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Measurement of Wall Zeta Potentials and Their Time-Dependent Changes Due to Adsorption Processes: Liposome Adsorption on Glass Miguel D. Reboiras,† Michael Kaszuba,‡ Malcolm T. Connah,‡ and Malcolm N. Jones* School of Biological Sciences, University of Manchester, Manchester, M13 9PT, United Kingdom, and Malvern Instruments Ltd., Engima Business Park, Graveword Road, Malvern, Worcestershire, WR14 1XZ, United Kingdom Received March 14, 2001. In Final Form: June 5, 2001 A novel method of measuring the zeta potential associated with the planar glass surface of an electrophoresis cell by laser Doppler velocimetry has been developed. The method exploits the observation that the terminal velocity of particles subjected to an electric field is reached at least an order of magnitude more quickly than the establishment of electroosomotic flow at the planar glass surface of the cell. By using a fast field reversal technique so that the particle velocity is unaffected by electroosomic flow in conjunction with a mixed mode measurement, it is possible to obtain the electrophoretic mobility of a particle without the need to make measurements at the stationary layer. In addition, the methods give data for the zeta potential at the cell wall. The cell wall zeta potential is sensitive to adsorption of solutes from solution and may be used to follow both the kinetics of adsorption and to construct an adsorption isotherm. The method has been applied to the study of the adsorption of cationic and anionic phospholipid liposomes. The wall zeta potentials reflect the adsorption process in that the cationic liposomes cause a reversal of the wall zeta potentials from the negative value of the clean glass surface to positive values after 1-3 h. In contrast, the anionic liposomes increase the negative wall zeta potentials. At equilibrium, limiting potentials may be fitted to a Langmuir isotherm. The results from the fitting are compared with the adsorption of liposomes to silica particles in suspension.
Introduction The zeta potential of colloidal dispersions has been measured using electrophoresis in a capillary cell for more than 50 years.1-4 Measurements using the stationary layer technique have been shown to give better quality data than any other proposed cell type; however, measurements at the stationary layer require that a number of practical considerations have to be taken into account. This means that the potential accuracy and resolution available are not always achieved. Electrophoresis is the phenomenon whereby particles with a net charge migrate when subjected to an electric field. Because micron-sized particles have a very low inertia, terminal velocity is reached in microseconds. The actual velocity is determined by the zeta potential of the particles, the viscosity and relative permittivity of the medium, and applied field strength. The velocity in a unit field (e.g., 1 V/m) is known as the particle electrophoretic mobility. The zeta potential can be calculated from the electrophoretic mobility using the Henry equation1 at low zeta potentials or the exact computer solutions of the mobility equations of O’Brien and White.5 In this study, * To whom correspondence should be addressed. E-mail: mjones@ fs1.scg.man.ac.uk. † Departmento de Quimica, Facultad de Ciencas, Universidad Autonoma de Madrid, Madrid 29049, Spain. ‡ Malvern Instruments Ltd. (1) Henry, D. C. Proc. R. Soc. London, Ser. A 1931, 133, 106. (2) Shaw, D. J. Introduction To Colloid And Surface Chemistry; Butterworth-Heinemann; Oxford, U.K., 1992. (3) Preece, A. W.; Brown, K. Advances in Electrophoresis. In Recent Trends in Particle Electrophoresis; Dunn, M. J., Radola, B. J., Eds.; VCH: New York, 1989. (4) Uzgiris, E. E. Prog. Surf. Sci. 1981, 10, 53. (5) O’Brien, R. W.; White, L. R. J. Chem. Soc., Faraday Trans. 2 1978, 74, 1607.
