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Total Internal Reflection Fluorescence Spectrometer To Study Dynamic Adsorption Phenomena at Liquid/Liquid Interfaces Michael J. Tupy, Harvey W. Blanch, and Clayton J. Radke* Department of Chemical Engineering, University of California, Berkeley, California 94720-1462
Adsorption at oil/water interfaces affects the performance of many industrial systems including oil recovery, extraction processes, cosmetic products, and food technology. However, no technique currently available can monitor adsorption dynamics using molecularly sensitive methods. We have constructed a novel total internal reflection fluorescence spectrometer (TIRFS) to follow dynamic adsorption events at the oil/water interface. The TIRFS monitors changes in fluorescence intensity and fluorescence spectra over time by maintaining an optical focus on the fluid interface during adsorption and desorption processes. Kinetic adsorption phenomena are examined by altering the composition of the aqueous phase and recording surface fluorescence response without mechanically disturbing the fluid/fluid interface. The spectrometer captures changes in the fluorescence intensity over tenths of seconds and maintains optical focus for periods of days. Mass transport of fluorescing surface-active material to and from the oil/water interface is accurately modeled using the simple one-dimensional diffusion equation. The geometry designed for this apparatus can be applied to other light-based techniques studying adsorption at liquid/liquid interfaces. Here, we apply the TIRFS apparatus to the study of β-casein adsorption and desorption at an aliphatic oil/water interface. The observed increase in interfacial fluorescence due to β-casein adsorption is slower than the diffusive flux, and desorption is found to be very slow if not irreversible. The TIRF spectrum indicates interaction of sorbed β-casein with the oil phase and subsequent rearrangement of the native structure. Introduction Understanding adsorption phenomena at the oil/ water (O/W) interface is of interest to numerous applications including oil recovery, extraction separations, cosmetics, and food technology. Few studies have applied optics-based techniques to liquid/liquid systems because of the difficult practical issues involved with studying deformable fluid interfaces. Nevertheless, some optical techniques have been applied to liquid/ liquid interfaces including total internal reflection fluorescence spectroscopy (Morrison and Weber, 1987; Wirth and Burbage, 1991; Watarai and Saitoh, 1995; Bessho et al., 1997), Raman resonance spectroscopy (Takenaka and Nakanaga, 1976), second-harmonic generation spectroscopy (Grubb et al., 1988; Higgins and Corn, 1993), sum frequency vibrational spectroscopy (Conboy et al., 1996), attenuated total reflection spectroscopy (Perera et al., 1992), and surface light scattering (Lofgren et al., 1984; Sauer et al., 1986). To date, total internal reflection (TIR) has only been used to study equilibrated liquid/liquid interfaces. Currently available instruments are unable to follow the adsorption/desorption dynamics. Although design approaches to obtain sorption kinetics may be found in solid/liquid total internal reflection fluorescence spectrometer (TIRFS) instrumentation (Harrick and Loeb, 1973; Burghardt and Axelrod, 1981; Van Wagenen et al., 1982; Lok et al., 1983a,b; Tilton et al., 1990; Zimmermann et al., 1990; Liebmann et al., 1991), the solid interface greatly simplifies application. The TIRFS apparatus discussed in this article is able to monitor, in a noninvasive fashion, the arrival of fluorescing surface-active * To whom all correspondence should be addressed. Telephone: 510-642-5204; Fax: 510-642-4778.
components at the O/W interface from the bulk aqueous phase and adsorption. The ability to measure TIRF spectra enables subsequent changes in the adsorbedlayer molecular configuration to be observed (Lakowicz, 1983). Additionally, the TIRFS design allows desorption to be studied without exposing the adsorbed layer to convective currents. Our apparatus is also uniquely able to follow sorption processes that occur over long time scales, such as with protein adsorption. Total Internal Reflection Spectroscopy A major difficulty in studying adsorption phenomena is separating the measured physical properties of the adsorbed species from the properties of the surrounding bulk media present in much larger quantities. Rather than using bulk-phase illumination, total internal reflection studies the adsorbed layer using a shallow surface energy wave, the evanescent wave, produced at the point of reflection. Light directed through medium one with index of refraction, n1, toward a medium of a lower index of refraction, n2, reflects at the interface if the angle of incidence is greater than the critical angle (φcritical ) sin-1(n2/n1)). At the point of reflection, some of the incident light penetrates into the second, less optically dense phase. The intensity of this evanescent wave, E, decreases exponentially with distance from the interface: E(z) ) E0 exp(-z/dp) where z is the distance into the second medium perpendicular to the interface. The depth of penetration, dp, is given by
dp )
λ 2πn1(sin2 φ - (n2/n1)2)1/2
S0888-5885(97)00924-X CCC: $15.00 © 1998 American Chemical Society Published on Web 06/09/1998
(1)
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Figure 1. Schematic of the cross-sectional view of the O/W TIRFS cell (not drawn to scale).
where λ is the wavelength of light and φ is the angle of incidence. Typical depths of penetration in our apparatus are 200 nm. The intensity of the TIRFgenerated fluorescence, F, is calculated by integrating over the region interacting with the evanescent wave:
∫0∞c(t,z) (t,z) ψ(t,z) E(t,z) dz
F(t) ) χ
(2)
where χ is an instrument parameter, c is the concentration of the fluorescent molecule, is the extinction coefficient, and ψ is the quantum yield, each written as functions of time and position to reflect the dynamic processes occurring in the adsorbed layer (Harrick, 1967). Although simple in principle, application of TIRFS to the study of adsorption dynamics at fluid interfaces presents numerous experimental challenges. Apparatus Description Design Criteria. To study sorption dynamics at liquid/liquid interfaces, several practical issues must be overcome. The most challenging issues center around maintaining focused optics on the fluid interface for the duration of the experiment. The location of the interface is constrained to the field of view and depth of focus of the detection optics and the coinciding focal point of the incident excitation light. The O/W interface can move due to a variety of disturbances. Room vibrations cause the interface to fluctuate. Adsorption/desorption processes alter the interfacial tension and the wetting competition between the oil and water at the contact line, changes that both affect interfacial curvature and thus the location of the O/W surface. Any convection near the interface induces interfacial flow and may cause the fluid interface to fluctuate or shear. Our apparatus mitigates these disturbances. Additionally, mass transfer of the surface-active agent to and from the interface must be understood quantitatively to make any surface kinetic measurements meaningful. We validate the mass-transfer behavior of the TIRFS cell by using a simple, nonadsorbing fluorescent probe, sodium fluorescein. (Interfacial tension measurements reported elsewhere (Tupy, 1998) demonstrate that sodium fluorescein adsorbs minimally at the O/W interface.) Then, the protein β-casein is studied using a covalently bound extrinsic fluorescent probe. Although our TIRFS apparatus is designed primarily to study protein adsorption at the O/W interface, the geometry of the spectroscopy cell can be applied to optical techniques other than fluorescence for the study of liquid/liquid interfaces.
