Chapter 5 SPM applied to lipid LB monolayers

In this chapter an introduction to Langmuir-Blodgett (LB) films will be given, after which the application of SPM to the characterization of these films will be demonstrated. Apart from their biological relevance (as model systems for biological membranes) these films are a challenge to any form of microscopy since they are transparent and very thin (about 2 nm). In the sixth and last chapter a new form of atomic force microscopy (AFM) will be applied to these films.

5.1 Biological membranes

In 1925 Gorter and Grendel came with their hypothesis that all living cells are surrounded by a lipid bilayer.1 They concluded this from an experiment that was met with a lot of criticism. In 1966 it was finally discovered that in fact they had made two mistakes that miraculously canceled each other out and that their conclusion was right. The only thing that had to be added to their model was the fact that in the natural membrane a variety of proteins (receptors) are embedded. This currently accepted model is known as the fluid mosaic model (see Fig. 5.1)2,3.

Fig. 5.1 Fluid mosaic membrane model: the natural membrane is a lipid bilayer, with embedded proteins.

Lipid bilayers consist of amphiphilic molecules which in their turn consist of a hydrophobic ('water-fearing') part, and a hydrophilic ('water-loving') headgroup. dimiristoylphosphatidylethanolamine (DMPE), is an example of a amphiphilic membrane component, with a hydrophilic headgroup and two hydrophobic alkylchains (see also Section 5.2.1 and Fig. 5.5).

Many essential functions of living cells are performed by events associated with their membranes. The amount of protein in biological membranes varies between 30 and 70% depending on the function of the cell. The basic functions of biological membranes can be divided in three groups:4

(i) Organization: energetically most favorable spatial arrangements of chemical reaction sites and cell components.

(ii) Interaction: transport of substances.

(iii) Information: cell recognition and adhesion, immune reactions.

Because of various enzymes and other functional proteins attached to or embedded in membranes, they are involved in many metabolic processes. The two important energy conversion processes, photosynthesis in chloroplasts and oxidative phosphorylation in mitochondria, are carried out in membranes.5 To understand these and many other processes, which are essential for living organisms, it is necessary to obtain knowledge of molecular structure and function of the membrane, preferably at a very high resolution.

5.1.1 Biomembrane models

The complexity of the biomembrane is such that frequently simpler systems are used as models for physical investigations. They are based on the spontaneous self-organization of the amphiphilic lipid molecules when brought in contact with an aqueous medium. The three most frequently used model systems are: (i) monolayers; (ii) black lipid membranes (BLM); (iii) vesicles or liposomes (see Fig. 5.2).

(a)(b) (c)Air Water

Fig. 5.2 Three of the most frequently used model systems: (a) monolayers at the air-water interface; (b) black lipid membranes (BLM); (c) vesicles or liposomes.

BLM and vesicles consist of lipid bilayers separating two aqueous compartments. They make the measurement of e.g., membrane resistance, capacity and permeability possible. Liposomes are spherical lipid multilayers.

An aspect of the model systems that has considerable relevance in the biological field, is their ability to undergo phase transitions (see Section 5.2.1). It is known that biomembranes tend to adapt to the ambient conditions to maintain themselves in a transitional state.6 In particular, it is thought that separate domains of lipid in different phases exist in real biomembranes. Because of this biological relevance, as well as their intrinsic interest, much attention has been paid to the phase transitions of simple model membrane systems.

A monomolecular layer of lipid molecules spread at the air-water interface is a system in which the transitions have been exhaustively studied.7 The hydrophilic parts of the molecules are then submerged in the water, while the hydrophobic parts remain outside and interact with each other to form a layer.

The monolayer at the air-water interface represents only one half of the naturally occurring bilayer, but allows the packing density of the spread lipid molecules to be varied by changing the available area per molecule. Therefore, monolayers can yield precise information on molecular packing and orientation, although they cannot provide information on membrane conductance, as is provided by BLM studies, or membrane permeability, as provided by liposome work.5 Thus, the monolayer can give precise information on membrane constituents organized in a membrane-like environment, but is less able to provide direct information on membrane functionality.