the data cover a range of zeta potentials from 0 to (40 mV. To avoid inconsistencies, the Henry equation was used throughout. Electroosmosis is the movement of a liquid containing ions, near to a charged surface when a field is applied along the surface. In a quartz cell, the velocity of the electroosmotic flow is the same order of magnitude as the particle velocity. This flow takes much longer to establish than electrophoresis, in the order of tens of milliseconds. In a capillary which is closed at both ends, the liquid flow along the cell wall returns through the cell and generates a parabolic profile with origin at the center of the cell. The particle mobility measured where the fluid flow parallel to the cell wall cancels out the return flow is the true particle mobility. The position where the two fluid flows cancel is called the stationary layer (Figure 1a). This method can give good resolution and accuracy; however, the stationary layer must be located very precisely. Some systems enable a mobility measurement at the stationary layer on both sides of the cell or a cell scan to confirm alignment, but this increases the cost and complexity of the instrument. A theoretical electrokinetic analysis has shown that after the application of an electric field to a capillary cell, colloidal particles suspended in the liquid reach terminal velocity at least an order of magnitude more quickly than the establishment of electoosmosis.6 Therefore, if an alternating electric field is applied with a sufficient high frequency then the liquid velocity due to electroosmosis becomes insignificant with respect to the electrophoretic mobility. This means that measurements do not have to be taken at the stationary layer in the capillary cell, (6) Minor, M.; van der Linde, A. J.; van Leeuwen, H. P.; Lyklema, J. J. Colloid Interface Sci. 1997, 189, 370.
10.1021/la010391a CCC: $20.00 © 2001 American Chemical Society Published on Web 07/27/2001
Measurement of Wall Zeta Potentials
Figure 1. (a) Electroosmotic flow and stationary layer at low frequency of field reversal. (b) Electroosmotic flow profile at high frequency of field reversal.
removing the need for alignment. This measurement strategy is called fast field reversal (FFR) and gives an accurate mean, but of lower resolution than the standard stationary layer technique. All systems that measure mobilities using LDV (laser Doppler velocimetry) reverse the applied field periodically. This is to reduce the polarization of the electrodes that is inevitable in a conductive solution. The field is usually reversed about every second to allow the fluid flow to stabilize (Figure 1a). If the field is reversed much more rapidly, it is possible to show that the particles reach terminal velocity, while the fluid flow due to electroosmosis is insignificant (Figure 1b). This means that the mobility measured during this period is due to the electrophoresis of the particles only and is not affected by electroosmosis. The mean zeta potential that is calculated by this technique is therefore very robust, as the measurement position in the cell is not critical. However, as the velocity of the particles is sampled for such a short period of time, information about the distribution is degraded. This is what is addressed by a new measurement technique call “mixed mode measurement” (M3).7 This is a new method of making zeta potential measurements that uses the best features of both stationary layer and FFR techniques in a capillary cell. The benefits are improved resolution, an insensitivity to cell alignment, and a reduced sensitivity to cell wall contamination. In addition, the zeta potential of the cell wall can be calculated. The M3 method incorporating FFR is standard software commercially available in the Malvern instruments. M3 consists of both slow field reversal and fast field reversal measurements. In addition to the zero field measurement, which gives a better measurement of the actual distribution width, two measurements are done for each zeta potential determination: first, a fast field reversal measurement to provide the accuracy and stability of the result and second, slow field reversal measurements to improve the resolution. The new M3 method uses a cell that positions the measurement zone in the middle of the cell rather than at the stationary layer. In principle, the M3 measurement could be done at any position in the cell; however, there (7) Connah, M. T.; McNeil-Watson, F. K. Patent Application GB 0010377.0, 2000.
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Figure 2. The zeta potential of a carboxyl modified latex as a function of cell position at two frequencies of field reversal: (b) 1 Hz and (9) 30 Hz. The latex has a certified zeta potential of -50 mV ((5 mV). The dotted lines indicate the position of the stationary layer of the cylindrical capillary cell (14.6 and 85.4% of the diameter, respectively).