Design. A schematic of the dynamic O/W TIRFS cell is shown in Figure 1. The body of the stainless steel cell contains the oil and aqueous phases. A quartz dove prism (CVI Laser, (9.0 × 6.5)-cm base, 1.5-cm height, 45° angle on the prism face) seals the top of the cell. A porous polymeric membrane (discussed below) separates the aqueous-phase portion of the cell into two parts, a stagnant layer and a flow channel. The membrane acts as a barrier to convective flow from the flow channel to the stagnant layer, but allows solute diffusive flux. Fluorescent-probe molecules under study are introduced to the TIRFS cell by circulating the appropriate aqueous solution in the flow channel. Surface-active fluorescent molecules approach the O/W interface by diffusing through the membrane and stagnant layer and adsorbing. Because the stagnant aqueous layer is only 0.1 cm in height and 2.3 cm in diameter at the O/W interface, use of the simple one-dimensional diffusion equation is permitted. As the time for diffusive transport scales with distance squared, the stagnant layer is designed with the smallest practical height. The membranes (Millipore RTTP04700) are 10-µm thick track-etched polycarbonate with a hydrophilic poly(vinylpyrrolidone) (PVP) surface coating, chosen because the straight-channel pore geometry minimizes the amount of surface area available for adsorption. A pore diameter of 1.2 µm was selected because a monolayer of adsorbed protein (nanometers thick) on the pore walls only marginally reduces the void volume. The oil-phase compartment, cylindrical in shape, is bounded by the prism, the water, and the stainless steel components. A Viton O-ring seals the quartz and stainless steel. As the oil is incompressible and is completely surrounded by impressible components, water can be circulated through the lower portion of the cell without changing the location of the O/W interface. A stainless steel knife-edge ring pins the contact line of the O/W interface and further reduces any movement of the interface during an experiment. The volume of the oil phase can be changed through the single port shown to the right in the figure. Prior to an experiment, oil is added or withdrawn through this port to force the interface to be flat, as discussed below. Closing a ball valve (Nupro SS-41S2) reseals this port and thus the oil-phase compartment, enabling aqueous solution to be circulated without altering the interface position. Because the interface has zero curvature and its contact line is pinned, subsequent changes in sorption and interfacial tension during an experiment do not alter the location of the interface.
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The flatness criterion for the O/W interface is determined by the depth of focus of the fluorescence-collection optics. Interfacial position must be retained within microns, a range that allows visual confirmation. By looking through the vertical side of the dove prism parallel to the plane of Figure 1, one can visually observe the O/W interface. Since the oil matches the index of refraction of the quartz, the interface is seen essentially distortion free. The angle at which the observer views the interface is greater than the critical angle, giving a flat O/W interface the appearance of a smooth mirror. Mounted vertically outside the cell, opposite to the observer, is graph paper with a fine grid spacing. When the interface is not flat, the image of the graph paper is magnified and the ruled lines in the reflected image appear distorted. By comparing the reflected image of the graph paper with the actual paper and by adjusting the oil-phase volume, the interface is reproducibly forced to be flat. Incident light is directed toward the cell parallel to the O/W interface. It enters the cell through the fusedsilica dove prism and travels toward the O/W interface at an angle greater than the critical angle. After reflection, a symmetric path guides the beam out of the cell. The oil is an aliphatic hydrocarbon mixture (Cargille Labs no. 50350) that matches the index of refraction of quartz over a wide range of wavelengths and was chosen to minimize reflections and scattering generated at the quartz/oil interface. The 45° angle on the prism face refracts the incident light such that the angle of incidence with the O/W interface is greater than the critical angle (nquartz(at 490 nm) ) noil(at 490 nm) ) 1.463 which gives φincidence ) 73.9 °> φcritical ) 65.4°). The reflection spot is the image of a 0.10-cm diameter aperture, not a collimated beam. As a result, the light reflects at the interface with a range of angles centered about 73.9°. Knowing the focusing-lens focal length and diameter (2.5 cm) and taking into account that the incident beam travels through media of different indices of refraction, the angle of incidence is calculated to be 69.6-77.8° over the area of reflection at the O/W interface. Details of the cell assembly are shown in Figure 2. The quartz prism is held in place with an aluminum clamp, four stainless steel screws, and a Buna-rubber pad to protect the prism from the clamp. Components assembled in the TIRFS cell basin are 5.5 cm in diameter, matching the basin dimensions. Two stainless steel disks, each with a sieved region in the center, sandwich the membrane and provide mechanical support. Sieve plates are 0.013-cm thick and have 0.051cm diameter holes drilled 0.076 cm apart on center over a region 2.5 cm in diameter in the center of the plate. A ring with a beveled inner edge is placed on the membrane supports and acts as a spacer for the stagnant layer, as well as the knife edge for pinning the O/W contact line. A stainless steel ring clamp with eight inset stainless steel screws secures and seals the assembly. The ring clamp has an inner diameter of 4.2 cm and is 0.38 cm in height, except directly below the light path where the ring is milled to 0.23 cm in height in order to not block the light. Beneath the assembly, the flow channel is located having dimensions of 2.5 × 2.5 × 0.1 cm. Those pieces in contact with the oil phase (the stainless steel ring clamp and the top of the knife-edge ring) are Teflon-coated to prevent water from wetting
Figure 2. Schematic of a vertical cross-section of the TIRFS cell assembly (drawn to scale). A: Aluminum clamp (thickness of 0.635 cm). B: Buna rubber pad (0.159 cm). C: Quartz dove prism (1.5 cm). D: Stainless steel ring clamp (0.38 cm). E: Stainless steel knife-edge ring (0.1 cm). F: Stainless steel sieve plate (0.0127 cm). G: Porous membrane (0.001 cm). H: Viton gasket (0.0508 cm). I: Viton O-ring (size 2-036). J: 0.67-cm deep by 5.5 cm-diameter cylindrical basin for pieces D-H. K: Stainless steel base. L: Flow lines.
