2636
Langmuir 2009, 25, 2636-2640
Preparation and Characterization of Multilayer Coated Microdroplets: Droplet Deformation Simultaneously Probed by Atomic Force Spectroscopy and Optical Detection Hainer Wackerbarth,† Peter Scho¨n,‡ and Ute Bindrich*,† German Institute of Food Technology, Prof.-Von-Klitzing-Str. 7, D-49610 Quakenbru¨ck, Germany, and Veeco Instruments GmbH, Life Science Team, Dynamostrasse 19, D-68165 Mannheim, Germany ReceiVed September 4, 2008. ReVised Manuscript ReceiVed December 31, 2008 We report the preparation and characterization of multilayer coated droplets in an emulsion. Stability and control of mass transport across the interface are the key issues for such coated microdroplets. Shelf life of cosmetic, pharmaceutical, and food formulations can be improved by increasing stability. Moreover, such emulsions have potential applications in drug delivery and storage. A primary oil-in-water emulsion with caseinate as an emulsifier was prepared. On the basis of attractive electrostatic interactions, polyelectrolytes with opposite charges were added layer by layer. The oil droplets (particle size around 10 µm) were successively coated with casein, pectin, whey proteins, pectin, whey proteins, and pectin. Laser diffraction spectroscopy, particle charge measurements, and confocal laser scanning microscopy were applied to characterize the multilayer droplets. The complementary results indicate that the inner layers merge and the packing density of the interface increases. AFM-induced mechanical compression of single oil droplets coated with casein and pectin is monitored by an inverted optical microscope, and simultaneously AFM force curves are recorded. Thus, the deformation of the droplet is reflected by its lateral expansion and the force curve. Force volume imaging is applied to probe the lateral distribution of mechanical properties of the droplet.
Introduction The sequential assembly of charged polyelectrolytes by a layerby-layer technique has become a useful tool for emulsion preparation.1 The stability of droplets against coalescence and thus the stability of emulsion can be improved by such an approach. Shelf life of cosmetic, pharmaceutical, and food formulations can be prolonged. In addition, emulsions based on such microstructures have potential applications in drug delivery and storage. In the food industry, proteins and polysaccharides are used for the control of microstructure, texture, flavor, and shelf life of emulsions.2,3 The stability of oil-in-water (O/W) emulsions containing protein-coated droplets can often be improved by adsorption of a polysaccharide layer on the droplet surface. These layers protect the droplets from aggregation by repulsive interactions.4 Emulsions based on multilayer coated oil droplets often show stability to environmental stresses such as thermal treatments, freezing, thawing, and mechanical agitation.5 Moreover, multilayer droplets of oil-in-water emulsions have potential in protecting lipid components and for controlled flavor release.6 A number of approaches have been developed to produce stable emulsions with multilayer droplets. The approaches can be divided into three groups: (1) saturation (i.e., all the polyelectrolytes from the solution are adsorbed), (2) centrifugation * To whom correspondence should be addressed. Tel.: +49 (0) 5431 183 130. Fax: +49 (0) 5431 183 144. E-mail:
[email protected]. † German Institute of Food Technology. ‡ Veeco Instruments GmbH. (1) Grigoriev, D. O.; Miller, R. Curr. Opin. Colloid Interface Sci. 2008, 14, 48–59. (2) Guzey, D.; McClements, D. J. AdV. Colloid Interface Sci. 2006, 128-130, 227–248. (3) Dickinson, E. Food Hydrocolloids 2003, 17, 25–39. (4) Guzey, D.; McClement, D. J. J. Agric. Food Chem. 2007, 55, 475–485. (5) McClements, D. J. Curr. Opin. Colloid Interface Sci. 2004, 9, 305–313. (6) Aoki, T.; Decker, E. A.; McClements, D. J. Food Hydrocolloids 2005, 19, 209–220.