are a number of reasons for choosing to work at the center of the cell: (1) The measurement zone is further from the cell wall and so reduces the chance of light flare from the nearby surface. (2) The alignment of the cell is even less critical. (3) The charge on the cell wall can be calculated. The measurement of the cell wall charge (or zeta potential) can be used to study the adsorption of particles to the surface of the capillary cell (silica). In this study, we have applied the M3 technique to the adsorption of cationic and anionic liposomes prepared from lipid mixtures of dimethyldioctadecylammonium bromide (DDAB)cholesterol (chol)-dipalmitoylphosphatidylcholine (DPPC) and DPPC-phosphatidylinositol (PI), respectively, to the silica capillary wall. The kinetics of both liposome adsorption and equilibrium adsorption has been studied as a function of liposome concentration. Theory The determination of the wall potential is based on the dependence of the measurement of electrophoretic mobility and derived zeta potential on the frequency of field reversal. Figure 2 shows the zeta potentials of latex particles (DTS 5050) derived from measurements of electrophoretic mobilities by use of the Smoluchowski equation as a function of position in the electrophoresis cell at high (30 Hz) and low (1 Hz) frequency. The parabolic profile of zeta potential at low frequency arises from the sum of particle electrophoresis and electroosmotic flow. The electroosmotic mobility (uE) is determined by the wall zeta potential (ζw) according to the equation8
uE ) -
0rζw η
(1)
where η, 0, and r are the viscosity of water, permittivity of vacuum, and relative permittivity, respectively. When ζw is negative, the space charge is positive and the liquid flows toward the negative electrode. The particle mobility when the particle radius is large relative to the double (8) Hunter, R. Zeta Potential In Colloid Science: Principles And Applications; Academic Press: London, 1981; p 61.
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Figure 3. The zeta potential (mV) of a carboxyl modified latex as a function of the field reversal frequency. The measurements were performed at the center of the capillary cell. The latex has a certified zeta potential of -50 mV ((5 mV).
layer thickness (i.e., high ionic strength) is given by the Smoluchowski approximation,
up )
0rζ η
(2)
where ζ is the zeta potential of the particle. A further requirement for the use of the Smoluchowski approximation is that the surface conductivity of the particles must be small (i.e., the Dukhin number, the ratio of particle surface to bulk conductivities, is small). At electrolyte concentrations >10-2 M, the surface conductivity becomes negligible.9 All the measurements reported here relate to electrolyte concentrations >10-2 M (see below). At a low frequency of field reversal, the measured mobility (uobs) is the sum of uE and up. Under these conditions, the electroosmotic flow can be established before the field reverses. At high frequency, the electroosmotic flow cannot follow the field and hence the particle mobility is not affected by electroosmotic flow and is independent of the position of measurement in the cell. As the frequency is changed from low to high values, a dispersion of mobility (and hence zeta potential) will be observed as shown in Figure 3. It should be noted that the low-frequency parabolic profile in Figure 2 crosses the high-frequency (position-independent) zeta potential line at the conventional stationary layer located at 14.6% and 85.4% of the diameter for a cylindrical cell. The mixed mode measurement makes use of the dispersion of mobility with frequency of field reversal to obtain both the zeta potential of the particles independent of position in the cell (high frequency) and wall zeta potential calculated from uE derived from the difference between uobs at low () uE + up) and high frequency (up). The data shown in Figures 2 and 3 are very similar to those described by Minor et al.6 for silver iodide particles. Experimental Section Materials. L-R-Dipalmitoylphosphatidylcholine (DPPC, product no. P0763), dimethyldioctadecylammonium bromide (DDAB, product no. D2779), and cholesterol (product no. C8667) were from Sigma Chemical Co., Poole, Dorset, U.K. Phosphatidylinositol (PI, from wheat germ) was from Lipid Products, South Nutfield, Redhill, U.K. PBS tablets (code BR 14a) were from Oxoid, Hants, U.K. Filters for preparing VETs were from Poretics, (9) Dukhin, S. S.; Derjaguin, B. V. Surface and Colloid Science; Matijevic, E., Ed.; Wiley-Interscience: New York, 1974; Chapter 2, p 210.
