Probing small unilamellar EggPC vesicles on mica surface by atomic force microscopy

Colloids and Surfaces B: Biointerfaces 34 (2004) 41–51

Probing small unilamellar EggPC vesicles on mica surface by atomic force microscopy Xuemei Liang, Guangzhao Mao, K.Y. Simon Ng∗ Department of Chemical Engineering and Materials Science, Wayne State University, 5050 Anthony Wayne Drive, Detroit, MI 48202, USA Accepted 30 October 2003

Abstract Sonicated small unilamellar egg yolk phosphatidylcholine (EggPC) vesicles were investigated using atomic force microscopy (AFM) imaging and force measurements. Three different topographies (convex, planar, and concave shape) of the EggPC vesicles on the mica surface were observed by tapping mode in fluid, respectively. It was found that the topography change of the vesicles could be attributed to the interaction force between the AFM tip and vesicles. Force curves between an AFM tip and an unruptured vesicle were obtained in contact mode. During approach, two breaks corresponding to the abrupt penetration of upper and lower bilayer of vesicle were exhibited in the force curve. Both breaks spanned a distance of around 4 nm close to the EggPC bilayer thickness. Based on Hertz analysis of AFM approach force curves, the Young’s modulus (E) and the bending modulus (kc ) for pure EggPC vesicles were measured to be (1.97 ± 0.75) × 106 Pa and (0.21 ± 0.08) × 10−19 J, respectively. The results show that the AFM can be used to obtain good images of intact and deformed vesicles by tapping mode, as well as to probe the integrity and bilayer structure of the vesicles. AFM force curve compare favorably with other methods to measure mechanical properties of soft samples with higher spatial resolution. © 2003 Elsevier B.V. All rights reserved. Keywords: Topography; Sonicated unilamellar vesicle; Elastic property; Force curve; Atomic force microscopy

1. Introduction Liposome technologies are very promising for highly sophisticated pharmaceutical products. In addition to the medical applications of the liposome, there is a broad spectrum of non-medical applications, such as cosmetics and biosensors [1–3]. Most recent liposome investigations use homogeneous unilamellar vesicles in the size range 50–150 nm because this size range is a compromise between loading efficiency of liposomes (which increases with increasing size), liposome stability (which decreases with increasing size), and ability to extravasate (which decreases with increasing size) [2]. Unilamellar liposomes are commonly obtained from the corresponding multilamellar vesicle (MLV) dispersions either by sonication [4] or extrusion [5]. From both the fundamental and applied perspectives, it is very important to have an in-depth understanding of the behavior of the liposome [3]. One of the primary issues is the topography and stability of vesicles, especially upon adsorption on ∗ Corresponding author. Tel.: +1-313-577-3805; Fax: +1-313-577-3810. E-mail address: [email protected] (K.Y. Simon Ng).

0927-7765/$ – see front matter © 2003 Elsevier B.V. All rights reserved. doi:10.1016/j.colsurfb.2003.10.017

a solid substrate. Much understanding of liposome stability has been gained from theoretical study [6–8] and mechanical measurement [9–14] of liposomes. Atomic force microscopy (AFM) allows simultaneous imaging and force measurement of soft, biomaterial interfaces, with dimensional resolution approaching one angstrom and force resolution approaching 10−12 N [15]. AFM images on liposome are quite challenging since they are soft and dynamic. Thus, definitive visualization of individual vesicles is still one of the most challenging tasks to date, due to vesicle deformation during scanning and possible artifacts induced by the AFM tip in contact mode [16–19]. Several papers found that both contact and tapping mode were able to capture soft samples with tapping mode showing slightly better resolution [20–22]. To the best of our knowledge, the tip effect on topography evolution by the tapping mode in fluid has not been reported. Moreover, obtaining high-resolution images of vesicles would be a first step toward the study of vesicle adsorption behavior and formation of the supported phospholipid bilayers (SPBs) process. Bending rigidity is a fundamental macroscopic property of lipid bilayers which is related to the stability, and strength of bilayer. A number of experimental methods, such as

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shape fluctuation method [9–11], magnetic-field-induced orientation [12], and micropipette aspiration method [13,14] were used to measure the bending modulus of vesicles. However, those methods are suitable for giant liposomes (several micrometer diameter) and need complicated mathematical analysis. As membranes are extremely flexible, measurement of mechanical properties has not been simple or direct; furthermore, well-characterized small force and sub-micrometer detection of membrane deformation are required. Thus, it has been a technology challenge to measure the bending modulus of small vesicles adsorbed on substrate. Besides imaging, force measurements can be used to obtain additional information, such as Derjaguin-LandauVerwey-Overbeek (DLVO) forces, hydration force, steric forces [23], and elastic properties [24–27]. The elastic properties can be deduced from AFM force plots, in which the AFM tip moves up and down over a point on the sample surface, probing the elastic and adhesive properties of materials [24]. In addition to force plots, force mapping giving a complete maps of heterogeneous sample has been used to study the surface properties. The Young’s modulus of the Torpedo synaptic vesicles with apparent diameters of around 90–150 nm were explored by Laney et al. [27] from force mapping by averaging approach and retraction force curves. They discarded the force plots with discontinuities which are important for understanding the structure of liposome. Several papers have described the force curves on different types of bilayer and a jump-in point was observed [28–30]. Franz et al. [28] interpreted the jump-in point as a penetration of the AFM tip through the bilayer. Recently, Teschke and Souza [31] reported individual approaching force curve on liposome. It was found that there are two repulsive regions in approach force distance curve. However, no retraction curve was reported in their work. In our work, small unilamellar egg yolk phosphatidylcholine (EggPC) vesicles (<60 nm made by the sonication method) adsorbed on a mica substrate were studied in solution by AFM tapping and contact mode. First, we attempted to gain a clear understanding of various factors that may affect the stability and topography change of small unilamellar vesicles upon adsorption. Second, we aimed to identify the structure and integrity of vesicles and to develop a method to measure the stability of the vesicles based on AFM approach and retraction force curve. Furthermore, the Young’s modulus and bending modulus measured based on approach force distance plot are reported.