the surfaces in the oil compartment and to inhibit weeping flow from the aqueous phase to the corners of the oil-phase compartment. The Teflon-coating procedure is detailed elsewhere (Tupy, 1998). A single coat, approximately 1-µm thick (Stewart, 1993), is reapplied after each experiment. The optical flow cell is the centerpiece of the overall TIRFS apparatus, which is illustrated in Figure 3. Light from a 150-W arc lamp (Oriel 6292) is focused onto a 0.10-cm aperture (Edmund Scientific J39,730), and collected and collimated using a plano-convex lens (Newport SPX022AR.10). It is then directed through a narrow band-pass filter (Oriel 59285 at 410 nm and Oriel 59335 at 490 nm), a polarizer (Oriel 27320), and a mechanical shutter (Melles Griot 4IES003). The polarizer is aligned parallel to the plane of incidence for all of the work described here. To protect the fluorescent probe from photobleaching, the shutter is open only during the short time (∼1 s) required to measure fluorescence intensity. A second lens (Newport SPX022AR.10) focuses the collimated light onto the interface. Collimating and focusing lenses have the same focal length (10.0 cm) to minimize chromatic aberration. Fluorescence generated in the cell is detected directly above the cell using two identical simple lenses (Oriel 41220) having a focal length of 2.5 cm. Collected fluorescence emission is focused into a monochromator (Acton SP-150-M) equipped with variablewidth entrance and exit slits. The intensity of the light exiting the monochromator is detected by a photomultiplier tube (PMT) (Hamamatsu R4632) and moni-
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Figure 3. Schematic of the O/W TIRFS apparatus.
tored in photon-counting mode (Hamamatsu C3866, Hewlett-Packard 53131A). To compensate for any drift of the arc-lamp intensity and other electronics over the course of an experiment, we take advantage of the light reflected off the face of the quartz prism at the air/quartz interface. A fiberoptic cable (Edmund Scientific M42,346), placed at the focal point of the reflected beam, directs the light to a second mechanical shutter (Newport 846HP) where a second fiber-optic cable (Edmund Scientific M42,346) transmits it to the PMT downstream of the monochromator. Cell fluorescence intensity is measured with this second shutter closed, and the light-source reference intensity is measured, in turn, with the shutter opened. The signal-referencing process is cycled several times per cell fluorescence-intensity interval, and the averages are saved. Both shutters, the monochromator, and the photon counter are all interfaced with a computer (IBM 386). To reduce perturbations resulting from room vibrations, the entire apparatus is mounted on a pneumatic vibration isolation table (Newport RS1000-48-8) and is isolated in a light-tight box. A piston pump (ISCO 100D) circulates the aqueous solutions through the optical flow cell. Technique. After assembly, more than 1000 flowchannel volumes of distilled, deionized water are flushed through the cell. Just preceding an experiment, the O/W interface is forced flat by adding or withdrawing oil through a hand-operated syringe, and the oil compartment is sealed. Aqueous solution containing the surface-active fluorescent probe is introduced to the cell by circulation through the flow channel. Probe molecules diffuse through the membrane and through the aqueous stagnant layer. As they near the O/W interface, light from the evanescent wave produced at the point of reflection is absorbed, and fluorescence is generated. The increase in fluorescence intensity is monitored over time. This process is referred to as loading. When the system is equilibrated or when a change in system conditions is desired, the aqueous solution concentration is changed by simply pumping a different solution composition through the flow channel. For example, to test for adsorption reversibility, a solution free of fluorescent surface-active agent is circulated in the flow channel. The decrease in fluorescence intensity at the interface is monitored as the fluorescent mol-
ecules desorb and diffuse away from the interface toward the flow channel to be flushed out of the cell. This process is referred to as washout. Fluorescent spectra are measured periodically during loading and washout. The monochromator diffraction grating is motor-driven and automated by computer control. Experimental Methods Materials. Crystalline sodium fluorescein, sodium azide (>99% purity), and β-casein (phosphorylated, greater than 90% purity by electrophoresis) were purchased from Sigma. Acetone, ethanol, dimethyl sulfoxide (DMSO), hydroxlamine, methanol, sodium bicarbonate, sodium phosphate dibasic, and sodium phosphate monobasic were obtained from Fisher at ACS-certified grade purity. Spectroscopic grade n-heptane was purchased from EM Science while sodium dodecyl benzene sulfonate (SDBS) was obtained from TCI. 1-(3-(succinimidyloxycarbonyl)benzyl)-4-(5-(4-methoxyphenyl)oxazol2-yl)pyridinium bromide (PyMPO-SE) from Molecular Probes served as the fluorescent protein label. The Teflon-coating solution was supplied by Dupont (“Teflon” solution TE5078A). All chemicals were used without further purification. Water was distilled and deionized with a four-stage Milli-Q reagent-grade water system from Millipore to a routine surface tension of over 72 mN/m. All experiments were performed at room temperature, 21 ( 1 °C. β-Casein was labeled with PyMPO-SE per the instructions provided by the manufacturer. A 20-cm3 solution of 1.0 mg/cm3 β-casein in 0.1 M sodium bicarbonate was prepared and combined with a 0.185-cm3 aliquot of a 1.0 mg/ cm3 solution of PyMPO-SE dissolved in DMSO to give a 4:1 molar ratio of PyMPO to β-casein. The reaction mixture was gently stirred at room temperature for 2 h, after which the reaction was stopped by adding 1.5 M hydroxylamine (pH 6.7) to bring the solution to a final concentration of 0.15 M. The solution was then gently stirred at room temperature for an additional hour. To separate the conjugates from the free dye, the reaction mixture was dialyzed eight times in succession against 0.1 M, pH 7.0 phosphate buffer in a light-free environment. Protein concentration of the conjugate solution was determined by the BioRad Coumassie Blue Protein Assay per instructions of the
