Accumulation and Separation of Membrane-Bound Proteins Using

Dec 14, 2010 - The separation of molecules residing in the cell membrane remains a largely unsolved problem in the fields of bioscience and biotechnol...
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Anal. Chem. 2011, 83, 604–611

Accumulation and Separation of Membrane-Bound Proteins Using Hydrodynamic Forces Peter Jo¨nsson,* Anders Gunnarsson, and Fredrik Ho¨o¨k* Department of Applied Physics, Chalmers University of Technology, SE-41296 Gothenburg, Sweden The separation of molecules residing in the cell membrane remains a largely unsolved problem in the fields of bioscience and biotechnology. We demonstrate how hydrodynamic forces can be used to both accumulate and separate membrane-bound proteins in their native state. A supported lipid bilayer (SLB) was formed inside a microfluidic channel with the two proteins streptavidin (SA) and cholera toxin (CT) coupled to receptors in the lipid bilayer. The anchored proteins were first driven toward the edge of the lipid bilayer by hydrodynamic forces from a flowing liquid above the SLB, resulting in the accumulation of protein molecules at the edge of the bilayer. After the concentration process, the bulk flow of liquid in the channel was reversed and the accumulated proteins were driven away from the edge of the bilayer. Each type of protein was found to move at a characteristic drift velocity, determined by the frictional coupling between the protein and the lipid bilayer, as well as the size and shape of the protein molecule. Despite having a similar molecular weight, SA and CT could be separated into monomolecular populations using this approach. The method also revealed heterogeneity among the CT molecules, resulting in three subpopulations with different drift velocities. This was tentatively attributed to multivalent interactions between the protein and the monosialoganglioside GM1 receptors in the lipid bilayer. The cell membrane consists of a fluid lipid bilayer in which a variety of other membrane-associated molecules, such as proteins and oligosaccharides, are embedded. In the effort to understand the complex biochemical reactions orchestrated by the cell membrane, significant research has been devoted to isolating membrane-associated molecules. In contrast to water-soluble biomolecules, for which chromatographic methods have long been established,1,2 the separation and identification of membraneassociated proteins is a significantly more demanding task. The main problem associated with membrane protein separation and isolation with existing methods is that the molecules must be extracted from the lipid bilayer using detergents before they can be purified by conventional chromatography, a procedure that can * Corresponding authors. E-mail: (P.J.) [email protected]; (F.H.) [email protected]. (1) Hage, D. S. Clin. Chem. 1999, 45, 593–615. (2) Issaq, H. J.; Conrads, T. P.; Janini, G. M.; Veenstra, T. D. Electrophoresis 2002, 23, 3048–3061.

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damage or alter the structure and function of the molecules.3-5 A method that allows membrane-associated molecules to be separated while still being embedded in their native lipid environment would, therefore, be of great value in the field of cell membrane proteomics. Methods of separating membrane-associated molecules have so far primarily utilized electric fields to drive charged molecules in a supported lipid bilayer (SLB), which is a common cell membrane mimic consisting of a planar lipid bilayer formed on a solid support.6-8 This has allowed the separation of different types of charged lipids9-12 as well as membrane-anchored proteins.13,14 An alternative approach that does not rely on the charge of the molecules is the self-spreading approach,15-17 in which a dried aggregate of lipids is hydrated, followed by spontaneous spreading of an SLB over the support. Separation of different lipids was shown to occur when an SLB spread through an array of metal nanogates.18 Although this technique can be used to move uncharged molecules, it is not possible to control the drift velocity or the direction of motion of the SLB and its components. In addition to these techniques, surface acoustic waves have also been used to separate membrane-associated molecules, where the molecules were observed to accumulate at either the nodes or antinodes of the standing wave.19,20 We recently discovered that it is possible to instigate and control the transport of an SLB formed on the walls of a microfluidic channel by applying a bulk flow of liquid inside the (3) (4) (5) (6) (7) (8) (9) (10) (11) (12) (13) (14) (15) (16) (17) (18) (19) (20)