2. Experimental section 2.1. Materials EggPC with 99% purity was purchased from Sigma (St. Louis, MO). Sodium chloride (ACS grade), acetone (HPLC grade), methanol (HPLC grade), and chloroform (99.9%)

were purchased from Fisher Chemicals (Fair Lawn, NJ). Ethyl alcohol (ACS grade) was purchased from Pharmco (Brookfield, CT). Deionized water with resistivity 18 MO-cm was obtained from a Barnstead Nanopure water purification system (Debuque, IA). Grade 3 ruby muscovite mica (New York, NY) was used. Mica consists of negatively charged layers bound together by positively charged interlayer potassium cations (K+ ). In aqueous solutions, a mica cleavage surface becomes negatively charged due to the dissociation of K+ ions. Mica was chosen because it is molecularly smooth and hydrophilic [32]. 2.2. Preparation of EggPC vesicles We followed a well-established recipe to prepare vesicle solutions [4]. Multilamellar vesicle solution was obtained by dissolving appropriate amounts of EggPC lipids in chloroform/methanol (2:1 v/v) and evaporating the solvent with nitrogen. After drying in a desiccator connected to a rotary vacuum pump for 30 min, the lipids were re-suspended by stirring them in an aqueous buffer solution (20 mM NaCl) at a concentration of 0.5 mg/ml. Sonicated unilamellar vesicles (SUVs) were produced from the MLV suspension by sonication to clarity (about 1 h) in a Branson 2200 bath sonicator (Danbury, CT). The suspension was kept in an ice bath during the sonication process. Sonicated samples were centrifuged for 1 h at 17,443 × g to remove large lipid fragments by Sorvall OTD70B ultraspeed centrifuge (Wilmington, DE). SPBs were prepared via the extrusion method. Multilamellar EggPC solution was extruded through a polycarbonate membrane with an average pore size of 200 nm using a LiposoFastTM extruder from Avestin Inc. (Ottawa, ON). The extrusion method produced larger vesicles that ruptured into bilayers upon adsorption on the substrate [16]. 2.3. AFM characterization AFM imaging and force measurement were conducted using a Nanoscope IIIa atomic force microscope from Digital Instruments (Santa Barbara, CA) equipped with an E scanner with a maximum scan area of 16 ␮m2 . The scanner was calibrated following the standard procedures provided by Digital Instruments. The fluid cell (Digital Instruments) was washed with deionized water, ethanol, and deionized water before each experiment. Freshly cleaved mica was mounted onto a stainless steel disk using a sticky tab (Latham, NY). After the substrate was brought to close to the AFM tip, the freshly prepared EggPC vesicle solution was injected through silicone rubber tubing into the fluid cell, sealed by a silicone rubber o-ring. The EggPC vesicle solution was allowed to incubate on mica at room temperature for 1 h. Excess vesicles were removed by flushing the fluid cell with buffer solution followed by pure water, and the images were obtained in pure water. The microscope was allowed to thermally equilibrate for 30 min before imaging. Scanning rates

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between 1 and 5 Hz were used. Room temperature was maintained at 22 ± 1 ◦ C. Images were recorded in contact mode or tapping mode using standard silicon nitride (Si3 N4 ) integral tips (NP type) (Digital Instruments) mounted on cantilevers with a manufacturer-specified spring constant of 0.22 N/m, length of 120 ␮m, width of 15 ␮m, and a nominal tip radius of curvature between 20 and 40 nm. The radius of the tip (33.20 ± 6.62 nm) was calibrated by imaging the TGT01 gratings (Mikromasch inc., Portland, Oregon) [33]. The spring constant of the cantilever was calibrated using the deflection method against a reference cantilever (Park Scientific Instruments, CA) of known spring constant (0.157 N/m) [34]. The calibrated spring constant (0.17 ± 0.05 N/m) was used in all force curves calculations. Force curves were obtained in contact mode only. Multiple force curves were obtained on the mica surface rinsed in pure water and on the phospholipid bilayer. Multiple force curves were also obtained around and on the vesicle to ensure that the force curve represented the interaction between the tip and vesicles. The typical time for a complete cycle was ∼1 s. The force calibration plot is converted to a force versus distance plot by defining the point of zero force and the point of zero separation [15]. Zero force is determined by identifying the region at a large separation, where the deflection is constant. Zero separation is determined from the constant compliance region at high force where deflection is linear with the expansion of the piezoelectric crystal. Both the approaching and retracting force curves are reported unless otherwise specified. 2.4. Calculating elastic properties We fit our approaching force distance data to the Hertz contact model assuming spherical shape for both tip and vesicle. The indentation from the difference between the cantilever distance z − z0 and cantilever deflection d − d0 is described as Eq. (1) [27]. |z − z0 | − (d − d0 ) 1/3
k2 (Rtip + Rves )(1 − υves 2 )2 (d − d0 )2/3 = 0.825 Eves 2 Rtip Rves (1) where z is the Z position, z0 is contact point, d is deflection of tip, d0 is noncontact deflection, υ is Poisson’s ratio, E is Young’s modulus, R is radius, subscript tip and ves mean AFM tip and vesicles, and k is spring constant of tip, respectively. The Poisson’s ratio for vesicles is taken to be 0.5 [27]. Bending modulus kc is deduced from Young’s modulus based on Eq. (2) [35] kc =

Eh3 12(1 − υ2 )

where h is the bilayer thickness.