Livermore, CA. All solutions were made up in double-distilled deionized water. Preparation and Characterization of Liposomes. Liposomes were prepared using the vesicle extrusion method10 with a Lipex extruder (Lipex Biomembranes, Inc., Vancouver, BC, Canada). All the liposomes contained DPPC as the major component. The required lipids (total mass of approximately 30 mg) were dissolved in either 3 mL of chloroform-methanol (4:1 by volume) or 20 mL of tert-butyl alcohol for cholesterol-containing liposomes in a 100 mL round-bottomed flask. The solvent was removed by rotary evaporation at 60 °C to leave a thin lipid film. The film was hydrated by the addition of 3 mL of phosphate buffered saline (PBS), pH 7.4, at 60 °C with vigorous mixing using a vortex mixer to give a suspension of multilamellar vesicles (MLVs). The MLVs were then extruded at 60 °C through two stacked polycarbonate filters (pore size of 100 nm) under a nitrogen pressure of 200-500 psi. The extrusion was repeated 5-10 times to produce a uniform suspension of largely unilamellar liposomes. Three liposome compositions were used: DPPC-cholesterol-DDAB mol % (80:11:9) and (71:11:18) and DPPC-PI mol % (91:9). The extruded liposomes were characterized in terms of size by photon-correlation spectroscopy (PCS) using a Malvern Zetasizer 3000. The liposome diameters were 125 nm (9 mol % DDAB), 129 nm (18 mol % DDAB), and 118 nm (9 mol % PI). Measurements of Wall Potentials. Wall zeta potential measurements, derived from electrophoretic mobility measurements, were made using the field-flow reversal (FFR) protocol as described above using the Malvern Zetasizer 3000. The measurements were made at 25 °C in 1/10 dilution PBS which has an ionic strength of 0.0188 M.11 The zeta potentials were calculated from the mobilities using the Smoluchowski approximation (eq 2) for the zeta potential (ζ) corresponding to the Henry factor of 1.51 where η, 0, and r were taken as 8.904 × 10-4 N m-2 s, 8.854 × 10-12 J-1 C2 m-1, and 78.5, respectively. For 1/ 10 dilution PBS, ionic strength of 0.0188 M, the Debye length (1/κ) is 2.190 nm, and hence for liposomes of diameter 124 nm (the mean diameter for the three types used) κa ) 56.52 which corresponds to a Henry factor of 1.43. The use of the Smoluchowski approximation will thus give an error in ζ of approximately 5% depending on the particular liposome preparation. The wall zeta potential measurements were an average of 10 independent measurements. Between measurements on each system, the zeta cell was exhaustively washed with 2% Decon C10 in water and finally 1/10 dilution PBS. The system was then checked using the electrophoresis standard polystyrene latex DTS5050 which has a zeta potential of -50 ( 5 mV. The wall zeta potential of the clean glass cell surface was obtained from measurements at very low cationic and anionic liposome concentrations and times before appreciable adsorption had occurred and was found to be -35 ( 1 mV.
Results Measurements were made on three liposome compositions, two cationic (9 and 18 mol % DDAB) and one anionic (9 mol % PI). The sizes and zeta potentials of the liposomes are shown in Table 1. Measurements were made over a range of liposomal lipid concentration both initially, within the first 10 min of putting the liposomes in the electrophoresis cell (ζ10), and again after the wall potentials had come to equilibrium (ζav) (see below). Ten measurements at each liposome concentration were made, so that the data for the zeta potentials given in Table 1 are averages of at least 50 measurements for each liposome composition. The ζ10 values are lower than the ζav values especially for the 9 mol % DDAB liposomes although the differences are not significant for the other compositions. The change in wall zeta potential as a function of time on exposure to cationic liposomes is shown in Figure 4a,b. (10) Hope, M. J.; Bally, M. G.; Webb, G.; Cullis, P. R. Biochim. Biophys. Acta 1985, 812, 55. (11) Sanderson, N. M.; Guo, B.; Jacob, A. E.; Handley, P. S.; Cunniffe, J. G.; Jones, M. N. Biochim. Biophys. Acta 1996, 1283, 207.