Lundstrom, K. Cell. Mol. Life Sci. 2006, 63, 2597–2607. Kashino, Y. J. Chromatogr. B 2003, 797, 191–216. Josic, D.; Clifton, J. G. Proteomics 2007, 7, 3010–3029. Sackmann, E. Science 1996, 271, 43–48. Castellana, E. T.; Cremer, P. S. Surf. Sci. Rep. 2006, 61, 429–444. Knoll, W.; Koper, I.; Naumann, R.; Sinner, E. K. Electrochim. Acta 2008, 53, 6680–6689. van Oudenaarden, A.; Boxer, S. G. Science 1999, 285, 1046–1048. Daniel, S.; Diaz, A. J.; Martinez, K. M.; Bench, B. J.; Albertorio, F.; Cremer, P. S. J. Am. Chem. Soc. 2007, 129, 8072–8073. Groves, J. T.; Boxer, S. G.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 935–938. Stelzle, M.; Miehlich, R.; Sackmann, E. Biophys. J. 1992, 63, 1346–1354. Groves, J. T.; Wulfing, C.; Boxer, S. G. Biophys. J. 1996, 71, 2716–2723. Tanaka, M.; Hermann, J.; Haase, I.; Fischer, M.; Boxer, S. G. Langmuir 2007, 23, 5638–5644. Nabika, H.; Takimoto, B.; Murakoshi, K. Anal. Bioanal. Chem. 2008, 391, 2497–2506. Nissen, J.; Gritsch, S.; Wiegand, G.; Radler, J. O. Eur. Phys. J. B 1999, 10, 335–344. Radler, J.; Strey, H.; Sackmann, E. Langmuir 1995, 11, 4539–4548. Nabika, H.; Iijima, N.; Takimoto, B.; Ueno, K.; Misawa, H.; Murakoshi, K. Anal. Chem. 2009, 81, 699–704. Hennig, M.; Neumann, J.; Wixforth, A.; Radler, J. O.; Schneider, M. F. Lab Chip 2009, 9, 3050–3053. Neumann, J.; Hennig, M.; Wixforth, A.; Manus, S.; Radler, J. O.; Schneider, M. F. Nano Lett. 2010, 10, 2903–2908. 10.1021/ac102979b  2011 American Chemical Society Published on Web 12/14/2010

Figure 1. (A) Membrane-associated proteins are accumulated at the edge of the lipid bilayer by hydrodynamic forces from a bulk flow above the SLB. (B) Applying a bulk flow in the opposite direction drives the proteins away from the edge of the bilayer with a drift velocity characteristic of each type of protein.

channel.21-23 The shear force from the flowing liquid above the SLB causes the upper leaflet of the lipid bilayer to move in the direction of the flow.21,22 Due to the strong coupling between the SLB and the support, the lower leaflet of the SLB remains essentially stationary, resulting in a rolling motion where the lipids in the upper leaflet move with an average velocity that is twice that of the front of the bilayer.21 It was also observed that fluorescently labeled lipids that are moving faster than the bilayer front can accumulate at the front of the lipid bilayer, whereas labeled lipid molecules that are slower do not.21 However, this type of molecular separation only works for a system consisting of two types of molecules where one of the molecules moves faster than the bilayer front, whereas the other type of molecule moves slower than the front of the lipid bilayer. In this work we present a more general method for the separation of molecules coupled to an SLB into different, spatially separated bands in the lipid bilayer. Protein molecules anchored to the upper leaflet of an SLB are expected to move with a velocity that is higher than that of the advancing bilayer front, when having a flow of liquid above the SLB. This will result in an accumulation of protein molecules at the front of the lipid bilayer. The reason for this accumulation is that the proteins are reluctant to move over the edge of the bilayer into the lower leaflet of the SLB (see Figure 1A). Reversing the direction of the bulk flow causes the accumulated protein molecules to move away from the edge of the bilayer, due to the same type of hydrodynamic force that caused them to move toward the edge of the lipid bilayer (see Figure (21) Jonsson, P.; Beech, J. P.; Tegenfeldt, J. O.; Hook, F. J. Am. Chem. Soc. 2009, 131, 5294–5297. (22) Jonsson, P.; Beech, J. P.; Tegenfeldt, J. O.; Hook, F. Langmuir 2009, 25, 6279–6286. (23) Jonsson, P.; Jonsson, M. P.; Hook, F. Nano Lett. 2010, 10, 1900–1906.