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2.5. Light scattering The size distribution of vesicles in solution was studied by light scattering (Nicomp 370 Autodilute Submicron Particle Sizer, Pacific Scientific). Polystyrene (diameter = 32 ± 1.3 nm, NanosphereTM size standards, Duke Scientific, CA) was used as the calibration standard. The light scattering cell was from VWR Scientific (West Chester, PA). A He–Ne laser (λ = 632.8 nm) was used. Data were taken at 23 ◦ C. 3. Results and discussion 3.1. Images in tapping mode versus contact mode Images of EggPC vesicles were obtained in both contact mode and tapping mode. Tapping mode is known to reduce frictional and adhesive forces that often interfere with imaging of soft samples. Fig. 1 shows the AFM images (amplitude and deflection images for tapping and contact mode, respectively) and the cross-sectional profiles from the height images of the adsorbed EggPC vesicles taken within the first 15 min after engagement. The images consisted of spherical particles on a flat background. The diameter of the particles was measured to be 48.6±11.4 nm and 69.3±12.8 nm based on tapping mode and contact mode image sectional analysis, respectively. The diameter corresponded to the width of the peak at the base in height images. The average vesicle diameter in solution was determined to be 37.0 ± 7.9 nm by dynamic light scattering. The latter method determined the free vesicles’ diameters while the AFM measured the vesicles after adsorption on the mica. The soft vesicles flatten on the substrate, resulting in larger lateral vesicles size [16,19]. In comparison with tapping mode, the vesicle diameter by contact mode is larger mainly due to tip convolution [36]. The height values by cross-sectional analysis of height images were measured to be 13.9 ± 2.2 nm in tapping mode and 3.9 ± 0.4 nm in contact mode, which are consistent with results from other studies [16,17]. The height value from the tapping mode is roughly about 40–50% of the actual vesicle diameter (30–40 nm). Causes for the unreasonably low height values from contact mode are postulated as follows: (1) the bound vesicle may move during contact mode imaging and (2) the vesicles are compressed by the scanning tip [16,18]. Unlike the contact mode, where the tip is dragged across the surface, the tapping mode causes less movement of vesicles by intermittently contacting the surface. Our experience suggests that both the contact and tapping modes were able to capture intact vesicles, with the tapping mode showing slightly better resolution [20]. Furthermore, a smaller image force can be more easily maintained in tapping mode. 3.2. Topography change of vesicles

(2) As reported in other studies, small SUVs used in our experiment should remain intact on mica [16]. Our AFM

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Fig. 1. AFM images and cross-section profiles of EggPC vesicles on mica substrate by tapping mode (a) and contact mode (b). The images were taken within the first 15 min after engagement in pure water and represented typical images from tapping mode and contact mode. The vesicle suspension was incubated on the mica surface for 1 h before flushing using buffer solution. Initial EggPC concentration was 0.5 mg/ml. The convex particles of size 48.6 ± 11.4 nm are EggPC vesicles, and the blank surface is mica substrate. The image size is 1 × 1 ␮m; and; (a) amplitude image; Z-scale is 20 nm, adsorbed convex vesicles (S), (b) deflection image; Z-scale is 5 nm, flat vesicles (F) caused by frictional or lateral forces in contact mode. The profiles were from the height images. Height value for the EggPC vesicles by tapping mode is 13–15 nm, and the value from contact mode is relatively low, ∼4 nm.

imaging and force measurement results largely supported the findings of enclosed, unruptured vesicles. However, the exact shape of the enclosed vesicles can vary as a function of image force in tapping mode.

With increasing AFM tip image force, i.e. a decreasing set point in tapping mode, EggPC vesicles exhibited three distinctive topographies: convex-shaped vesicles shown in Fig. 2(a), planar vesicles shown in Fig. 2(b),

Fig. 2. Different topographical images and profiles of EggPC vesicles obtained by tapping mode. The images were height representative images of the convex (a), planar (b), and amplitude image of concave (c) of EggPC vesicles. The images were captured in pure water and with initial EggPC concentration of 0.5 mg/ml. The image size is 0.75 × 0.75 ␮m, and Z-scale is 20 nm; (a) convex vesicles. Topographic profiles across two convex vesicles (single dashed line). The height of the vesicles is around 15 nm; (b) planar shape (P). Topographic profile across the two planar vesicles showed that the center of the disk is flat (c) concave vesicles. The profile showed that the center part had been compressed.