of excess polyelectrolyte, and (3) filtration of excess electrolyte. The key issue is to avoid droplet aggregation due to the presence of excess free polyelectrolyte.2 Sodium caseinate, pectin, and whey protein are commonly encountered in manufactured food colloids.3 Sodium caseinate and whey protein have a positively charged surface at a pH below the isoelectric point (IP). The characteristic structure of pectin is a linear chain of R-(1-4)-linked D-galacturonic acid. Because of the carboxyl groups, pectin is negatively charged in the deprotonated form. Thus, attractive interactions between caseinate-pectin and whey protein-pectin at low pH are expected. Surface charge of particles can be monitored by streaming potentials and streaming currents. These arise when a solution flows through a capillary tube or a porous plug and charged particles are adsorbed to the surface, while the counterions in the diffuse layer of the particles adjacent to the surface migrate with the fluid.7,8 A cross section of an emulsion droplet can be monitored by confocal laser scanning microscopy (CLSM), as it visualizes two-dimensional images of three-dimensional microscopic structures.9 It is a commonly used characterization technique in biological, pharmaceutical, and food science. Atomic force microscopy (AFM) has become a powerful tool to visualize colloidal particles directly in liquid medium. Moreover, force curves between the AFM tip and a droplet or two droplets can be recorded to investigate the mechanical properties of single droplets.10-12 Information about surface (7) Kam, S.-k.; Gregory, J. Colloids Surf., A 1999, 159, 165–179. (8) Barron, W.; Murray, B. S.; Scales, P. J.; Healy, T. W.; Dixon, D. R.; Pascoe, M. Colloids Surf., A 1994, 88, 129–139. (9) Sheppard, C. J. R.; Shotton, D. M. Confocal Laser Scanning Microscopy; Royal Microscopical Society Microscopy Handbook Series 38; BIOS Scientific Publishers: Oxford, 1997. (10) Dagastine, R. R.; Stevens, G. W.; Chan, D. Y. C.; Grieser, F. J. Colloid Interface Sci. 2004, 273, 339–342. (11) Gunning, A. P.; Mackie, A. R.; Wilde, P. J.; Morris, V. J. Langmuir 2004, 20, 116–122.
10.1021/la802898k CCC: $40.75 2009 American Chemical Society Published on Web 02/06/2009
Characterization of Multilayer Coated Microdroplets
Langmuir, Vol. 25, No. 5, 2009 2637
tension and reciprocal droplet size can be derived from force curves providing insight into stability of emulsions.13 In the socalled force volume imaging mode, an array of force curves is collected over a selected area. Such an array gives information about the lateral distribution of mechanical and adhesive properties of a single droplet. In this study, an oil-in-water emulsion with multilayer droplets was prepared. On the basis of the attractive interactions, the oil droplets were coated in the following sequence with six layers: casein, pectin, whey proteins, pectin, whey proteins, and pectin. Laser diffraction spectroscopy, confocal laser scanning microscopy, and particle charge measurements were applied for characterization of the multilayer coated droplets. First results of AFM force spectroscopy and simultaneously probed optical detection as well as force volume imaging on oil droplets coated with casein and pectin are presented.
Experimental Section Materials. Sunflower oil was purchased from the supermarket. Oil, apple pectin (degree of esterification 30%, Herbstreith & Fox KG), whey protein (Globulal 70 A, Meggle), and sodium caseinate (Fonterra, ca. 4% moisture, spray dried) were used without further purification. Citric acid, 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), tetrasodium pyrophosphate, and phosphate had analytical grade. Distilled water was used for the preparation of all solutions. Primary Emulsion. First, a primary oil-in-water sodium caseinatestabilized emulsion was prepared (Figure 1A) with 68.7 wt % water, 1.1 wt % sodium caseinate, and 30.2 wt % sunflower oil. The aqueous phase containing sodium caseinate was adjusted to pH 3.1 with 0.1 M citric acid and stirred for 120 min at 23 °C. Pre-emulsifying with sunflower oil with a crown stirrer (1800 min-1, 2 min, 60 °C) was performed after oil and aqueous solution were set to 60 °C for 20 min. The pre-emulsion was dispersed by an Ultra-Turrax (16 000 min-1, 2 min) and finally cooled to 25 °C. As the sodium caseinatestabilized emulsion was prepared at pH 3.1, the surface of the coated oil droplet was positively charged. Multilayer Droplet Preparation. Washing Procedure. Before adsorption of the second layer the droplets were washed. The washing procedure included two steps: (1) centrifugation (Cyrofuge 8000, Heraeus) at 1000 min-1 and 25 °C for 10 min followed by separation of the continuous phase and (2) redispersing aqueous solution pH 3.1 (citric acid). The washing procedure was performed two times (Figure 1B). Adsorption of Anionic Layer. Pectin solution (0.8 wt %) was solved in water and adjusted to pH 3.1. The solution was added to the washed primary emulsion (pH 3.1). After being mixed, the emulsion was centrifuged at 200 min-1 for 10 min to separate the continuous phase. The emulsion was adjusted to pH 3.1 (citric acid) and redispersed in aqueous solution (pH 3.1, citric acid). Centrifugation and redispersing were performed twice. Adsorption of Cationic Layer. Whey protein (1.4 wt %) solution was set to pH 3.5 and mixed with the redispersed emulsion. The emulsion was centrifuged at 1000 min-1 for 10 min, adjusted to pH 3.1 (citric acid), and redispersed in aqueous solution (pH 3.1, citric acid). Centrifugation and redispersing were performed twice. Laser Diffraction Spectroscopy. A Malvern Mastersizer X laser diffraction spectroscope was used. A prerequisite for the laser diffraction spectroscopy was low particle concentrations. Low concentrations enabled us to focus on single particles within a defined volume. Moreover, because of the minimization of the free surface energy, the droplets took a spherical shape in emulsion with low concentration, which was the ideal case for the laser diffraction spectroscopy. Thus, the emulsions were diluted 1:20 000 in tetrasodium pyrophosphate solution. (12) Butt, H.-J.; Cappella, B.; Kappl, M. Surf. Sci. Rep. 2005, 59, 1–152. (13) Gillies, G.; Prestidge, C. A. AdV. Colloid Interface Sci. 2004, 108-109, 197–205.