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Table 1. Zeta Potentials of Cationic (DDAB-DPPC-Cholesterol) and Anionic (DPPI-PI) Liposomes in Dilute PBS, Ionic Strength of 0.0188 M, at 25 °C liposome composition (mol %)
dw (nm)
concentration range (mM) (no. of concentrations)
ζ10a (mV)
ζava (mV)
DDAB-DPPC-cholesterol (9/80/11) DDAB-DPPC-cholesterol (18/71/11) DPPC-PI (91/9)
125 129 118
0.02-0.20 (5) 0.005-0.3 (9) 0.3-1.5 (5)
21.4 ( 7.4 32.9 ( 2.7 -38.1 ( 3.0
44.8 ( 8.0 38.6 ( 3.3 -40.6 ( 3.4
a ζ is the zeta potential measured within the initial 10 min; ζ is the zeta potential measured after the wall potential became constant 10 av (40-130 min depending on concentration).
Figure 5. Wall zeta potential as a function of time on adsorption from DPPC-PI (91/9 mol %) liposomes in 0.1 PBS at 25 °C. The liposomal lipid concentrations were as follows: 9, 0.5 mM; b, 0.7 mM; 2, 1.0 mM.
Figure 4. (a) Wall zeta potential as a function of time on adsorption from DDAB-DPPC-cholesterol (9/80/11 mol %) liposomes in 0.1 PBS at 25 °C. The liposomal lipid concentrations were as follows: ), 0.008 mM; 4, 0.009 mM; O, 0.01 mM; 0, 0.02 mM; 3, 0.03 mM; 1, 0.04 mM; 2, 0.06 mM; b, 0.20 mM; 9, 0.34 mM. (b) DDAB-DPPC-cholesterol (18/71/11 mol %) liposomes: 9, 0.005 mM; b, 0.006 mM; 2, 0.008 mM; 1, 0.01 mM; (, 0.015 mM; 3, 0.02 mM; 0, 0.05 mM; O, 0.10 mM; 4, 0.30 mM.
For both compositions of liposomes, the wall potentials change sign from negative values at very low liposomal lipid concentrations to positive values. The development of a stable wall potential at the higher concentrations was slow and took 1-3 h. The change in sign suggests that the cationic liposomes and/or cationic lipid is adsorbing on the silica glass surface of the electrophoresis cell. For the 9 mol % DDAB liposomes, one particular liposome concentration (0.02 mM) gave a biphasic kinetic profile possibly suggesting that the mode of the adsorption changed after approximately 1 h. The kinetic behavior of the anionic liposomes was different from that of the cationic liposomes. In the concentration range of 0.05-0.3 mM, the wall potentials were independent of time; above 0.3 mM, the potentials
Figure 6. Equilibrium wall zeta potential as a function of concentration on adsorption from DDAB-DPPC-cholesterol liposomes in 0.1 PBS at 25 °C. Solid line (9), 18 mol % DDAB; dashed line (b), 9 mol % DDAB.
became more negative in a linear fashion although the absolute changes were not large, amounting to approximately 25% after 100 minutes (Figure 5). These data suggest that anionic lipid and/or liposomes are adsorbing to the silica glass and increasing the negative charge on the surface. Figure 6 shows the limiting wall zeta potentials as a function of liposomal lipid concentration. Both curves show a steep rise from the negative wall potential of the clean silica glass surface to a positive limiting value when the surface becomes saturated with cationic liposomes. Discussion The application of the fast field reversal method in conjunction with the mixed mode measurement has been
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Figure 7. Adsorption data from Figure 6 fitted to the Langmuir equation (solid line): (a) DDAB-DPPC-cholesterol liposomes (9/80/11 mol %) and (b) DDAB-DPPC-cholesterol liposomes (18/71/11 mol %).