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Fig. 3. AFM image of EggPC vesicles on mica surface. (a) Amplitude image size: 3.0 × 3.0 ␮m; Z-scale: 20 nm; the lower boxed area had been scanned for 40 min; the upper area was never scanned by the tip. The scanned area consisted of concave vesicles and the new area consists of convex vesicles. (b) Magnified image of the new area, indicating convex particles. (c) Magnified image from the repeated scanning area, showing compressed vesicles.

and concave-shaped vesicles shown in Fig. 2(c). The topographical changes were reversible with image force. The convex-shaped vesicles were characterized by a dome-like structure and conical relief shape [17,37]. The planar vesicles were flat across the top. The planar vesicles are similar to the saucer-like or pancake-like vesicles reported previously [16,17,38]. The concave vesicles were characterized by a depressed central portion and a higher periphery. The diameter of the vesicles was found to be 73.7 ± 11.5 nm for planar shape and 89.8 ± 6.6 nm for concave shape. There were several possible explanations for the formation of planar vesicles: (1) the vesicle wets or partially fuses with a bilayer [17]; (2) a multilamellar vesicle adsorbs and spreads on the substrate [17,38]; and (3) the vesicle ruptures into bilayer fragments [16]. All these scenarios are not likely in this study because of the small size of the vesicles and the fact that the shapes are reversible by manipulating the image force. Moreover, to the best of our knowledge, AFM image of concave vesicles has not been reported. In our experiments, there are two factors that may influence the topography change of the vesicle: substrate interaction and tip effect. Substrate interaction determines the stability of the vesicles on the mica surface. A strong adhesion force between the vesicles and the substrate may eventually cause the deformation of the vesicles. Based on theoretical calculations, the favorable energy gained by the vesicle upon adsorption was size-dependent [6,7]. In our experiments, the unbinding of the vesicles is driven by the energetic mechanism because of the vesicle size is below the critical radius Rc (200 nm) [6]. Thus, the substrate interaction would not contribute significantly to the shape transition of the vesicles. This is also consistent with our experimental observations. To investigate the significance of this effect further, we designed the following experiments: when we first observed the convex vesicles, we stopped scanning for 1–2 h. Then, the surface was imaged again, only convex vesicles were observed. However, with continuous scanning, concave vesicles appeared. Another illustration of the scanning effect is

that when we observed the concave shape in a certain area, the scan area was then increased [Fig. 3(a)]. It was noticed that convex vesicles were observed [Fig. 3(b)] outside the original scan area [Fig. 3(c)]. Based on these results, it can be concluded that the primary cause of the topography change of the vesicles is not substrate interaction, and that small vesicles are relatively stable on mica surface [16]. The second possible factor is tip effect. A schematic model to illustrate the interaction between the tip and the vesicle is shown in Fig. 4. A convex shape is obtained at the lowest indentation region; a planar shape is obtained at an intermediate indentation, and a concave shape is obtained at the highest indentation. For the concave shape, when the tip moves across the vesicle with an applied force held more or less constant, the amount of indentation is largest at the center of the vesicle. Several factors may contribute to maximum depression at the center. (1) It is known that the mechanical response of thin films couples to the substrate, which results in an increase (50%) of the apparent modulus [39]. The coupling increases with decreasing film thickness. Therefore, the tip indents more in the central region of a sessile droplet with higher thickness. (2) A high volume percentage of material is bound by the surface near the edge of a droplet and spreads less upon compression of the tip. Contrarily, at the center of the vesicle, less resistance exists against squeezing of the trapped liquid portion. The tapping mode is less likely to damage the sample than the contact mode because it eliminates lateral forces (friction or drag) between the sample and the tip. However, the vertical forces in tapping mode are large enough to deform the surface of soft or elastic materials [40]. Therefore, the concave images represent a mixture of topographic and elastic properties of the vesicle surface caused by tip-induced vertical artifacts on vesicles. It may also be helpful to know that the AFM tip can cause deformation of vesicles if the same area is scanned repeatedly. Fig. 3 shows that flattened and concave vesicles dominated after prolonged AFM scanning. Regions spared repeated scanning, e.g. at the beginning of an experiment or areas outside a repeatedly scanned area,

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Fig. 4. Schematic of the tip effect on vesicle topography. (a) Convex vesicle; lowest compression occurs on the vesicle and the convex vesicle is imaged. A vesicle adsorbed on the mica will be somewhat flattened due to adhesion force, so that the height of the vesicle is less than the real height of the vesicle in suspension; (b) planar vesicle; the tip slightly contacts the vesicle and the loading force will further flatten it; (c) concave vesicle; the tip contacts the vesicle, the middle part of the vesicle has been compressed, and the tip senses the concave vesicle.