1B). In this scenario, each type of protein can be expected to have a characteristic drift velocity determined by the size and conformation of the protein molecule, as well as the frictional coupling between the protein and the lipid bilayer. A large protein that protrudes significantly into the bulk solution is expected to have a higher drift velocity than a small, flat protein remaining close to the SLB, as the large protein should experience a higher hydrodynamic force from the flowing liquid. In addition, a protein with many anchors to the SLB is expected to have a lower drift velocity than a similarly sized protein with fewer anchors to the SLB, as the protein with many anchors experiences a stronger frictional drag from the lipid bilayer. To evaluate the potential of the proposed method, the accumulation and separation of two membrane-bound proteins of similar molecular weight (∼60 kDa)24 was investigated: (i) the B vitamin biotin-binding protein streptavidin (SA), which is often used as a model protein in, for example, lipid-bilayer-aided twodimensional protein crystallization25,26 and numerous biotechnological and diagnostic applications,27 and (ii) the monosialoganglioside GM1 (GM1)-binding protein cholera toxin (CT), here consisting of five B subunits each of which can bind a GM1 molecule.28 Besides studies related to its toxicity, CT has been much used as a model protein for studies of multivalent interactions between proteins and cell membrane receptors.29-31 Since different numbers of anchors between the protein and the SLB are expected to affect the drift velocity, the method presented here also has the potential to detect and investigate multivalentdependent heterogeneity in the coupling between CT and GM1. EXPERIMENTAL PROCEDURE To experimentally explore the issues discussed above, and in particular the possibility of separating different membrane-bound proteins, we made use of the microfluidic device depicted in Figure 2. The channel consisted of a polydimethylsiloxane (PDMS) replica bonded to a glass slide (see Supporting Information for details on the microfluidic setup). An SLB was formed in the left part of the channel by injecting lipid vesicles into arm 1, at the same time as a buffer solution was injected into arm 4. The vesicles are adsorbed onto the glass surface where they subsequently fuse to form a planar SLB. The SLB consisted of 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) with either 0.0025 wt % 1,2-dioleoyl-sn-glycero3-phosphoethanolamine-N-(cap biotinyl) (biotin-PE) or 0.01 wt % GM1, unless otherwise stated. Here SA binds to the biotin-PE lipids, whereas CT binds to GM1. To enable the molecules to be visualized with epifluorescence microscopy, (24) http://www.sigmaaldrich.com (accessed August 2, 2010). (25) Darst, S. A.; Ahlers, M.; Meller, P. H.; Kubalek, E. W.; Blankenburg, R.; Ribi, H. O.; Ringsdorf, H.; Kornberg, R. D. Biophys. J. 1991, 59, 387–396. (26) Lou, C.; Wang, Z.; Wang, S. W. Langmuir 2007, 23, 9752–9759. (27) Laitinen, O. H.; Nordlund, H. R.; Hytonen, V. P.; Kulomaa, M. S. Trends Biotechnol. 2007, 25, 269–277. (28) Zhang, R. G.; Westbrook, M. L.; Westbrook, E. M.; Scott, D. L.; Otwinowski, Z.; Maulik, P. R.; Reed, R. A.; Shipley, G. G. J. Mol. Biol. 1995, 251, 550– 562. (29) Lauer, S.; Goldstein, B.; Nolan, R. L.; Nolan, J. P. Biochemistry 2002, 41, 1742–1751. (30) Ruprecht, V.; Brameshuber, M.; Schutz, G. J. Soft Matter 2010, 6, 568– 581. (31) Shi, J. J.; Yang, T. L.; Kataoka, S.; Zhang, Y. J.; Diaz, A. J.; Cremer, P. S. J. Am. Chem. Soc. 2007, 129, 5954–5961.

