrophobic chains of the lipids are exposed at the surface. This is corroborated by the observation that on AP-mica after incubation with the liposomes the contact angle is very small, indicating a hydrophilic surface with the charged liposome headgroups uppermost (see below). To investigate the manner in which the liposomes contact the surface and form such a layer, a time series of AFM images of the liposomes interacting with the mica was taken at the 1000-fold dilution. Equivalent volumes were deposited on equal areas of freshly cleaved mica (in practice this was achieved by recleaving the same piece of mica), left for a given time and then rinsed with excess buffer, gently dried in a stream of dry nitrogen gas, and imaged using tapping-mode in air. Figure 2 shows a series of images obtained from such an experiment. As one can see from the images and corresponding cross sections, the liposomes come down to the surface at many places and then they slowly collapse and rearrange their structure to form flat patches with the hydrophobic chains exposed.
Figure 2. A series of tapping-mode images in air taken after different incubation times of the 20 µM (1000-fold dilution of stock) liposome dispersion with the mica. The incubation time is shown in the top right of each image in seconds. The height scales (Z) for the cross-section plots are (50 nm for (a) to (d), (25 nm for (e), and (5 nm for (f) to (h). The liposomes deposit randomly on the surface and then slowly collapse and fuse to form a monolayer on the mica. In contrast to Figure 1, if the liposome solution is left longer than the time required to form a monolayer, excess liposomes can be seen on the surface (h).
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Figure 3. Tapping-mode image taken under the 10 mM Tris buffer (pH ) 8) showing that individual liposomes can be resolved on an AP-mica surface. The liposomes close pack to form a layer on the surface from which the size of the liposomes can be readily determined. After scanning at high resolution (a) for a few images the liposomes are induced to fuse by the tip, which is observed by zooming out to a lower resolution (b).
Figure 4. Representative tapping-mode in air image of liposomes on AP-mica afer drying. The liposomes have fused together to form multiple bilayers on the surface. Eight levels of these bilayers can be seen in this image.
If the solution is incubated on the surface for long enough, then a complete layer is formed and an image similar to that in Figure 1 is obtained (see Figure 2h). In contrast to Figure 1, there are some liposomes on top of the layer of collapsed liposomes. This indicates that it is possible for liposomes to adhere to the collapsed layer. These were not seen in the dilution series because the mica was rinsed very soon after the drop of liposome solution reached equilibrium. Longer incubation times (600 s in Figure 2h) allow time for additional binding of the liposomes to the mica/liposome surface. Both the dilution series and the time series at high dilution (1/1000) suggest that the layer is formed from the collapse of single liposomes which then spread and coalesce on the mica. The process of collapse in liquid is on a far too short time scale to be captured through direct imaging in solution. Liposomes on AP-Mica. On AP-mica, the liposomes are stable enough to be imaged intact without collapse. Tapping-mode in fluid gives the highest resolution images on these samples. Figure 3 resolves individual liposomes closely packed on the surface. The indistinct edges of the liposomes are attributed to tip-induced disturbance of the
soft, viscoelastic liposomes. Indeed, continued scanning at high enough magnification induces fusion of the lipsomes on the surface. Figure 3a shows the effect of tapping at 1 × 1 µm (256 × 256 pixels) for more than one image and then zooming out to 2 × 2 µm (Figure 3b). In the initial image, single liposomes were resolved, but on subsequent scans, the liposomes were seen to fuse into larger nonspherical structures. Zooming out enables the fused area to be reproducibly imaged. The slow scan frequency (i.e., rate at which scan lines are acquired) and the pixel density were kept constant and the area increased 4-fold; therefore the energy input per unit surface area was decreased 4-fold. In reality, dissipation into the sample must be localized around the tip-sample contact area. Since the tapping frequency, scan frequency, and pixel resolution remained the same, the number of taps that the tip makes per pixel on the surface is constant for both images (about 10 in this case). Fusion of liposomes was not seen in the lower magnification image, which implies that not enough energy is dissipated per tap to cause adjacent liposomes to fuse. This means that in the 1 × 1 µm image the contact area of the tip must be larger than the pixel area, enabling more energy to be dissipated per
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Figure 5. Cross-section analysis of a software zoom of the image in Figure 4 showing (a) the height of the lipid layers to be about 6 nm, which is consistent with a bilayer, and (b) some vesicle-like structures which have not been completely incorporated into one of the bilayers.