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manufacturer. PyMPO concentration was measured by absorbance using a Beckman DU-6 spectrometer and an extinction coefficient of ) 26 000 M-1 cm-1 at 415 nm (Haugland, 1996). The labeling ratio was determined to be 2.4 PyMPO probes per protein. Conjugate solution was stored at 4 °C in a light-free environment. Bulk-solution spectra, measured using the TIRFS apparatus by replacing the TIRFS cell with a standard optical cuvette, confirm that the spectral response of the TIRFS apparatus is consistent with that measured by a standard fluorescence spectrometer (Perkin-Elmer LS3). Procedure. Dynamic TIRFS experiments with nonadsorbing fluorescein salt are carried out in the presence of 1.44 mM sodium dodecyl benzene sulfonate (SDBS) to reduce adsorption of fluorescein onto the polymeric membranes and onto the O/W interface. Loading experiments begin by circulating aqueous 0.01 mg/cm3 fluorescein solution in 1.44 mM SDBS through the flow channel until equilibrium is reached. At this time, washout experiments begin by introducing fluoresceinfree 1.44 mM SDBS solution. This solution is circulated in the flow channel until the fluorescence intensity returns to its initial value. Excitation light is 490 nm selected with a band-pass filter (10 nm band-pass), and fluorescence emission is measured at 520 nm (10 nm band-pass). Dynamic TIRFS experiments with β-casein conjugates are carried out in pH 7.0, 0.1 M sodium phosphate buffer with 2 mM sodium azide added as a bacteriocide. The solution has a 0.1 mg/cm3 protein concentration with 10% of the protein labeled and the remainder native protein. Washout is performed by flushing with proteinfree phosphate buffer solution. Excitation light is centered at 410 (10 nm band-pass), and fluorescence emission is measured at 550 nm (10 nm band-pass). TIRF spectra are measured periodically during loading and washout to monitor changes in time of the chemical environment of the fluorescent marker in the adsorbed protein layer. Only the final loading and final washout spectra are reported here to demonstrate the capabilities of the apparatus. Mass-Transfer Characterization To validate the mass-transport behavior of the TIRFS cell, fluorescence-intensity dynamics are followed for a simple, nonadsorbing solute, sodium fluorescein. The first step is to account for the dead time needed to flush the system with a new solution. This is done by installing a nonporous membrane and determining at each flow rate the time for the exiting solution fluorescence to reach 95% of the inlet concentration. We take one-half of this value as the dead time with typical values of a few minutes in our apparatus. Next, loading and washout experiments are performed with the nonadsorbing sodium fluorescein. Figure 4 shows examples of loading and washout TIRFS data for sodium fluorescein in the aqueous SDBS solution at flow rates of 0.054 and 0.83 cm3/min. Relative fluorescence intensity is plotted as a function of time. Note that at early times the loading and washout data both do not change. This time lag is a result of the finite thickness of the stagnant layer. A short lag time is required for the fluorophore to diffuse across the stagnant layer, reach the interface, and interact with the evanescent wave. Also, notice that the
Figure 4. TIRFS loading and washout intensity histories for 0.01 mg/cm3 sodium fluorescein in 1.44 mM SDBS solution at channel flow rates of 0.054 and 0.83 cm3/min. The best fit of the masstransport model is shown by the superimposed heavy lines.
time scales for the loading and washout steps are the same, confirming that any convective flux of material in the stagnant layer is indeed negligible. A flow-rate dependence of the TIRFS dynamics is clearly demonstrated in Figure 4. Higher flow rates lead to faster loading of the stagnant layer. A similar trend is found for the washout curves with higher flows, leading to faster depletion. A mass-transfer resistance at the flow-channel membrane boundary explains this observed flow rate dependence. Solid lines, shown superimposed on the data in Figure 4, reflect a one-dimensional diffusion model with a uniform initial concentration, a no-flux boundary condition at the O/W interface, and a flux boundary at the channel membrane characterized by a convective masstransfer resistance in the flow channel at the membrane boundary. (Model details are given in Appendix A.) Transient concentration profiles (eq A-6) are integrated over the region illuminated by the evanescent wave in obedience to eqs 1 and 2. The quantum yield and extinction coefficient are taken as constants for the nonadsorbing fluorescein and are combined into the instrument parameter. The diffusive path length, hD ) 0.15 cm, the channel length, L ) 2.5 cm, and the channel height, hF ) 0.15 cm, are set by the TIRFS-cell geometry (Tupy, 1998). A known diffusion coefficient for fluorescein in water, D ) 6.3 × 10-6 cm2/s (Zero et al., 1983), is adopted. Adsorption onto the sieve plate and membrane surfaces is neglected, and the diffusivity in the pores is assumed to be that in bulk water. The mass-transfer coefficient, km, is used as the single-fitting parameter by minimizing the square difference of the diffusion model and the experimental data for a given flow rate. Comparison of the mass-transfer model and data in Figure 4 demonstrate excellent agreement. Figure 5 displays mass-transfer coefficients regressed from the data in Figure 4 along with loading and washout experiments performed at eight additional flow rates. Values of the mass-transfer coefficients, reported as Nusselt numbers, Nu ) kmhF/D, are plotted as a function of flow rate in terms of the Graetz number, Gr ) 2〈v〉hF2/DL, where 〈v〉 is the average fluid velocity. The solid line is the solution to the Le´veˆque problem in a channel (Edwards and Newman, 1985) and is in excellent agreement with the experimental data over a range
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Figure 5. Local Nusselt number plotted as a function of the Graetz number for measured mass-transfer coefficients at the membrane in the flow channel of the O/W TIRFS cell. The result for the Le´veˆque problem in a channel is also shown for comparison. At Gr g 200 Starling flow is seen in the stagnant layer.