exhibited largely convex vesicles. It clearly indicated that repeated AFM scanning not only removed some vesicles but also caused deformation of the vesicles. 3.3. Force measurement on the EggPC vesicles A force curve taken on bare mica rinsed in pure water before adding EggPC solution is shown in Fig. 5(a). In the approach stage, it shows a weak and long-range repulsion with a maximum of 0.11 ± 0.02 nN at a separation of 2.55 ± 0.26 nm, followed by a strong attraction with a minimum of −0.10 ± 0.04 nN just before contact. The discontinuity in the force curve is called the jump-in point. Fig. 5(a) agrees with previous measurements [41]. In order to distinguish the characteristic force curves between vesicles and a bilayer, we prepared EggPC bilayer film using extrusion method [16]. Fig. 5(b) and (c) are the force curve and image on the EggPC bilayer patches, respectively. Fig. 5(b) showed a strong repulsion starting at 6.58 ± 0.37 nm, and a jump-in point at 4.57 ± 0.27 nm with a maximum force of 1.75 ± 0.27 nN. We refer to the load at which the jump-in occurs as the breakthrough force. The jump-in (4.57 ± 0.27 nm) corresponds to the removal of the bilayer fragment between the tip and substrate, most probably by a lateral push-out mechanism [28]. The maximum steric barrier was reported to be proportional to the surface excess of the film, and can be used to compare the packing density of films with similar adsorption mechanisms [42]. The onset of repulsion (6.58±0.37 nm) coincided with the thickness (6.27 ± 0.57 nm) measured by AFM images sectional analysis [Fig. 5(c)]. The value (6.27 ± 0.57 nm) from AFM images sectional analysis is larger than the distance between the EggPC hydrophilic headgroup (∼4 nm). The discrepancy (∼2 nm) is probably due to the water layer thickness between the phospholipid bilayer and the mica

substrate [16,43]. For the force curve, the onset of repulsion (∼6.58 nm) corresponds to the relatively unperturbed hydrated thickness of the EggPC bilayer film sandwiched between the tip and substrate, 6.3 nm, as measured by X-ray scattering [44]. It is reported that the repulsive force region (from 6.58 ± 0.37 to 4.57 ± 0.27 nm) is a combination of hydration/steric forces and mechanical deformation [29]. According to our experience, the water layer thickness (∼1–2 nm) information is also included in the region (6.58–4.57 nm) [16,43] although the accurate value is still hard to get. The force curves (not shown) measured on the flat background in Fig. 1 were similar to Fig. 5(a). Therefore, it was concluded that the flat background is mica and that there was no significant bilayer formation in our sonicated vesicle samples which is also consistent with the work done by Reviakine and Brisson [16]. Fig. 6(a) represents a typical force curve captured during force measurements on vesicles. While the exact location of the tip with respect to the vesicle is difficult to assess during these measurements, we took the following precautions. When a small scan area with a vesicle in the center of the image is observed, the force curves are captured. The force curves in the immediate areas are also obtained to show the difference between mica substrate and vesicles. The onset force distance, 32.19 ± 4.18 nm (Point 1) measured in Fig. 6(a), indicates that the tip was at or near the center of the vesicle. The onset values measured on vesicles fell between 30 and 40 nm, consistent with the vesicles’ sizes. At a force of 0.72 ± 0.09 nN (Point 2), a discontinuity with a gap spanning 4.79 ± 0.38 nm was observed. After that the force continued to increase till another jump-in at 5.41 ± 0.61 nm (Point 4). During retraction, one or two abrupt changes were observed in the retracting force curves (in Fig. 6(a) only one is shown). The jump-in and -out gaps were consistently

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Fig. 5. Approaching force curves on mica surface in pure water (a) and EggPC bilayer patches adsorbed from solution (b). The data was taken in pure water. There exists a weak and long-range repulsion with a maximum of 0.11 ± 0.02 nN at 2.55 ± 0.26 nm, followed by a strong attraction near the surface with a minimum of −0.10 ± 0.04 nN before contact. The small repulsive force and the strong attraction force are due to the van der Waals force. (b) Shows that a repulsive force starts at 6.58 ± 0.37 nm and the jump-in distance near 4.57 ± 0.27 nm on the EggPC bilayer patch, (c) is the AFM amplitude image and section analysis of EggPC bilayer patches formed by the extrusion method. The thickness of the EggPC bilayer patches is 6.27 ± 0.57 nm by section analysis, which agrees with 6.58 ± 0.37 nm from force curves.

measured at around 4 nm, that is, in the range of bilayer thickness [45]. This suggests that these jump-in and -out gaps correspond to the sudden opening and closing of portions of the vesicle bilayer during approach and retraction force measurements. This finding is consistent with the jump-in point observed on bilayer [28]. The discontinuities on approach force curve on liposomes were also reported [29]. It should be noted that sometimes force curve with one jump-in point around 9.78 ± 1.14 nm was observed (not shown). The force curve should not be the result of isolated bilayer patches because the repulsive onset distance (∼30 nm) is significant higher than that of a typical bilayer (∼6 nm). One possibility is due to deformed vesicles (as a result of repeated scanning), therefore, with double bilayer thickness jump-in (9.78 ± 1.14 nm). The exact nature of the one jump-in is being further investigated. The stepwise mechanical deformation events constructed based on the force curves are illustrated in Fig. 6(b). When the tip is brought into contact with the vesicle, further advancement of the tip is deterred by the steric repulsion and elastic deformation of the vesicle [Fig. 6b-1]. The slope of the curve should correlate to the local bending modulus of the vesicle. At a critical force (0.72 ± 0.09 nN), the tip breaks through the upper portion of the bilayer in direct contact with the tip, probably through a push-out mechanism