Figure 1. Schema for the preparation of the primary emulsion (A) and multilayer coated oil droplets (B).
Charge Analyzing System. The procedure to determine the charge of the particle surface was carried out with charge analyzing system (Emtec). Particles with a cationic surface were titrated with sodium poly(vinyl styrol) (SPVS) (0.001 N) (Emtec); for anionic surfaces poly(diallyl dimethylammonium) chloride (PDDA) (0.001 N) (Emtec) was used. Titration was performed until a zero streaming potential was obtained (corresponding to zero streaming current). The consumption of electrolyte was noticed. Confocal Laser Scanning Microscopy. A Nikon Eclipse E600 confocal laser scanning microscope was used. Immobilization was carried out with carboxymethyl cellulose. To visualize proteins, the dye rhodamin B (Sigma-Aldrich) was used. Atomic Force Microscope. A Bioscope II atomic force microscope (Veeco Instruments, Inc.) combined with a Leica DMI 6000B inverted optical microscope (Leica Microsystems) was used in the present study. The Bioscope II AFM was equipped with a NanoScope V controller (Veeco Instruments). AFM data were collected and analyzed using Nanoscope v7.20 software. Force spectroscopy and force volume imaging experiments were performed in aqueous solution using DNP cantilevers (Veeco Probes) with nominal spring constants of 0.32 N/m. Emulsions were diluted with phosphate-
2638 Langmuir, Vol. 25, No. 5, 2009
Wackerbarth et al.
Figure 3. Consumption of cationic (PDDA) and anionic (SPVS) electrolyte to reach zero streaming potential after adsorption of a new layer.
Figure 2. X50.3 value (A) and specific surface area (B) of droplets dependent on the number of layers.
buffered saline (pH ca. 7.4) and exposed to polylysine-modified glass bottom dishes for 1 h and then directly mounted to the instrument.
Results and Discussion O/W emulsions based on multilayer coated droplets were prepared using the layer-by-layer technique. First, a primary emulsion was prepared by adsorption of sodium caseinate at the interface (Figure 1A). Then the pectin and whey protein were alternately adsorbed by tuning the pH to generate electrostatic attraction (Figure 1B). In this way, layer by layer was added until the oil droplets were coated with six layers (caseinate, pectin, whey protein, pectin, whey protein, and pectin). The outer layer of the droplets was due to the pectin molecules negatively charged stabilizing the emulsion by repulsive electrostatic forces. Particle Size Distribution. Particle size distribution and the specific surface area were determined by laser diffraction spectroscopy. The specific surface area is an integral value reflecting the state of dispersion. Additionally, the X50.3 values are given. The parameter, X, characterizes a physical property uniquely related to the particle size. It is assumed that a unique relationship exists between the volume and the diameter unequivocally defining “size”. The 50 indicates that 50% of all particles have a diameter smaller than the indicated one. The droplet size of the primary emulsion increased by adding the first pectin layer from 10.18 to 11.64 µm as displayed by the X50.3 value in Figure 2. By adding further layers, the droplet size was not significantly affected. The droplets covered by six layers show an X50.3 value of 11.61 µm. The same tendency was observed
for the specific surface area. The primary emulsion had a specific surface area of 1.7 m2/g, which shrank to values close to 1.4 for further layers. Generally, the larger the specific surface area, the smaller the particles are in the emulsion. Thus, both the X50.3 value and the specific surface area show a saturation of the growth of the droplet volume after the first pectin layer. Particle Charge Measurements. Almost all colloidal substances or solids in aqueous solutions carry an electrical charge. Therefore, the surface is surrounded by a counterion cloud. A flow driven distortion of the counterion cloud leads to a stream potential or current that can be detected by electrodes. When both electrodes are held at the same potential, the streaming current