An advantage of the lipid monolayer as a model is that it can be transferred to a solid substrate, using the Langmuir-Blodgett (LB) technique, allowing it to be studied using a whole range of techniques. The spreading of lipid monolayers, monolayer experiments and the Langmuir-Blodgett technique will be discussed in the next section.

5.2 Monomolecular layers

In this section monolayer spreading and deposition is treated. The measurement of isotherms is described, and the interpretation of isotherms is discussed.

5.2.1 Monolayer spreading

Monomolecular films on a water surface (named Langmuir films, after Irving Langmuir 1881-1957), can be prepared by spreading: when a lipid solution in a water-immiscible solvent such as chloroform is dropped onto a water surface, the solution spreads rapidly to cover the available area. After the solvent is evaporated, the lipid molecules are left behind at the air-water interface.7

The amphphilic lipid molecules will naturally orient themselves such that the hydrophilic headgroups are immersed in the water surface and the hydrophobic tailgroups remain outside and interact with each other. When a large water surface area per molecule is available, the film molecules move independently on the surface. Such a film is called a 'gaseous' film, in two dimensions. The monolayer can be compressed by decreasing the surface area with a moveable barrier (see Fig. 5.3). The barrier is in contact with the water and the lipid molecules cannot pass the barrier.

barrier subphase

Fig. 5.3 Schematic drawing of a monolayer at an air-water interface. The monolayer can be compressed by moving the barrier to the left.

For a compressed monolayer, the van der Waals forces are very large, the film molecules are not allowed to move about freely, and thus the film resembles a ‘solid’ in a two-dimensional crystalline phase. When the van der Waals forces between the molecules are weaker at larger molecular areas, the result is a ‘liquidlike’ film.


Fig. 5.4 Different phases of a monolayer: (a) gaseous phase; (b) Liquid-Expanded phase; (c) Liquid-Condensed phase; (d) Solid-Condensed phase.

Generally, a distinction is made between films in a liquid-expanded state (LE) and films in a liquid-condensed state (LC). It is not clear what the conformation of molecules in those phases looks like, but generally the conformations are envisaged as shown in Figure 5.4. The problem is that, as far as the conformation is concerned, it is not known where to draw the line between both phases. In the next section it will become clear why this distinction is made. Möbius gave a slightly different interpretation of the phase transitions.8 He divided the molecule under study in three sub-units, with each phase transition corresponding to one sub-unit being lifted from the water surface.

Molecules in the solid phase, have a dense packing of the headgroups or of the alkyl chains, depending on their relative sizes. Further decrease of the surface area results in the collapse of the monolayer by formation of undefined multilayers. The collapse areas for stearic acid and DMPE are determined by their alkyl chains.

(C14) (C14) (C18) NH3 COOH POOOOOOOO +- (a)(b)

Fig. 5.5 Molecular structure of: (a) stearic acid; (b) dimyristoylphosphatidylethanolamine (DMPE).

It is obvious that the role of the interaction of the subphase, i.e. water, with the film-forming substance is very important. For example, the dipolar nature of the molecules (see Fig. 5.5) has the effect of aligning the dipoles at the air-water interface. If there are sufficient ions in the subphase, this electrostatic interaction will be more or less shielded, which will reduce the interaction between the dipoles.

The presence of a molecular film on the surface of the water has a pronounced effect on the physical properties of the air-water interface. In the next section, the influence of a monolayer on the surface pressure and its measurement will be discussed.

5.2.2 Surface pressuremeasurement

The water surface on one side of any imaginary line pulls perpendicular to this line with a force equal and opposite to that exerted by the surface on the other side of the line. The force acting perpendicularly to a line of unit length is called the surface tension \(\gamma\), measured in N/m or mN/m. The surface pressure \(\pi\) of the monolayer is defined as the lowering of the surface tension due to the presence of the monolayer on the water surface:


It can be interpreted as the two-dimensional analog of the three dimensional pressure. The measurement of the surface pressure as a function of the available area per molecule can provide information concerning molecular dimensions and intermolecular interactions.