greater than an order of magnitude in Gr (5-200), corresponding to flow rates of 0.04-1.6 cm3/min for fluorescein solutions. At small values of the Graetz number, which reflect low flow rates or long channels, the Le´veˆque solution breaks down due to the assumption of a linear velocity profile. At large values of Gr, the mass transport is complicated by convection through the membrane and into the stagnant layer or Starling flow (Apelblat et al., 1974). Appendix B gives an order of magnitude estimate to predict the onset of Starling flow, as demarked in Figure 5. Thus, mass-transfer coefficients determined at higher flow rates deviate from the Le´veˆque solution, confirming the initiation of Starling flow. Below this critical flow rate, the proposed diffusion model accurately describes mass transfer in the TIRFS cell. Additionally, fluorescein concentrations were varied from 0.003 to 0.03 mg/cm3 to assess any possible fluorescein adsorption loss to the membrane or cell walls. In the presence of the dilute surfactant SDBS, no effect of fluorescein concentration was found. Accordingly, Figures 4 and 5 confirm that solute mass transfer in our optical flow cell is indeed by onedimensional diffusion through a stagnant aqueous layer. It is now possible to apply the diffusion model toward elucidation of protein sorption dynamics. β-Casein/PyMPO Sorption TIRF Intensity Dynamics. Figure 6 shows TIRFS experimental results for loading (open circles) and washout (filled triangles) intensity curves for PyMPO/ β-casein conjugate sorption at a channel flow rate of 0.3 cm3/min. Arrival of 0.1 g/cm3 of β-casein/PyMPO conjugate at the interface is monitored over a 24-h period. The lag at early times for β-casein is about 40 min, longer than that observed for fluorescein due to a smaller diffusion coefficient and possibly due to some adsorption of protein onto the membrane and other components of the apparatus. Washout is performed with protein-free phosphate buffer circulating in the flow channel, again at a flow rate of 0.3 cm3/min. After 30 h of washout there is little change in the TIRF intensity, although sufficient time has clearly elapsed for any reversibly adsorbed material
Figure 6. TIRF loading (open symbols) and washout (filled symbols) intensity histories for β-casein/PyMPO conjugate adsorption at the O/W interface from a 0.1 mg of protein/cm3 solution at a 0.3 cm3/min channel flow rate. Results for the irreversible Langmuir adsorption kinetic model are shown in the heavy lines for adsorption rate constants of k1 ) 10, 1, and 0.1 cm3/g/s.
to diffuse away. Thus, following 24 h of exposure to the O/W interface, β-casein is essentially irreversibly adsorbed. To quantify the measured protein adsorption dynamics, the following simple model is proposed. Transport in the stagnant layer is by one-dimensional diffusion with zero initial concentration and a mass-transfer-flux boundary condition at the membrane boundary, as in the nonadsorbing solute case. Protein loading at the O/W interface, however, is assumed to obey simple, irreversible Langmuir adsorption kinetics:
dΓ(t) ) k1c(t,0) (Γsat - Γ) dt
(3)
where Γ is the adsorption, Γsat is the saturation adsorption, c(t,0) is the protein concentration in the solution sublayer, and k1 is the adsorption rate. Equation 3 serves as the boundary condition at the O/W interface. Details of the mathematical model are outlined in Appendix C. It is interesting to note the difference between this diffusion/adsorption model and the commonly applied Ward-Tordai analysis (1946). In the Ward-Tordai model, a clean interface at zero time interacts with a solution of uniform concentration, whereas in the TIRFS apparatus, the solute must first diffuse across the stagnant layer in order to adsorb, resulting in a lag time, as observed in the TIRFS data. This difference also appears in that the characteristic diffusion length for the TIRFS experiments, hD, the thickness of the stagnant layer, while in the WardTordai model the adsorption length, Γsat/c, is the appropriate characteristic diffusion length scale. Results of the irreversible Langmuir kinetic model are shown as solid lines in Figure 6 in terms of the fractional adsorption, θ ≡ Γ/Γsat, for various values of the adsorption rate constant. Fractional adsorption, θ, is compared directly to normalized interfacial fluorescence intensities. Thus, the maximum fluorescence, F ) 1, corresponds to Γ ) Γsat. In the model-predicted intensities, we do not include any contribution from the protein concentration below the surface because the adsorbed protein dominates the TIRF signal with little contribution from the bulk, as demonstrated by the
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washout data in Figure 6. As with the sodium fluorescein experiments, the quantum yield and extinction coefficients are assumed constant. Adsorption of proteins onto the apparatus surfaces is neglected in the modeling because the amount of protein lost to the various surfaces of the TIRFS cell has a minor effect at the bulk concentration used here. Remaining parameters in the model are determined by experimental conditions. The diffusivity, D, of β-casein is 6.05 × 10-7 cm2/s (Sullivan et al., 1955), and the mass-transfer coefficient is calculated using the Le´veˆque solution, eq A-5, for a channel flow rate of 0.3 cm3/min or an average fluid velocity of 〈v〉 ) 0.013 cm/s. Maximum β-casein adsorption, Γsat ) 4 mg/m2, is taken from that measured at a toluene/water interface by Graham and Phillips (1979). As seen in Figure 6, an adsorption rate constant of 1 cm3/g/s gives good agreement with the loading data. The square root of the dimensionless adsorption rate constant, κ ≡ k1ΓsathD/D, is equivalent to the Thiele modulus for this problem and has a value of 0.3, indicating that the system is surface adsorption ratelimiting. The slow arrival of β-casein at the interface by diffusion is still faster than the rate at which β-casein can reconfigure into the adsorbed layer. The assumption of constant and ψ is likely in error as the β-casein strongly adsorbs at the O/W interface, forcing the PyMPO into an environment that is different than that in bulk water and that likely varies within the adsorbed-layer region. Additionally, the adsorbed layer is more densely packed with protein than is the bulk phase, providing a higher probability of fluorescence quenching. Lacking a detailed description of the chemical nature of the β-casein adsorbed layer, we take and ψ as constants. The TIRFS-loading curve displays a small maximum at about 1.5 h. This maximum could be a result of a number of phenomena including photobleaching, rotation of the flourescence excitation dipole, competitive adsorption between labeled and unlabeled β-casein, concentration quenching, and structural rearrangements in the adsorbed layer (Robeson and Tilton, 1996). The most likely explanation is quenching, although structural rearrangements in the adsorbed layer and interactions of the PyMPO label with the oil phase could also lead to a maximum in dynamic TIRFS (Tupy, 1998). TIRF Spectra. To gain understanding of the fluorescent-probe molecular environment at the O/W interface, we periodically collect fluorescence emission spectra. Figure 7 displays such spectra measured after the 24-h loading step and again after the 30-h washout step reported in Figure 6. The loading and washout TIRF spectra exhibit fluorescence-emission maxima at wavelengths of 528 and 526 nm, respectively. For comparison, the bulk-solution spectrum measured for a protein concentration of 1.0 mg/cm3 is also shown. At the end of loading, the TIRF spectrum is blueshifted by 27 nm from the bulk, indicating that the PyMPO fluorescent probe in the adsorbed β-casein layer is in a less polar environment than that in the aqueous solution (Lakowicz, 1983). A blue shift of 29 nm is measured in the TIRFS-washout spectrum, again signifying a greater contribution to fluorescence from PyMPO molecules in a nonpolar environment. During washout, those PyMPO molecules in a polar environment either diffuse away from the interface and are washed from the cell, or they become incorporated into the adsorbed layer, thereby moving into a less polar region. Both processes
Figure 7. TIRF spectra after 24-h loading and subsequent 30-h washout steps for the irreversibly adsorbed β-casein/PyMPO conjugate from a 0.1 mg of protein/cm3 solution. The bulk-solution spectrum of the β-casein/PyMPO conjugate is shown for comparison.