Figure 6. Distribution of liposome diameters as measured from images such as those in Figure 3. The spread of liposome size is fairly narrow and has a mean and standard deviation of 70.5 ( 8.4 nm. The median value is 69.6 nm.
unit area, thereby causing the liposomes to fuse. The pixel area for each of the images is 15 and 60 nm2, respectively, which means that the effective contact area of the tip to the surface, or perhaps, more accurately, the area over which the dissipation takes place, is between 15 and 60 nm2. These data also imply that the relaxation time of the liposomes is longer than the pixel dwell time, about 1 ms in this case. Clear images of the liposomes, such as those in Figure 3, were not obtained on every sample, and after scanning for a few minutes, the imaging often became obscured through tip contamination. The reason for this irreproducibility is clear once a sample has been dried and imaged in tapping-mode in air. Figure 4 shows a typical image of
liposomes deposited on AP-mica after it has been dried and imaged using tapping mode in air and reveals large extended flat areas divided, in this case, into eight levels in the 10 × 10 µm area. Cross-sectional analysis indicates that these layers are 6.0 ( 1.5 nm high (see Figure 5a), consistent with bilayers of the lipids. X-ray studies of dimyristoyl phosphotidylcholine (DMPC) multilamellar films have determined the bilayer thickness to be about 5.3 nm, depending upon the temperature and the level of hydration.32,33 The bilayer thickness of cholesterol is (32) Smith, G. S.; Sirota, E. B.; Safinya, C. R.; Plano, R. J.; Clark, N. A. J. Chem. Phys. 1990, 92 (7), 4519. (33) Harroun, T. A.; Heller, W. T.; Weiss, T. M.; Yang, L.; Huang, H. W. Biophys. J. 1999, 76, 937.
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smaller, at 3.4 nm.34 To our knowledge, the multilamellar structures of EDMPC and EDMPC/cholesterol mixtures have not been investigated to date, but since EDMPC has an additional ethyl group compared with DMPC, we might expect the bilayer thickness of EDMPC to be a little larger than 5.3 nm. The fact that the lipid headgroups are uppermost on AP-mica is supported by contact angle measurements which showed that after application of lipids, the surface became more hydrophilic. On bare AP-mica the contact angle of water was 36.6 ( 3.4° (average of five measurements). After incubation with a liposome dispersion the contact angle was too low to be accurately measured, approximately 5°. On some layers there were globular features consistent in diameter with individual liposomes (see Figure 5b). It is assumed that these are collapsed and have no appreciable water contained within them since they are also about 6 nm high and the sample was dry. Little detail could be resolved within the layers suggesting that they are fully fused bilayers, which presumably occurs on drying, although spontaneous fusion was also observed while imaging in contact-mode under buffer. The liposomes on AP-mica could be imaged using contact-mode producing images not dissimilar from Figure 3, though not quite as clear (data not shown). No obvious tip-induced fusion was observed in contact mode. However, larger scans (> 2 × 2 µm) showed patches where the liposomes had apparently spontaneously fused on the surface. These patches were about 1 nm higher in these images than the rest of the close-packed liposome layer. This could be due to a real change in the height due to a geometry change in the lipid or a change in the tip-sample interaction as a consequence of the geometry change. The sizes of the liposomes imaged in fluid tapping-mode on AP-mica (see Figure 3) were measured. Figure 6 shows the distribution of the liposome diameters. The liposomes in Figure 3 do not appear exactly spherical due to their motion and/or distortion by the tip. To estimate their diameters, the perimeter of each liposome was drawn by (34) Craven, B. M. Nature 1976, 260, 727.
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hand, the area encompassed was determined, and the corresponding diameter of an equivalent circle was calculated. These are the values that are plotted in Figure 6. The mean and standard deviation of the distribution are 70.5 ( 8.4 nm, and the median value is 69.6 nm. This is comparable with light scattering data, which yielded a diameter of 80 nm. Tip convolution does not appear to be significant because the liposomes are in a close packed layer, and in such cases the AFM will correctly measure periodicity. The interaction of these EDMPC/cholesterol cationic liposomes with a negatively charged mica surface and a postively charged AP-mica surface are quite different. On the basis of the experiments presented here, we suggest that the liposomes bind randomly to the surface, controlled by diffusion in solution. On mica they collapse over about 5 min to form a fused monolayer on the surface with the hydrophobic chains exposed. On AP-mica the liposomes pack closely together on the surface to form multilayers, which spontaneously fuse to form bilayers on the time scale of minutes. The liposomes can be induced to fuse by increasing the energy input to the surface via the AFM tip in tapping-mode by altering the magnification. Conclusions Individual cationic liposomes have successfully been imaged by AFM using fluid tapping-mode on positively charged AP-mica surfaces. The observed structure is strongly dependent upon the interaction the liposomes have with the surface. On AP-mica the liposomes have dimensions comparable with those measured by light scattering in solution. Fusion of the liposomes on these surfaces is spontaneous, although slow, but can be induced by the AFM tip. Successful imaging of the liposomes in fluid illustrates the potential AFM has for determining structures of gene therapy vectors that use cationic liposomes as the delivery vehicle. Acknowledgment. The postdoctoral funding for N.H.T. was supplied by Glaxo Wellcome. LA991256P