(a)(b) thwwater lq

Fig. 5.6 Wilhelmy plate with dimensions indicated, immersed to a depth h in water, with a contact angle \(\theta\): (a) front view; (b) side view.

A method frequently used to determine the surface pressure resulting from monolayer compression is the Wilhelmy plate method (see Fig. 5.6). It is a highly accurate method for the measurement of the surface tension. Using the Wilhelmy method, a measurement is made by determining the force due to surface tension, on a plate which is suspended so that it is partially immersed in the water subphase. The forces acting on the plate consist of gravity and surface tension which pull downward (for a hydrophilic plate), and buoyancy due to displaced water which gives an upward force. For a rectangular plate of dimensions l, w and t, and material of density \(\rho_P\), immersed to a depth h in a liquid of density \(\rho_L\) the net downward force is given by:


where \(\gamma\) is the surface tension, \(\theta\) is the contact angle on the solid plate, and g is the gravitational constant. Materials that are generally used for the Wilhelmy plate are clean filter paper or platinum. When completely wetted, the contact angle will almost be zero (\(\theta\approx 0°\)).

5.2.3 Monolayer isotherms

By quasi-statically moving the barrier and simultaneously measuring the surface pressure, the surface pressure can be determined as a function of the area of water available to each molecule. Because the temperature is kept constant, this relation is known as the surface pressure vs. area isotherm, usually abbreviated to 'isotherm', the two dimensional analog of the three dimensional P/V diagram. If the area per molecule is sufficiently large, ideal gas films obey the relationship:

\[\pi A=kT,\tag{5.3}\]

where k is the Boltzmann constant. This relationship is analogous to the three-dimensional gas law (i.e., PV = kT). In general, ideal gas behavior is observed only when the distances between the molecules are very large and the surface pressure is very small (less than 0.1 mN/m).

SCLCLEG Surface Pressure (mN/m) Mean Molecular Area (nm2) 040010 504030 20100 0.40.5

Fig. 5.7 The isotherm of stearic acid. The inset does not represent measured values, but serves as illustration for the behavior at very high available area.

In Figure 5.7 the isotherm for stearic acid is given (taken from Ref. 9). On examining this isotherm, a number of distinct regions become immediately apparent. If the surface area is reduced from an initial high value, with the monolayer in a gaseous state, there is a gradual onset of surface pressure until an approximately horizontal region is reached (inset Fig. 5.7). This is the gaseous-liquid expanded (LE) transition in the stearic acid isotherm. In the horizontal region the hydrophobic tailgroups, which were originally lying almost flat on the water surface, are subsequently being lifted from that surface. The inset in Figure 5.7 does not represent measured values, because the surface pressure in this region is so low that this portion of the isotherm is not resolved by measurements. Analogous to a 3D-gas, the compressibility is defined as:

\[\kappa=-\frac{1}{A}\left\lgroup\frac{\partial A}{\partial \pi}\right\rgroup_T.\tag{5.4}\]

The gaseous-liquid expanded transition of stearic acid is an example of a first order transition, which is characterized by a horizontal slope in the isotherm, with the compressibility approaching infinity.

There follows an abrupt transition to the liquid-condensed (LC) phase at an area per molecule of 0.28 nm2. The different phases are observed as portions of the isotherm with different slopes. In the liquid condensed phase the slope of the isotherm, and thus the compressibility, is approximately constant.


Fig. 5.8 The collapse mechanism by which undefined multilayers are formed.