result in a slight shift of the washout TIRF spectrum to the blue, indicative of greater interaction of adsorbed β-casein with the oil phase. Discussion β-Casein is a surface-active protein that is an important agent in stabilizing milk emulsions (Dickinson, 1994). The amphiphilic nature of the molecule suggests fast adsorption kinetics at a clean O/W interface. However, the experimentally observed Thiele modulus of 0.3 and the long time frame in Figure 6 indicate slow incorporation of β-casein into the adsorbed layer at a rate less than that of the diffusive flux. Simple kinetic reorientation of the adsorbed β-casein cannot be the explanation as all tagged molecules fluoresce within the penetration depth, dp, independent of orientation. The negligible desorption found after long residence times at the interface demonstrates strong interaction within the adsorbed protein layer. Blue shifts in the TIRF spectra evidence exposure of the fluorescent probes to a more oil-like environment as the adsorption process continues. During the slow, irreversible incorporation into the adsorbed layer, β-casein molecules progressively expose their PyMPO tags to a hydrophobic environment. These observations may be reconciled with the following simplified description of the adsorption process. Native protein molecules arriving at the clean O/W interface adsorb quickly. They then start to reconfigure and allow their hydrophobic moieties to interact with the oil phase. Concomitantly, more β-casein molecules diffuse to the interface, thereby building the sublayer concentration and the adsorbed amount and raising the TIRF intensity. These molecules also begin to unfold and interact both with the oil phase and with the previously adsorbed protein molecules. This causes the blue shifts seen in the TIRF spectra. The adsorbed region grows in thickness, and the adsorbed molecules slowly intermesh into an entangled network structure that apparently does not permit desorption. Thus, the picture of β-casein adsorption at the O/W interface is one of inexorable denaturation and eventual building of gel-like layers. This picture is consistent with previous observations of irreversible protein adsorption at the O/W boundary (Cumper and Alexander, 1951; Cheesman and Davies, 1954; Izmailova, 1979; Beverung, 1996).
3166 Ind. Eng. Chem. Res., Vol. 37, No. 8, 1998
Utility of the TIRFS Cell
Nomenclature
The optical flow cell developed here for TIRFS study of dynamic adsorption at a liquid/liquid interface can be directly applied to other optical techniques. Fluorescence methods that are more sensitive to molecular configuration, such as tryptophan fluorescence and fluorescence resonance energy transfer, require little additional effort. Raman spectroscopy is now possible to probe molecular conformation changes during transient adsorption processes. As noted in the Introduction, reflection techniques, such as ATR-FTIR, CD, ellipsometry, reflectometry, and surface light scattering, have been previously applied to fluid interfaces and now may be extended to the study of sorption dynamics.
c(t,z) ) dimensional concentration (g/cm3) C(τ,Z) ) dimensionless concentration, chD/Γsat D ) diffusion coefficient (cm2/s) dp ) depth of penetration for the evanescent wave (nm) E(t,z) ) evanescent-wave intensity F(t) ) TIRF intensity Gr ) Graetz number, 2〈v〉hF2/DL hD ) diffusive path length (cm) hF ) height of the flow channel (cm) hS ) height of the stagnant layer (cm) k1 ) dimensional rate constant for adsorption (cm3/g/s) km ) dimensional mass-transfer coefficient (cm2/s) L ) length of the flow channel (cm) Lp ) membrane hydraulic permeability (cm2 s/g) n1, n2, noil, nquartz ) indices of refraction N ) dimensionless mass-transfer coefficient, kmhD/D Nu ) Nusselt number defined in eq A-5 ∆P ) pressure drop (dyn/cm2) Pe ) Peclet number defined in Appendix B Q ) channel flow rate (cm3/min) t ) time (s) V(τ,Z) ) dimensionless concentration defined in eq C-6 vm ) maximum fluid velocity in stagnant layer (cm/s) 〈v〉 ) average fluid velocity in the flow channel (cm/s) W ) width of the flow channel (cm) z ) dimensional distance from the O/W interface (cm) Z ) dimensionless distance from the O/W interface, z/hD
Conclusions A novel total internal reflection fluorescence spectrometer (TIRFS) has been constructed to study adsorption dynamics at the O/W interface. The unique design of our apparatus allows introduction and removal of surface-active species without mechanically perturbing the interface and also permits long-time adsorption phenomena to be examined, even as interfacial tensions change. Nonadsorbing solute transport in the TIRFS cell is well-described by a simple one-dimensional diffusion equation, as confirmed by transport studies with 0.01 mg/cm3 sodium fluorescein in 1.44 mM SDBS. The observed flow-rate dependence of the TIRF intensities is accurately described by the Le´veˆque mass-transfer problem in a channel over a range of Graetz numbers from 5 to 200, corresponding to flow rates of 0.04-1.6 cm3/min for aqueous sodium fluorescein solution concentrations between 0.003 and 0.03 mg/cm3. β-Casein/PyMPO conjugate adsorption is studied in 0.1 M, pH 7.0 phosphate buffer at a 0.1 mg/cm3 concentration and at a flow rate of 0.3 cm3/min. TIRF intensity monitored for 24 h of loading and a subsequent 30 h of washout displays no perceptible desorption, signifying the irreversible and entangled nature of β-casein adsorbed at O/W interfaces. The TIRF loading history exhibits best agreement with the Langmuir irreversible adsorption kinetic model when the adsorption rate constant is near 1 cm3/g/s. A Thiele modulus of 0.3 emerges, indicating slow kinetic-controlled reconfiguration of β-casein into the adsorbed layer. TIRF spectra measured after the completion of loading and washout evidence fluorescence-emission maximum wavelengths of 528 and 526 nm, respectively, corresponding to blue shifts of 27 and 29 nm with respect to the bulksolution spectrum. Adsorbed β-casein molecules interact both with the nonpolar oil phase and neighboring adsorbed proteins. The adsorption process for β-casein at the O/W interface is apparently one of denaturation and slow entanglement into a gel-like domain. The new TIRFS apparatus follows adsorption and desorption dynamics quantitatively and garners information on molecular changes in the adsorbed layer. Further, the new flow-cell design can be applied to different fluorescence studies, as well as to other lightbased techniques for probing liquid/liquid interfacial phenomena.