was measured directly as the electrical current or potential flowing through the electrodes. The sign of the streaming potential indicates the particle charge. Titration of polyelectrolyte with opposite charge related to the surface charge of the particle was performed. The consumption of cationic or anionic polyelectrolyte to reach the zero streaming potential is given (Figure 3). Cationic electrolyte (PDDA) was needed to reach zero streaming potential, if the outermost layer consisted of pectin; an outermost layer of whey proteins needed anionic electrolyte (SPVS) for charge neutralization. The titration was performed after preparation of each new layer of the droplet. The data clearly reflect the change of the surface charge of the microdroplets after adding a layer. Thus, it is evident that the microdroplet is coated with alternating charged layers. The consumption of electrolyte should be related to the charge density of the surface. When we take into account the unaltered specific surface area of each layer (Figure 2), whey proteins obviously constitute a surface with a higher charge density than the pectin at pH 3.1. Confocal Scanning Laser Microscopy. In contrast to conventional light microscopy, CLSM restricts the illumination to a single point of the specimen (point scanning illumination), and by inserting a confocal imaging aperture in the optical system, almost all light emanating from regions above and below the focal plane is physically prevented to contribute to the image. Therefore, to a first approximation the image contains only infocus information and permits noninvasive optical sectioning of various z-axis planes.9 For imaging of coated droplets in an emulsion the protein phase was colored with rhodamin B, resulting in red fluorescent light by 543-nm excitation. A CLSM image of several droplets is shown in Figure 4. The diameter of a droplet in the enlargement is around 10 µm, agreeing well with the determined X50.3 value of 11.61 µm of the multilayer coated
Characterization of Multilayer Coated Microdroplets
Figure 4. CLSM image (57 µm × 51 µm) and enlargement of a droplet of the multilayer droplet emulsion; the protein phase is colored with rhodamin B, and excitation is 543 nm.
Figure 5. Schema of the structure of the coated droplets.
droplets. The dark area indicates the absence of protein, and the red area indicates the localization of protein layer. Thus, the red circles visualize the protein layer around the oil droplets. The thickness of the outer layer can be approximated to 200 nm. The inner layer is in the same range, but blurred toward the center. Moreover, one protein layer cannot be resolved, indicating that the inner layers merge. It looks as though the protein diffuses also in the oil phase. The particle charge measurements clearly indicate alternating surface layers, and at the same time the particle size distribution indicates no further increase after addition of the first pectin layer. The CLSM provides an explanation. The inner layers merge and diffuse maybe even in the oil phase, whereas the outer layer is quite defined, enabling the adsorption of opposite charged molecules. The merging of the inner layers might be driven by attraction of opposite charged molecules, causing a dense packing. In Figure 5, the resulting multilayer droplet structure is schematically shown. Addition of layers, thus, results in a more compact interface. By adjusting the compactness of the interface by the amount of adsorbed layers, the mass transport of molecules across the interface can be controlled, which can be used for the prevention of release of functional ingredients (vitamins, Ω-3 fatty acids) and flavors. On the other side, the multilayer coatings can protect labile substance and function as a shuttle in drug delivery. AFM Measurements. Scanning probe microscopies are still rarely used in food science.14 AFM including force spectroscopy has been applied to investigate colloidal particles.11 In the present study, force spectroscopy combined with transmission light (14) Morris, V. J. Trends Food Sci. Technol. 2004, 15, 291–297.