At a surface area of just over 0.2 nm2 per molecule there is an abrupt increase of slope. Clearly, this is due to a phase change as well and represents a transition to an ordered solid-like arrangement of the molecules, known as the solid condensed (SC) or solid phase. In this phase the molecules are close-packed, and an all-trans (straight) chain configuration can be achieved (see Fig. 5.4). If this second linear portion of the isotherm is extrapolated to zero surface pressure, the intercept gives the area per stearic acid molecule that would be expected for the hypothetical state of an uncompressed close-packed layer. This value of 0.22 nm2 per molecule is close to that occupied by stearic acid molecules in single crystals, which confirms the interpretation of a compact film as a two-dimensional solid. The mean molecular area in close-packed configuration is equal to the cross-sectional area per CH2 chain.10 Further reduction of the surface area results in the collapse of the monolayer (see Fig. 5.8). Because molecular layers are riding on top of each other the compressibility then approaches infinity again.

For a DMPE monolayer a different isotherm is measured (see Fig. 5.9; taken from Ref. 11). For mean molecular areas around 0.5 nm2 the compressibility approaches infinity, indicating a first order phase transition from the LE to the LC phase. In first order phase transitions, phases coexist in separate domains, as will be discussed in Section 5.3.

The LC-SC transition is indicated by an abrupt increase in slope in the already steep portion of the isotherm. Note that the collapse area is approximately twice that of stearic acid because of the second CH2 chain of the DMPE molecule. The characteristics of the isotherms however, do not depend only on the molecules. The subphase is also very important, in particular the pH and the ion strength of the subphase. The influence of the temperature is analogous to that for a 3D-gas, but for monolayers there can be more phase transitions. 0.81.0 01530 4560 SCLCLEG Surface Pressure (mN/m) Mean Molecular Area (nm2)

Fig. 5.9 The DMPE isotherm.

5.2.4 Langmuir-Blodgett deposition on a solid substrate

For some microscopy techniques it is necessary to transfer a monolayer from the water surface to a solid substrate. The technique to do this has become universally known as the Langmuir-Blodgett (LB) technique.12 For LB deposition, a substrate is first lowered through the monolayer so that it dips into the subphase, and then withdrawn. If the substrate used is hydrophilic, then deposition will follow the sequence of events shown in Figure 5.10.

During the lowering of the slide, the water wets the slide's surface and the meniscus turns up, but there is no deposition at this stage. As the slide is withdrawn, the meniscus is wiped over the slide's surface and leaves a monolayer behind. During this process the surface pressure is kept constant, using the surface pressure as a feedback signal for adjusting the barrier position. The value of surface pressure that gives best results depends on the nature of the monolayer but is usually more than 10 mN/m. In the deposited monolayer the hydrophilic groups are turned toward the hydrophilic surface of the slide. Initially there is liquid film between the monolayer and the slide's surface and bonding is completed after the water layer has drained away or evaporated.


Fig. 5.10 Deposition of multilayers by the Langmuir-Blodgett technique: (a) first immersion; (b) first withdrawal; (c) second immersion; (d) second withdrawal.

The rate at which the slide can be withdrawn from the water while a homogeneous film is deposited, is typically 1 mm/s. This rate depends on the rate at which the water film drains from the monolayer-slide interface and on the viscosity of the monolayer. A highly viscous monolayer will be unable to maintain a homogeneous film in the neighborhood of a rapidly moving slide.

To deposit a second layer onto the slide, the slide is again lowered through the monolayer. This time the slide's surface is hydrophobic; the meniscus turns down and the second monolayer is deposited with its hydrophobic tailgroups in contact with the exposed tailgroups on the slide. At the second withdrawal the next monolayer is deposited like the first, onto the hydrophilic headgroups of the last monolayer. If the monolayer material shows poor adhesion to the substrate, then the second immersion in the water can simply lead to the monolayer peeling off the slide and respreading on the water surface. Repeated dipping will then lead to a single deposited monolayer on the slide.

Because during dipping the pressure is kept constant, the barrier will move to compensate for the deposited molecules. By measuring the change in available surface on the water and the surface area of the substrate, the transfer ratio can be determined.

5.3 Domain formation in lipid monolayers

In a previous section it was mentioned that during a phase transition in a monolayer, different phases coexist, organized in separate domains. A problem encountered when describing domain formation is that the description is completely different for different compounds and conditions. In this section only the most essential and general aspects are discussed.