applied to the study of the adsorption of liposomes and/or lipid to the planar silica glass surface of the electrophoresis measuring cell. It has been shown that the measurement of the wall zeta potential is a novel way to follow both the kinetics and equilibrium adsorption of the liposomes/lipid to planar silica glass. Both cationic and anionic liposomes adsorb to the glass, increasing the wall zeta potentials to more positive and negative values, respectively. Cationic liposome/lipid adsorption results in surface charge reversal. The data in Figure 6 suggest that adsorption is of the Langmuir type. To test this hypothesis, the data were fitted to the Langmuir isotherm (type 1)12 in the form
ζw )
ζlim [L] Kd + [L]
(3)
where ζw and ζlim are the wall zeta potential and its limiting value at high liposomal lipid concentration [L] and Kd is the dissociation constant. The data in Figure 6 can be fitted to eq 3 provided they are normalized with respect (12) Brunauer, S. Physical Adsorption of Gases and Vapours; Oxford University Press: 1944.
to the potential of the clean glass surface (-35 ( 1 mV). The fits to the data are shown in Figure 7, from which dissociation constants (Kd) of 0.0308 ( 0.0151 mM (9 mol % DDAB) and 0.0375 ( 0.0110 mM (18 mol % DDAB) were obtained. These values of Kd may be used to calculate an association constant (1/Kd) and hence Gibbs energies of association of -25.8 and -25.3 kJ mol -1, respectively, at 25 °C. A number of diverse effects have been observed on the adsorption of phospholipid liposomes on glass surfaces depending on the nature of the glass and the conditions of deposition. Bilayer formation has been found on adsorption of DPPC and dimyristoylphosphatidylcholine (DMPC) unilamellar vesicles.13,14 However, other studies15,16 have shown that for DPPC liposome adsorption on Ballotini glass beads (diameter of 67 nm, surface area of 0.1 m2 g-1) a bilayer is formed below the lipid chain melting temperature (Tc ∼ 41° C) but an expanded monolayer is formed above Tc.16,17 For cationic and anionic liposomes of the type used here, adsorption on silica particles of diameters 440, 330, and 1560 nm was found to be largely in the form of intact liposomes; no significant amount of liposome disruption was detected on adsorption.17 These later studies that were made using a solution depletion technique gave considerably greater Gibbs energies of adsorption. Specifically, for adsorption to 330 nm diameter silica particles for DDAB-DPPC-cholesterol liposomes the values were -57.9 kJ mol -1 (9 mol % DDAB) and -61.9 kJ mol -1 (18 mol % DDAB) and for DPPC-PI liposomes the value was -57.0 kJ mol-1 (9 mol % PI). For adsorption to larger silica particles (1560 nm diameter), the values were -52.3 kJ mol-1 (9 mol % DDAB), -58.4 kJ mol-1 (19 mol % DDAB), and -50.5 kJ mol-1 (9 mol % PI). These data suggest that the Gibbs energy of adsorption decreases with decreasing curvature of the surface so that the lower values found in this work for planar glass may not be out of line. Convex solid surfaces with high radius of curvature might possibly be capable of distorting adsorbed liposomes more than a planar surface, and this might increase the strength of adhesion between liposomes and glass. Finally, one may question to what extent the wall zeta potential measurements may have more general applications to surfaces other than glass. Any material which could be used to coat the glass with a sufficiently optically transparent surface may well be used as the adsorbing surface in this novel technique. Since only a thin surface layer on the glass is required, the technique may well be applied to a wide range of surface layers prepared from polymeric and other surface coatings. Thus, wall potential measurements offer a convenient means to investigate adsorption processes in situ. LA010391A (13) Bayerl, T. M.; Bloom, M. Biophys. J. 1990, 58, 357. (14) Nollert, P.; Kiefer, H.; Ja¨hnig, F. Biophys. J. 1995, 69, 1447. (15) Jackson, S. M.; Jones, M. N.; Lyle, I. G. Colloids Surf., A 1986, 20, 171. (16) Jackson, S. M.; Reboiras, M. D.; Lyle, I. G.; Jones, M. N. Colloids Surf., A 1987, 27, 340. (17) Bennett, T. C.; Creeth, J. E.; Jones, M. N. J. Liposome Res. 2000, 10, 303.