Langmuir, Vol. 25, No. 5, 2009 2639
microscopy and force volume imaging on a double layer coated oil droplet was performed in buffer solution at pH 7.4. The interface of the O/W emulsion was stabilized with sodium caseinate covered by pectin. The double layer microdroplets were adhered to polylysine-modified glass substrates. The negatively charged droplets were adsorbed via electrostatic interaction with the positively charged lysine groups of the surface. By optical microscopy, single droplets could be identified and the AFM tip straightforwardly navigated to the droplet of interest (Figure 6). The AFM tip was moved on top of the droplet (Figure 6A). The tip was then moved further toward the droplet so that a pressure was applied. The pressure caused deformation of the droplet as depicted in Figure 6. Figure 6B,C shows the impact of an increasing compression force: high force induced an expansion of the projected diameter of the droplet from 12 to 15 µm, which is 25% (Figure 6B). The expansion becomes evident by focusing on the two smaller droplets (ca. 8 and 5 µm in diameter) near the droplet below the tip, whose size remained unchanged as no force was applied to them (Figure 6A,B). When we reduced the force, the droplet shrank to its original diameter, indicating the reversibility of the compression and the elasticity of the droplet interface. The force curve recorded in the center of the droplet is shown in Figure 6C. It contains the force as function of travel distance in both approach (black) and retract (red) directions. Several points in the force curve can be associated with different events during approach/retract: 1 indicates the engagement of the AFM tip onto the microdroplet, regime 2 indicates the indentation of the microdroplet, point 3 indicates the reversal of the z travel motion, and regime 4 shows adhesion caused by sticking of the AFM tip to the microsphere. At point 5, the tip releases from the droplet. Generally, a force curve could provide a spring constant that is proportional to the surface tension and the reciprocal of the droplet radius according to Gillies and Prestidge.13 A disadvantage of force measurements on soft materials is that there is no clear transition from noncontact to contact. Surface deformations occur before contact because of the extended range of surface forces and fluid dynamics.15 The tip might cause an indentation or flattening of the droplet. Force-distance functions can be used to describe rheological properties of interfaces of real emulsion droplets and may be an additional tool for correlations of composition, structure, and functionality. Another tool for imaging surfaces is force volume imaging, in which an array of force curves is collected over a selected area. In contrast to AFM imaging modes such as contact mode in which the AFM tip is scanned with constant force in continuous contact with the sample, force volume imaging eliminates lateral or shear forces by retracting the tip and moving without contact to the selected coordinates of the surface. By that, damage of fragile samples can be avoided. Recently, the advantages of this technique were demonstrated on probing a bacterial surface.16 The force image obtained in the force volume mode (Figure 7) shows a topography map of the droplet surface (diameter of the surface ca. 12 µm) obtained by indenting with a defined force set by a trigger value (see also Figure 6C, point 3). The color schema is height encoded (the brighter, the higher) and reflects the curvature of the droplet. At the same time, the force volume indicates homogeneous elasticity/softness of the droplet, which can be interpreted as a homogeneously covered surface. The preliminary AFM data clearly show the potential of this approach and will pave the way for further investigations. (15) Withers, J. R.; Aston, D. E. AdV. Colloid Interface Sci. 2006, 120, 57–67. (16) Gaboriaud, F.; Parcha, B. S.; Gee, M. L.; Holden, J. A.; Strugnell, R. A. Colloids Surf., B 2008, 62, 206–213.
2640 Langmuir, Vol. 25, No. 5, 2009
Wackerbarth et al.
Figure 6. (A) Bright field image and sketch of the droplet and tip; no contact between tip and droplet. (B) Bright field image and sketch of the droplet deformation caused by the AFM tip. (C) Force curve recorded on the droplet. The oil droplet is surrounded by a caseinate-pectin layer; the measurements are performed in solution. The force curve contains the force as function of travel distance in both approach (black) and retract (red) directions. Several points in the force curve can be associated with different events during approach/retract: 1 indicates the engagement of the AFM tip onto the microdroplet, regime 2 indicates the indentation of the microdroplet, point 3 indicates the reversal of the z travel motion, and regime 4 shows adhesion caused by sticking of the AFM tip to the microsphere. At point 5 the tip releases from the droplet.
Figure 7. Force image obtained on an oil droplet surrounded by a caseinate-pectin layer; z-scale: 12 µm.
Variations of the multilayer structure and interface in combination with AFM techniques will provide insights into the relationship between interfacial structure and deformability of droplets. This would result in a better understanding of coalescence, which is of great interest for academia and industry.
Conclusions An O/W emulsion with multilayer coated droplets was prepared by a layer-by-layer technique. The primary emulsion was stabilized with caseinate. Using attractive interactions between oppositely charged polyelectrolytes, we alternated coating the
droplets of the primary emulsion with pectin and whey protein, leading to a droplet with six layers where the outermost layer consisted of pectin layer. The particle size distribution was determined by laser diffraction spectroscopy. Both the X50.3 value and the specific surface area show a saturation of the growth of the droplets after adsorption of the first pectin layer. The surface charges of the droplets were determined by titration of the streaming potentials, indicating that after each adsorption step the charge changed and thus the addition of a new layer was confirmed. The saturation of the particle size and the alternating surface charge can be explained by an increase of the packing density of the polyelectrolytes on the surface. This interpretation agrees well with observation of CLSM where cross sections of the oil droplets were monitored. The deformation of the droplet was observed by transmission light spectroscopy while an AFM tip pressed on the droplet. The detected force curves showed the reversibility of the mechanical deformation. Force volume imaging was performed on the droplet, indicating a homogeneous surface layer. Acknowledgment. This work was supported by “Bundesministerium fu¨r Wirtschaft und Technologie (BMWi)” via the AiF/ FEI Project, No. 15218N. LA802898K