5.3.1 Domain nucleation and growth

In this section we will consider domain nucleation and growth in a film in an LE-LC phase transition. LC domains will be referred to as 'solid' and LE domains as 'fluid' or 'liquid'. In the LC domains the molecules attain a near close-packed configuration. In the LE domains the molecules have more freedom to move, with flexibility increasing along the chain with the distance from the anchoring hydrophilic group.

Consider a lipid monolayer composed of liquid and solid domains, and for simplicity assume that the pure liquid and pure solid phases are incompressible. For a given water surface area, the fractions of the area covered by fluid and solid phases are then fixed. A state of global equilibrium is achieved once the number and shape of the solid domains correspond to the minimum energy state of the film.

On first slowly compressing a monolayer, a certain number of solid domains appears by a nucleation process.13,14 On further compression, these domains usually grow in size but not in number. If the monolayer is compressed rather quickly or if the impurity content is high, a large number of nuclei are formed. If compression speed is decreased the domain size is increased.14,15 In Ref. 16 the onset of the phase transition was detected before solid domains were visible with fluorescence microscopy. Therefore, it was suggested that domain formation results from aggregation of unresolved particles of condensed lipid.

Domain shapes are reversible with respect to growth and decay, even if these shapes are complex:14,17 the same kind of shapes are observed before and after compression and subsequent decompression. The changes are also reversible with respect to the temperature.18

The alignment of polar or charged molecules in the solid domains causes long-range electrostatic forces. Due to the electrostatic repulsion between the solid domains, they are separated by an approximately constant distance. For fusion of domains, a favorable relative orientation of colliding domains is necessary. This can be explained by the fact that the lattices of the crystalline domains must match. In that case the van der Waals forces between the touching lipid chains of both domains exceed the electrostatic repulsion.19

5.3.2 Domain shapes

The shape of domains depends on the monolayer compound and the experimental conditions, and is determined by a thermodynamic equilibrium. More precisely, there is a competition between the long-range electrostatic dipolar repulsion Fel and the line tension \(\lambda\). The line tension represents the tendency to minimize boundary length. The shape of an individual solid domain is determined by the interplay of these forces:13,18,20-22

\[F=\lambda p+F_{el}\tag{5.5}\]

Here p is the perimeter of the solid domain. The electrostatic repulsion favors elongation and narrowing, while the line tension favors short boundaries and thus round domains.

Initially, the domains cover a small fraction of the area of the air-water interface, and are relatively far apart. Therefore, the electrostatic interaction is small, and circular shapes result. As the domains grow in area they come nearer to each other and tend to thin in one dimension because of the long-range dipolar forces. Experiments have confirmed the existence of the electrostatic interaction. Domains were microscopically observed while repelling each other over distances of tens of micrometers and could be moved by applying an electric field.18

In the absence of charged headgroups, the minimum energy domain shape is determined entirely by line tension and is circular for an isotropic solid. In the other extreme, when the charge is large and the line tension is weak, and if the domain does not break into pieces, the minimum energy shape is highly elongated. This effect can be observed when a small amount of cholesterol is added to a charged monolayer. This results in a strongly decreased line tension and the mentioned longer and thinner domains.21

Because the electrostatic repulsion influences the domain formation, domain shapes also depend on the ionic conditions of the water subphase. In Ref. 23 it was shown that an increase in the concentration of monovalent ions increased the pressure corresponding to the LE-LC phase transition of the lipid dilauroylphosphatidic acid (DLPA). Increasing the concentration of divalent ions had the opposite effect. These effects were ascribed to an increase in surface charge density in the former case, and to screening of electrostatic forces in the latter. Thus, it was possible to vary the size and shape of the domains at the air-water interface with the ionic conditions.