[Fig. 6b-2]. Further steady increase in repulsion after the initial jump-in event indicates that the tip feels the other steric force and hydration force from the vesicle bilayer attached to the mica [Fig. 6b-3]. After reaching the maximum force, a second jump-in occurs, corresponding to the abrupt disruption of the vesicle bilayer that is attached to the substrate [Fig. 6b-4]. After the second jump-in near the substrate, the force increases rapidly, since the tip now presses against the hard surface. During retraction, when the force is gradually released, a jump-out near the surface represents resealing of the lower bilayer. The tip remains in contact with the vesicle until the final jump-out event occurring at a distance larger than the size of the vesicle. Rearrangement of lipid molecules may occur near the contact zone, giving rise to the adhesive force between the AFM tip and the vesicle. The fact that after the tip breaks the upper bilayer the vesicle maintains its elasticity suggesting that the tip adheres to the edge of the bilayer. The same adhesion force prolongs the detachment process of the tip from the vesicle upon retraction [Fig. 6b-5]. The schematic model illustrating the interaction between the tip and the vesicle probes the bilayer structure (two jump-in points) and the cohesive energy (breakthrough force) of the vesicles. Teschke and Souza [31] interpreted the interaction between tip and dried liposome during approaching process in a similar fashion. The difference is that

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2

Approach

(a)

Retraction

4 2

Force (nN)

1

0 -5

1

3 5

15

25

35

45

65

75

5

-1

-2

55

Distance (nm) (b)

2

1

5 3 4

Fig. 6. Typical force curves on the EggPC vesicle (a) and schematic procedures simulating the tip-vesicle interaction (b). When the tip is approaching the vesicle, the tip feels confinement repulsion around 32.19 ± 4.18 nm, which corresponds to the height of vesicle (Cartoon 1). At a force of 0.72 ± 0.09 nN, a jump-in (Cartoon 2) with gap distance 4.79 ± 0.38 nm occurs because the tip breaks through a portion of the upper vesicles bilayer. Another repulsion occurs (Cartoon 3) with a second jump-in at 5.41 ± 0.61 nm (Cartoon 4) while the tip is continuing down. The two long-range repulsive forces are from the confinement and compression of the vesicles, and the two jump-in points correspond to the two processes of the tip penetrating the upper and lower bilayers of the EggPC vesicles. During retraction, one abrupt change near 4 nm was observed. When the force is gradually retracted, the final jump-out point occurs at a distance larger than the size of the vesicle, due to the elongation effect on the vesicle by the adhesive force between the AFM tip and the vesicle. (Cartoon 5).

they suggested that bilayers are formed on the surface as a result of the rupture of the vesicles, and no retraction curve is provided in their paper. One the other hand, our model reveals the elastic reversible morphological changes of vesicles as evidenced by a series of reproducible force curves at the same spot. Thus, the elastic properties (Young’s and bending modulus) of adsorbed vesicles can be deduced. 3.4. Data analysis according to Hertz model Laney et al. [27] calculated elastic properties from averaged data of approach and retraction force curves and they

discarded force plots with discontinuities. In our work, individual force curve was obtained. The slope of the approaching force curve reflects the mechanical properties of the vesicles under compression. It is important to use the approach part of the force curve to calculate the indentation because significant adhesive forces can affect the measurement of indentation [25]. The first repulsive force region in the approach process was used to calculate Young’s modulus based on Hertz model [24,27] and data for indentation less than 10 nm was used for model fitting. The power equation δ = AFb (δ is an indentation (compression) on the vesicle, F is load force) is used to fit the

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Indentation (nm)

10 8 6 4 Experiment data

2

Hertz model

0 0

0.4

0.8 1.2 Load force (nN)

1.6

Fig. 7. Force curve data fit with Hertz model. The squares are experimental data. Experimental data can be described by power equation δ = 8.2752F 0.6433 . The solid line is the Hertz model δ = AF2/3 with A = 8.2752. The measured compression (indentation) versus loading force agrees with the Hertz model in the case of the first repulsive force region.

experimental data since the power equation is the basis for the Hertz model. From the power equation, b is found to be 0.6433 and A is equal to 8.2752. The exponent b = 0.6433 is close to 2/3 as described by the Hertz model for spherical contact [27]. Thus, all calculations for determining the mechanical properties of the vesicles surface are based on fitting the force distance curves to the Hertz model for spherical contact. Fig. 7 shows that in the beginning of the compression (indentation), the Hertz model can simulate the experimental findings very well. A deviation from the

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model is observed at high load force (larger indentation) illustrating the limitation of the model. Young’s modulus was calculated using Rtip = 33 nm, Rves = z0 /2 (the onset point of repulsive force is regarded as the size of the vesicles), υ = 0.5, and k = 0.17 N/m. Bending modulus is a characteristic property of the vesicles, which is closely related to the activities of liposomes and the gel-liquid phase transition of liposome’s bilayer membrane. According to solid-state mechanics, Young’s modulus is related to bending modulus as Eves = kc /I where I is the cross-sectional moment. The value I for a three-dimensional, isotropic planar surface is h3 /[12(1 − υ2 )] where h is the thickness of the bilayer and ␷ is Poisson’s ratio [27,35]. The bending modulus kc was calculated using Eq. (2) with h = 4.57 × 10−9 m, and υ = 0.5. The Young’s modulus and the bending modulus are found to be 1.97 ± 0.75 MPa and (0.21 ± 0.08) × 10−19 J, respectively (see Tables 1 and 2). The Young’s modulus of the EggPC vesicle from force plots is one order of magnitude smaller than the value (∼107 MPa) reported [46]. The discrepancy is probably due to different methods used and different environment the vesicles are in. Hantz et al. [46] gave the bulk average Young’s modulus value in solution while our data was obtained on the individual vesicle adsorbed on hard substrate. Our data is comparable with the value reported for adsorbed synaptic vesicles (0.2–1.3 MPa) on mica [27] from force mapping method. However, the accurate value of Young’s modulus derived from force curve is affected by several factors such as accurate tip radius (Rtip ), tip spring constant (k), contact point (z0 ) and liquid environment [27].