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Figure 2. Schematic illustration of the microfluidic setup used to concentrate and separate the membrane-bound proteins.

the SA molecules were labeled with the fluorescent group Cy3, and the CT molecules with the fluorescent group fluorescein isothiocyanate (FITC). After removal of excess vesicles from the microfluidic setup by rinsing with a buffer solution, a protein solution with either ∼170 nM SA-Cy3 or ∼30 nM CT-FITC (consisting of only the pentameric B subunits) was injected into arm 1, at the same time as a buffer solution was injected into arm 4 to prevent the protein molecules from adsorbing onto the bare glass surface in front of the SLB. During this process, the protein molecules bind to the receptors in the previously formed SLB. The anchoring of the protein molecules to the SLB was monitored by total internal reflection fluorescence microscopy. The proteins were injected until the adsorption of molecules had reached steady state (∼30 min). The bound proteins were subsequently driven toward the edge of the lipid bilayer by applying a bulk flow of buffer solution between the inlet at arm 1 and the outlet at arm 4, using a bulk flow rate of 200 µL/min unless otherwise stated. After accumulation of the proteins at the edge of the lipid bilayer, the bulk flow was reversed, causing the accumulated proteins to move away from the edge of the bilayer. For further details on the experimental procedure see the Supporting Information. RESULTS AND DISCUSSION Accumulation at the Edge of the Lipid Bilayer. Figure 3A shows how SA-Cy3 molecules accumulated at the edge of the lipid bilayer when applying a bulk flow of 200 µL/min from left to right in the figure. The maximum fluorescence intensity, corresponding to the highest concentration of protein molecules in the SLB, is observed near the edge of the lipid bilayer at the center of the channel. The maximum fluorescence intensity in this region increased after the bulk flow was applied but reached a constant value after typically 20-30 min (see Figure 3B). At the same time, a gradual broadening of the high-intensity region was observed. The maximum intensity at the edge of the bilayer leveled off at a value that was 78 ± 14 (n ) 3) times higher than before initiating the bulk flow. This corresponds to a surface coverage of ∼3.5 ± 0.6 × 10-9 mol/m2 of SA-Cy3 molecules (see Supporting Information for details). The accumulation of protein molecules was found to be dependent on the surface coverage of the receptors in the SLB. When increasing the amount of biotin-PE in the SLB from 0.0025 wt % to 0.025 wt %, thus 606

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increasing the initial coverage of SA-Cy3 by a factor of 10, the maximum value of the relative intensity increase at the center of the channel was 17 ± 2 (n ) 3), corresponding to a surface coverage of ∼7.7 ± 0.7 × 10-9 mol/m2. The maximum fluorescence intensity at the edge of the lipid bilayer is also dependent on the bulk flow rate, and increased by 10-30% when the flow rate was increased by a factor of 2 (see Figure 4A). The maximum surface coverage of SA-Cy3 is at least 1 order of magnitude lower than the surface coverage previously determined for the corresponding crystalline structure, where the proteins are in close proximity to each other.32 This suggests that the accumulated protein molecules can, in theory, be packed even more densely at the edge of the lipid bilayer than observed here. There are several possible explanations to this observation: (i) The surface coverage of the protein is higher than predicted by the observed fluorescence intensity, as a lower protein concentration is measured due to self-quenching of the fluorescent dye. (ii) A fraction of the protein molecules is continuously lost from the high-concentration region at the center of the channel to the sides of the lipid bilayer. (iii) The repulsive intermolecular forces between the protein molecules start to become important at the protein concentrations observed in the high-concentration region at the edge of the lipid bilayer. The first of these explanations is less likely, since the observed maximum surface coverage increases when using a higher concentration of biotin-PE in the SLB. This indicates that the surface coverage is below a value corresponding to densely packed protein molecules. Furthermore, when increasing the bulk flow rate in the channel the maximum intensity near the front of the lipid bilayer was also observed to increase, whereas the total intensity, corresponding to the number of protein molecules accumulated at the edge of the lipid bilayer, was not notably affected by the change in the bulk flow rate (see Figure 4). If there was significant self-quenching, then the total intensity would decrease as the protein molecules become more closely packed, as indicated by the increase in peak intensity, but this was not observed here. An observation that supports the second scenario, that a fraction of the protein molecules is continuously lost by an outflow from the central part of the bilayer, is the discrepancy between the shape of the advancing bilayer front and the parabolic shape of the hydrodynamic force arising from the bulk flow.22 To maintain the stationary curvature of the bilayer front seen in Figure 3A, an outgoing flow of lipids from the center of the SLB to the sides is necessary, where the velocity of the SLB resulting from the hydrodynamic flow is low. This observation could also explain why the maximum intensity at the edge of the lipid bilayer is lower than that arising from a densely packed layer of protein molecules. If it is assumed that a certain number of protein molecules leave the high intensity region at the center of the bilayer per unit time, then the number of protein molecules in this region will level off when the loss of protein molecules from the region is equal to the number of protein molecules arriving to the region. (32) Gast, A. P.; Robertson, C. R.; Wang, S. W.; Yatcilla, M. T. Biomol. Eng. 1999, 16, 21–27.