Another important factor is whether the packing of the molecules is determined by the chains or by the headgroups. For phospholipids and fatty acids, where crystallization is basically determined by the chain arrangement, the solid domains exhibit rounded boundary lines. If the packing is determined by the headgroups, domains exhibit pointed perimeters (e.g. the polymerizable diacetylenic lipid Bronco).14

5.4 LB film characterization with SPM

Most of the facts discussed in the previous sections of this chapter were originally discovered using fluorescence microscopy.14,24 However, there are many other monolayer characterization methods to determine domain shapes, layer thickness or lattice constants of crystalline domains, as mentioned in the first chapter.

Now we will show how surface plasmon microscopy (see also Chapter 1) can be used to image and characterize phase-separated lipid LB films.

5.4.1 Experimental section

The substrate preparation and setup that was used is described in the previous chapter. The 2.5 nm thick SiO2 layer (without pattern) is in this case needed to make the substrate hydrophilic for deposition of the Langmuir-Blodgett lipid monolayer. LB films were obtained by spreading a 1 mg/ml solution of the lipid dimiristoylphosphatidylethanolamine (DMPE) in 3:1 (v/v) chloroform/methanol onto the subphase (deionized water, pH~7.5). The monolayer was transferred to the substrate at a surface pressure of 20 mN/m and a molecular area of 0.5 nm2; the transfer ratio was about 1. The DMPE monolayer is under those conditions in an LE-LC phase transition (see Fig. 5.9). All experiments were carried out at room temperature.

5.4.2 Results and discussion

Using the image acquisition techniques that were described in Chapter four, the DMPE monolayer was imaged for a wavelength of 568.2 nm. Figure 5.11 shows the resulting images for p and s polarization, the ratio of the two and the result after Fourier filtering to remove some residual fringes. For this wavelength the optical thickness variations in the monolayer under investigation hardly give any contrast

50 mm (a)(b)(c)(d)

Fig. 5.11 SPM images of a phase-separated DMPE monolayer: (a) for p polarization; (b) for s polarization; (c) the ratio of the two; (d) the result after Fourier filtering to remove some residual fringes.

in the separate images. After division of the two however, a good contrast results and the lateral resolution is higher than it would have been for a longer wavelength (usually a wavelength of 632.8 nm is used). Due to the high dynamic range of the images, quantization levels do not become visible after the expansion of the gray scale. The dark 'islands' in the image are the LC domains surrounded by the LE phase. Because the monolayer was imaged under an angle only the middle part is optimally focussed.

50 mm

Fig. 5.12 SPM image of a DMPE monolayer in a transition from the LE to the LC phase. Contrast inversion was obtained by choosing a different angle of incidence. On the left and right side the LC and LE part are at resonance, respectively.

By choosing either the resonance angle for the LE phase or the one belonging to the LC part of the monolayer as angle of incidence, the contrast can be inverted (see Fig. 5.12). We estimate the lateral resolution that was obtained to be 3 \(\mu\)m (NA 0.19).

To evaluate the thicknesses of the different domains, microscopic ATR scans were made of three different areas: the bare gold-SiO2, an LE domain, and an LC domain. The ATR curves for the covered areas were shifted relative to that of the bare area with 0.17° and 0.23°, respectively (Fig. 5.13). In Fig. 5.14 the possible thickness and refractive index combinations are given for both domains. Assuming a refractive index of 1.5 (Ref. 25) for both domains, the imaged thickness difference is less than 0.4 nm. When a higher refractive index is chosen for the condensed domain and a lower one for the expanded domain, the resulting thickness difference is even smaller. These values have been found by others as well, for similar systems.26,27 If it were possible to determine refractive index and monolayer thickness separately, then the nature of the monolayer packing could be further studied. The reflectance difference in this image, based on the shift of the reflectance curves, is 0.5 %. This is about the limit for practical SPM imaging, since the contrast should be sufficient to focus using the real-time image. 0.30.4 -3.0-2.5-2.0 -1.5-1.0 LELCAu LCLE RMS difference Reflectance (P/S) Angular Shift Df (deg.) External Angle f (deg.) (a)(b)

Fig. 5.13 (a) Microscopic reflectance scans for the DMPE layer, for a bare, LE, and LC region in an image. (b) The root-mean-square value of the difference of the LE and LC curves with the curve of the uncovered part when they are shifted towards this curve along the f-axis. LELC Refractive Index Thickness (nm)

Fig. 5.14 Possible refractive index and thickness combinations as determined from the angular shifts for the LE and LC domains. The thickness difference of both domains is indicated for a refractive index of 1.5.