Subscripts 0 ) initial condition ∞ ) concentration far from the interface sat ) saturation adsorption Greek Symbols χ ) spectrometer instrument parameter (t,z) ) extinction coefficient (cm3/g/cm) φ ) angle (deg) Γ ) adsorption (mg/m2) κ ) dimensionless adsorption-rate coefficient or the square of the Thiele modulus, k1ΓsathD/D λ ) wavelength of light (nm) λn ) eigenvalue for the series solution U(τ,Z) µ ) viscosity (g/cm/s) µn ) eigenvalue for the series solution for the step change boundary condition within the Duhamel superposition integral νn ) eigenvalue for the asymptotic solution in eq C-7 θ ) fractional adsorption, Γ/Γsat τ ) dimensionless time, Dt/hD2 ψ(t,z) ) quantum yield
Appendix A: A Diffusion Model for Nonadsorbing Fluorophores In this appendix we describe the mass-transport model of our optical flow cell for a nonadsorbing solute. The one-dimensional diffusion equation reads
∂c ∂2c )D 2 ∂t ∂z with the initial and boundary conditions of
c(0,z) ) c0
Acknowledgment The work was funded by the U.S. Department of Energy, Basic Energy Sciences under Grant No. DEF903-94ER14456.
(A-1)
D
∂c(t,hD) ) km(c∞ - c) ∂z
(A-2) (A-3)
Ind. Eng. Chem. Res., Vol. 37, No. 8, 1998 3167
and
∂c(t,0) )0 ∂z
(A-4)
where c(t,z) is the local concentration of the fluorescing species, t is the time, D is the diffusion coefficient, z ) 0 locates the O/W interface, and z ) hD corresponds to the side of the membrane facing the flow channel. Thus, hD is the diffusion path length. The initial concentration is c0 and the concentration in the flow channel far from the membrane is c∞. To account for the observed flow rate dependence on the loading/washout curves in Figure 4, we adopt a mass-transfer coefficient, km, from the Le´veˆque problem in a channel with mass transfer on one side of the channel (Edwards and Newman, 1985), given in terms of the Nusselt number:
Nu )
kmhF ) 0.9244Gr1/3 - 0.2 - 0.10025Gr-1/3 D (A-5)
where Gr is the Graetz number defined in the text. Solutions to eqs A-1-A-4 is straightforward:
c(t,z) - c∞ c0 - c∞
∞
)
∑ n)1
2λn sin(λn)
λn2
( )
drop across the membrane. The Hagen-Poiseuille law in a slit is applied to calculate ∆Pflow channel. From Darcy’s law, the Hagen-Poiseuille law, and the definition of the Peclet number, we obtain
Pe )
where N ) kmhD/D and the eigenvalues, λn, are determined by λn tan(λn) ) N. The solution is valid provided the time scale associated with the mass transfer in the flow channel is much shorter than that in the stagnant layer and provided that there is no convective flow in the stagnant layer.
Appendix C: A Diffusion-Langmuir Kinetic Model for Adsorbing Fluorophores A diffusion model with Langmuir irreversible adsorption kinetics is outlined in this appendix. In reduced variables we have that
∂C ∂2C ) 2 ∂τ ∂Z
(C-1)
with the initial condition
C(0,Z) ) 0
(C-2)
boundary condition of
∂C(τ,1) ) N(C∞ - C) ∂Z
Appendix B: Starling Flow Estimate At higher flow rates, Starling flow (Apelblat et al., 1974) commences in the stagnant layer by hydrodynamic convection through the membrane. Flow lines travel from the flow-channel inlet through the membrane, along the stagnant layer, and back down through the membrane into the flow channel. Starling flow has a much higher resistance than that directly along the flow channel, but the resistance is finite. At higher channel flow rates, the convective flux due to Starling flow becomes comparable to the diffusive flux of material through the membrane, and the simple one-dimensional diffusion equation no longer applies. To estimate when Starling flow becomes important, we examine the magnitude of the Peclet number in the stagnant layer, Pe ) vmhS/D, where hS is the height of the stagnant layer and vm is the fluid velocity in the membrane pores. Small Pe means that convective flux due to Starling flow may be neglected compared to that due to diffusive flux. The velocity, vm, is estimated from Darcy’s law (vm ) Lp∆Pmembrane) using measured values of the membrane hydraulic permeability provided by the manufacturer (Lp ) 3.3 × 10-6 cm2s/g) and an estimate of the pressure drop across the membrane, ∆Pmembrane. We adopt a conservative estimate of the pressure drop across the membrane as ∆Pmembrane ) ∆Pflow channel/2, where ∆Pflow channel is the pressure drop across the flow channel. Implicit in this estimate are the assumptions that Starling flow is negligible compared to the flow in the channel and that the pressure drop across the stagnant layer is negligible compared to the pressure
(B-1)
DhF3W
where hS ) 0.10 cm, Q is the flow rate in the flow channel, and µ is the viscosity of water. The length, width, and height dimensions of the flow channel are known: L )2.5 cm, W ) 2.5 cm, and hF ) 0.15 cm, respectively. When Pe > 0.1, convective flux is large enough that the simple diffusion models described in this paper no longer apply. This upper bound estimate is labeled in Figure 5 and agrees with the experimental observation of Starling flow in our flow cell.