Table 1 Young’s modulus of biological samples Material

Young’s modulus E (MPa)

Method

Remarks

Reference

Synaptic vesicles DMPC vesicles DOPC vesicles EggPC vesicles

0.2–1.3 15

Force mapping Osmotic swelling

Size: 90–150 nm adsorbed on mica Size: 160–180 nm in solution

[27] [46]

Force plot

Size: <60 nm adsorbed on mica

This work

1.97 ± 0.75

DMPC: dimyristoylphosphatidylcholine; DOPC: dioleoylphosphatidylcholine. Table 2 Comparison of bending modulus of egg yolk phosphatidylcholine Method

Size/shape

Bending modulus kc (× 10−19 J)

T (◦ C)

Reference

Phase contrast microscopy

Long unilamellar tubular vesicle (11 ␮m < L < 34 ␮m, 17 < L/r < 83) Cylindrical vesicle (>10 ␮m) Quasi-spherical vesicle (>10 ␮m) Spherical (>10 ␮m) Cylindrical rods (5–30 ␮m diameter, <200 ␮m long)

2.3 ± 0.3

22.0

[9]

1–2 1–2 0.4–0.5 0.4

25

[10] [11] [47,48] [12]

Magnetic-field-induced orientation AC electric field

AFM force curve

Spherical vesicle (diameter >20 ␮m) Spherical vesicle (∼15–70 ␮m diameter) Spherical vesicle (diameter <60 nm) on mica substrate

0.247 0.66 ± 0.06 0.45 ± 0.05 0.21 ± 0.08

25

[49] [50] 22 ± 1

This work

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X. Liang et al. / Colloids and Surfaces B: Biointerfaces 34 (2004) 41–51

The bending modulus calculated ((0.21±0.08) × 10−19 J) from force plots is in the same magnitude range (10−20 to 10−19 J) of literature reported value for the same EggPC lipid [9–12,47–50] (see Table 2) regardless of different geometry and size of the samples. Variations in bending modulus measured have been reported and the reason for the discrepancy still needs further analysis [49–51]. Several possibilities are proposed to explain the variations of bending modulus observed: (1) different experimental method: Niggemann et al. [51] found that the values of the rigidity for the same lipid differed significantly between methods. Our data was derived from Young’s modulus based on AFM force curves and others use phase contrast microscopy [9–11,47,48], magnetic-field [12], and electric field method [49,50]; (2) bending modulus (Eq. (2)) is based on three-dimensional, isotropic planar surface assumptions, while the vesicles on substrate is not prefect isotropic planar surface; (3) our data is based on the result on the adsorbed small size vesicles while other methods deal with giant liposomes (>10 ␮m) vesicles. Although there is some variation for the bending modulus value from different methods, our data from AFM force curve can measure the mechanical properties of adsorbed vesicles with much smaller size. The values obtained are comparable with other techniques, which deal with much larger vesicles. In a subsequent paper, we will use the force plot to quantify the micromechanical property changes of the cholesterol-modified vesicles.

4. Conclusion In this study, images of small unilamellar EggPC vesicles are obtained by tapping and contact mode in the fluid. Different topographies (convex, planar, and concave) of the EggPC vesicles observed on mica substrate can be attributed to the interaction between the AFM tip and the vesicles. It is found that the tapping mode can cause vertical force-induced soft vesicle deformation. The force curve on the vesicle provides evidence for the stepwise opening and closing of the upper and lower portions of the vesicle bilayer during approach and retraction of the AFM tip. The force curve can be used to pinpoint the steps, such as elastic compression and stretching, breakthrough and resealing of upper and lower bilayers, in the deformation of a single vesicle. Based on force curve analysis, the Young’s modulus (E) and the bending modulus (kc ) for pure EggPC vesicles are found to be (1.97 ± 0.75) × 106 Pa and (0.21 ± 0.08) × 10−19 J. These findings compare favorably with other techniques to measure the micromechanical properties of vesicles. This study demonstrated that AFM force measurement provides a unique approach to probing not only the topography and structure, but also the micromechanical behavior of adsorbed small vesicles.

Acknowledgements We wish to thank the Petroleum Research Fund for grants 36149-ACS and 33036-ACS, administered by the American Chemical Society, and the Institute for Manufacture Research at Wayne State University for financial support. We also wish to thank J. S. Jourdan at BASF in Wyandotte, Michigan for help with the light scattering measurements.