5.5 References

(1) Gorter, E.; Grendel, F. J. exp. Med. 1925, 41, 439.

(2) Singer, S. J. In Membrane Structure and Function; Rothfield, L. I., Ed.; Academic Press: New York, 1971.

(3) Singer, S. J.; Nicolson, G. L. Science 1972, 175, 720.

(4) Booij, H. L.; Daems, W. T., Eds., Biomembranen 50 jaar na Gorter en Grendel; Centrum voor landbouwpublicaties en landbouw documentatie: Wageningen, 1976.

(5) Benga, G., Ed., Structure and properties of cell membranes, volume III; CRC Press: Boca Raton, 1985.

(6) Earnshaw, J. C.; Winch, P. J., Transitions in model membranes, SPIE 1403, 316-325 (1990).

(7) Birdi, K. S. Lipid and biopolymer monolayers at liquid interfaces; Plenum Press: New York, 1989.

(8) Vogel, V.; Möbius, D. Thin Solid Films 1985, 132, 205.

(9) Menger, F. M.; Wood, M. G.; Richardson, S. D.; Zhou, Q. Z.; Elrington, A. R.; Sherrod, M. J. J. Am. Chem. Soc. 1988, 110, 6797.

(10) Helm, C. A.; Möhwald, H.; Kjær, K.; Als-Nielsen, J. Europhys. Lett. 1987, 4, 697.

(11) Heckl, W. M.; Lösche, M.; Scheer, H.; Möhwald, H. Biochim. Biophys. Acta 1985, 810, 73.

(12) Langmuir-Blodgett Films; Roberts, G. G., Ed.; Plenum Press: New York, 1990.

(13) McConnell, H. M.; Moy, V. T. J. Phys. Chem. 1988, 92, 4520.

(14) Möhwald, H. Thin Solid Films 1987, 159, 1.

(15) Lösche, M.; Rabe, J.; Fischer, A.; Rucha, B. U.; Knoll, W.; Möhwald, H. Thin Solid Films 1984, 117, 269.

(16) Vogel, V.; Möbius, D. Thin Solid Films 1985, 132, 205.

(17) Gaub, H. E.; Moy, V. T.; McConnell, H. M. J. Phys. Chem. 1986, 90, 1721.

(18) Keller, D. J.; Korb, J. P.; McConnell, H. M. J. Phys. Chem. 1987, 91, 6417.

(19) Flörsheimer, M.; Möhwald, H. Thin Solid Films 1988, 159, 115.

(20) Moy, V. T.; Keller, D. J.; Gaub, H. E.; McConnell, H. M. J. Phys. Chem. 1986, 90, 3198.

(21) McConnell, H. M.; Keller, D. J. Proc. Natl. Acad. Sci. USA 1987, 84, 4706.

(22) McConnell, H. M.; Keller, D. J.; Gaub, H. E. J. Phys. Chem. 1986, 90, 1717.

(23) Lösche, M.; Helm, C.; Mattes, H. D.; Möhwald, H. Thin Solid Films 1985, 133, 51.

(24) Heckl, W. M.; Miller, A.; Möhwald, H. Thin Solid Films 1987, 159, 125.

(25) Kooyman, R. P. H.; Krull, U. J. Langmuir 1991, 7, 1506.

(26) Lösche, M.; Rabe, J.; Fischer, A.; Rucha, B. U.; Knoll, W.; Möhwald, H. Thin Solid Films 1984, 117, 269.

(27) Hickel, W.; Knoll, W. J. Appl. Phys. 1990, 67, 3572.