z (-λn2Dt)/(hD2) cos λn e hD + N cos (λn) (A-6) 2
12hSLpQµL
(C-3)
and adsorption kinetics at the O/W boundary of
∂θ(τ) ∂C(τ,0) ) ) κC(1 - θ) ∂τ ∂Z
(C-4)
where C ) chD/Γsat, τ ) Dt/hD2, Z ) z/hD, N ) kmhD/D, θ ) Γ/Γsat, and κ ) k1ΓsathD/D. Solution to eqs C-1C-3 with a time-dependent boundary condition at Z ) 0 is obtained by invoking superpostion:
C(τ,Z) )
[
∞
C∞ 1 -
]
( )
2λn sin(λn)
z (-λn2Dt)/(hD2) cos λn e + hD λn2 + N cos2(λn) V(τ,Z) (C-5)
∑ n)1
and by applying Duhamel’s superposition integral (Hildebrand, 1962):
V(τ,Z) ) ∞
[
∂V(τ′,0)
∫0τ
∑ n)1
∂τ′
1-
N
ZN+1
2N
µn(N + cos (µn))
]
sin(µnZ)e-µn (τ-τ′) dτ′ (C-6) 2
2
Eigenvalues, λn, are the same as those defined in Appendix A, while the eigenvalues, µn, are determined by N tan(µn) ) -µn. To solve for V(τ,0), the adsorption boundary condition in eq C-4 is invoked, and the
3168 Ind. Eng. Chem. Res., Vol. 37, No. 8, 1998
equations are solved numerically for θ(t) according to a fully implicit, forward time marching scheme, as summarized by Acrivos and Chambre` (1957). Nonlinearities are resolved at each time step by Newton-Raphson iteration. Additional mathematical details are available in the thesis of Tupy (1998). Literature Cited Acrivos, A.; Chambre, P. L. Laminar Boundary Layer Flows with Surface Reactions. Ind. Eng. Chem. 1957, 49, 1025. Apelblat, A.; Katzir-Katchalsky, A.; Silberberg, A. A Mathematical Analysis of Capillary-Tissue Fluid Exchange. Biorheology 1974, 11, 1. Bessho, K.; Uchida, T.; Yamauchi, A.; Shioya, T.; Teramae, N. Microenvironments of 8-Anilino-1-Naphthalenesulfonate at the Heptane-Water Interface: Time-Resolved Total Internal Reflection Fluorescence Spectroscopy. Chem. Phys. Lett. 1997, 264, 381. Beverung, C. J. Dynamics of Protein and Polypeptide Adsorption at the Oil/Water Interface. M.S. Dissertation, University of California, Berkeley, CA, 1996. Burghardt, T. P.; Axelrod, D. Total Internal Reflection/Fluorescence Photobleaching Recovery Study of Serum Albumin Adsorption Dynamics. Biophys. J. 1981, 33, 455. Cheesman, D. F.; Davies, J. T. Physicochemical and Biological Aspects of Proteins at Interfaces. In Advances in Protein Chemistry; Anson, M. L., Bailey, K., Edsall, J. T., Eds.; Academic Press: New York, 1954. Conboy, J. C.; Messmer, M. C.; Richmond, G. L. Investigation of Surfactant Conformation and Order at the Liquid-Liquid Interface by Total Internal Reflection Sum-Frequency Vibrational Spectroscopy. J. Phys. Chem. 1996, 100, 7617. Cumper, C. W. N.; Alexander, A. E. Proteins at Interfaces. Rev. Pure Appl. Chem. 1951, 1, 121. Dickinson, E. Emulsion Stability. In Food Hydrocolloids: Structures, Properties, and Functions; Nishinari, K., Doi, E., Eds.; Plenum Press: New York, 1994; p 387. Edwards, V.; Newman, J. The Asymmetric Graetz Problem in Channel Flow. Int. J. Heat Mass Transfer 1985, 28, 503. Graham, D. E.; Phillips, M. C. Proteins at Liquid Interfaces. II. Adsorption Isotherms. J. Colloids Interface Sci. 1979, 70, 415. Grubb, S. G.; Kim, M. W.; Rasing, T.; Shen, Y. P. Orientation of Molecular Monolayers at the Liquid/Liquid Interface as Studied by Optical Second Harmonic Generation. Langmuir 1988, 4, 452. Harrick, N. J. Internal Reflection Spectroscopy; Interscience Publishers: New York, 1967. Harrick, N. J.; Loeb, G. I. Multiple Internal Reflection Fluorescence Spectrometry. J. Anal. Chem. 1973, 33, 687. Haugland, R. P. Handbook of Fluorescent Probes and Research Chemicals; Molecular Probes, Inc.: Eugene, OR, 1996; Data Table 1.7. Higgins, D. A.; Corn, R. M. Second Harmonic Generation Studies of Adsorption at a Liquid-Liquid Electrochemical Interface. J. Phys. Chem. 1993, 97, 489. Hildebrand, F. B. Advanced Calculus for Applications; PrenticeHall: Englewood Cliffs, NJ, 1962; p 461. Izmailova, V. N. Structure Formation and Rheological Properties of Proteins and Surface-Active Polymers of Interfacial Adsorption Layers. In Progress in Surface and Membrane Science; Cadenhead, D. A., Danielli, J. F., Eds.; Academic Press: New York, 1979; p 141. Lakowicz, J. R. Principles of Fluorescence Spectroscopy; Plenum Press: New York, 1983; p 189. Liebmann, L. W.; Robinson, J. A.; Mann, K. G. A Dual Beam Total Internal Reflection Fluorescence Spectrometer for Dynamic Depth Resolved Measurements of Biochemical Liquid-Solid Interface Binding Reactions in Opaque Solvents. Rev. Sci. Instrum. 1991, 62, 2083.
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Received for review December 15, 1997 Revised manuscript received April 9, 1998 Accepted April 9, 1998 IE9709244