References [1] R.R.C. New, in: R.R.C. New (Ed.), Liposomes: A Practical Approach, Oxford, New York, 1990. [2] D.D. Lasic, Tibtech 16 (1998) 307. [3] Y. Barenholz, Curr. Opin. Colloid Interface Sci. 6 (2001) 66. [4] C.-H. Huang, Biochemistry 8 (1969) 344. [5] M.J. Hope, M.B. Bally, G. Webb, P.R. Cullis, Biochim. Biophys. Acta 821 (1985) 55. [6] U. Seifert, R. Lipowsky, Phys. Rev. A 42 (1990) 4768. [7] U. Seifert, Adv. Phys. 46 (1997) 13. [8] V.P. Zhdanov, B. Kasemo, Langmuir 17 (2001) 3518. [9] R.M. Servuss, W. Harbich, W. Helfrich, Biochim. Biophys. Acta 436 (1976) 900. [10] J.T. Schneider, M.B. Jenkins, W.W. Webb, Biophys. J. 45 (1984) 891. [11] J.T. Schneider, M.B. Jenkins, W.W. Webb, J. Phys. France 45 (1984) 1457. [12] I. Sakurai, Y. Kawamura, Biochim. Biophys. Acta 735 (1983) 189. [13] E. Evans, W. Rawicz, Phys. Rev. Lett. 64 (1990) 2094. [14] R.E. Waugh, J. Song, S. Svetina, B. Žekš, Biophys. J. 61 (1992) 982. [15] C.B. Prater, P.G. Maivald, K.J. Kjoller, M.G. Heaton, Probing Nano-scale Forces with the Atomic Force Microscope, Application note no. 8 from Digital Instruments, 1995, see also references therein. [16] I. Reviakine, A. Brisson, Langmuir 16 (2000) 1608. [17] S. Kumar, J.H. Hoh, Langmuir 16 (2000) 9936. [18] T. Shibata-seki, J. Masai, T. Tagawa, T. Sorin, S. Kondo, Thin Solid Films 273 (1996) 297. [19] B. Pignataro, C. Steinem, H.-J. Galla, H. Fuchs, A. Janshoff, Biophys. J. 78 (2000) 487. [20] N.H. Thomson, I. Collin, M.C. Davies, K. Palin, D. Parkins, C.J. Roberts, S.J.B. Tendler, P.M. Williams, Langmuir 16 (2000) 4813. [21] J. Jass, T. Tjärnhage, G. Puu, Biophys. J. 79 (2000) 3153. [22] A.S. Muresan, K.Y.C. Lee, J. Phys. Chem. B 105 (2001) 852. [23] H.-J. Butt, Biophys. J. 60 (1991) 1438. [24] A.L. Weisenhorn, M. Khorsandi, S. Kasas, V. Gotzos, H.-J. Butt, Nanotechnology 4 (1993) 106. [25] A. Vinckier, G. Semenza, FEBS Lett. 430 (1998) 12. [26] I. Lee, R.E. Marchant, Colloid Surf. B 19 (2000) 357. [27] D.E. Laney, R.A. Garcia, S.M. Parsons, H.G. Hansma, Biophys. J. 72 (1997) 806. [28] V. Franz, S. Loi, H. Muller, E. Bamberg, H.-J. Butt, Colloid Surf. B 28 (2002) 23. [29] Y.F. Defrene, T. Boland, J.W. Schneider, W.R. Barger, G.U. Lee, Faraday Discuss. 111 (1999) 79. [30] J. Schneider, Y.F. Dufrene, Jr., Barger, G.U. Lee, Biophys. J. 79 (2000) 1107. [31] O. Teschke, E.F. Souza, Langmuir 18 (2002) 6513. [32] Z.V. Leonenko, A. Carnini, D.T. Cramb, Biochim. Biophys. Acta 1509 (2000) 131. [33] J.S. Villarrubia, J. Res. Natl. Inst. Stand. Technol. 102 (1997) 425. [34] M. Tortonese, M. Kirk, SPIE 3009 (1997) 53. [35] E.A. Evans, Biophys. J. 14 (1974) 923.

X. Liang et al. / Colloids and Surfaces B: Biointerfaces 34 (2004) 41–51 [36] A. Doron, E. Joselevich, A. Schlittner, I. Willner, Thin Solid Films 340 (1999) 183. [37] H. Egawa, K. Furusawa, Langmuir 15 (1999) 1660. [38] S. Singh, D.J. Keller, Biophys. J. 60 (1991) 1401. [39] J. Domke, M. Radmacher, Langmuir 14 (1998) 3320. [40] Manual of “Scanning Probe Microscope”, from Park Scientific Instruments, Sunnyvale, CA P5. [41] O. Teschke, E.F. de Souza, G. Cetto, Langmuir 15 (1999) 4935. [42] K. Eskilsson, B.W. Ninham, F. Tiberg, V.V. Yaminsky, Langmuir 15 (1999) 3242. [43] E. Sackmann, Science 271 (1996) 43. [44] T.J. McIntosh, A.D. Magid, S.A. Simon, Biochemistry 26 (1987) 7325.

51

[45] J. Marra, J. Israelachvili, Biochemistry 24 (1985) 4608. [46] E. Hantz, A. Cao, J. Escaig, E. Taillandier, Biochim. Biophys. Acta 862 (1986) 379. [47] J.F. Faucon, M.D. Mitov, P. Méléard, I. Bivas, P. Bothorel, J. Phys. France 50 (1989) 2389. [48] H.P. Engelhardt, H. Duwe, E. Sackmann, J. Phys. Lett. France 46 (1985) L395. [49] M. Kummrow, W. Helfrich, Phys. Rev. 44 (1991) 8356. [50] M.I. Angelova, S. Soleau, P. Meleard, J.F. Faucon, P. Bothorel, Progr. Colloid Polym. Sci. 89 (1992) 127. [51] G. Niggemann, M. Kummrow, W. Helfrich, J. Phys. II France 5 (1995) 413.