Figure 3. (A) Fluorescence micrograph of SA-Cy3 accumulated at the edge of the lipid bilayer (x ∼ 0) together with a line profile of the intensity through the center of the channel, normalized to the intensity before the bulk flow was applied. (B) Overlaid intensity profiles of SA-Cy3 at the bilayer front at 10-min intervals after a 200 µL/min bulk flow was applied at t ) 0 s. The arrow indicates increasing time.

Figure 4. (A) The maximum intensity at the edge of the lipid bilayer from SA-Cy3 as a function of time at different bulk flow rates: (i) 200 µL/min, (ii) 300 µL/min, (iii) 200 µL/min, and (iv) 100 µL/min. (B) Corresponding accumulation of protein molecules at the edge of the bilayer. The solid line is a linear fit to the initial slope of the data.

The maximum concentration and the shape of the concentration profile may also be affected by repulsive intermolecular forces between the protein molecules at the observed protein concentrations. Electrostatic and steric forces, as well as entropic effects, all contribute to the repulsive force at higher surface coverage.33 For a surface coverage of 2 × 10-9 mol/m2, which is around half the maximum surface coverage of SA-Cy3 molecules in Figure 3A, the average distance between two nearest-neighbor protein molecules is ∼30 nm. At this distance the repulsive intermolecular force will be of a comparable magnitude to the hydrodynamic force acting on a single protein in the SLB (see Supporting Information), which could limit the accumulation of protein molecules at the edge of the lipid bilayer. Similar behavior to that observed when accumulating SA-Cy3 at the edge of the lipid bilayer was observed with CT-FITC. However, in contrast to SA-Cy3, the accumulated CT-FITC molecules had the maximum intensity in the center of the channel occurring a distance of ∼10 µm from the edge of the lipid bilayer (see Figure 5). That the intensity maximum occurs a distance away from the bilayer front, as shown in Figure 5, has also been observed by Groves et al. when driving two types of charged lipid molecules, NBD-PE and cardiolipin, toward a stationary barrier using an (33) Evans, D. F.; Wennerstro ¨m, H. The colloidal domain; VCH Publishers, Inc.: New York, 1994.

Figure 5. Fluorescence micrograph of CT-FITC molecules at the bilayer front (x ∼ 0) together with a line profile of the intensity through the center of the channel, normalized to the intensity before the bulk flow was applied.

electric field.11 Cardiolipin preferentially accumulated at the barrier, excluding NBD-PE molecules from this area, which resulted in a concentration profile of NBD-PE similar to the one seen in Figure 5. An explanation of the behavior observed here could be that the intermolecular interactions between different membrane-associated molecules favor a separation of the CT-FITC molecules to a region behind the edge of the lipid bilayer. It is Analytical Chemistry, Vol. 83, No. 2, January 15, 2011

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Figure 6. Micrographs of the fluorescence intensity from SA-Cy3 at (A) t ) 0 s, (B) t ) 100 s, and (C) t ) 200 s after a bulk flow of 200 µL/min is applied from right to left in the figure. The dashed line shows the position of the edge of the bilayer at t ) 0.

also possible that there are nonlabeled molecules in the SLB that accumulate with a higher preference than CT-FITC at the edge of the lipid bilayer. The maximum value of the relative intensity increase was also smaller for CT-FITC than for SA-Cy3, with values of 7.9 ± 1.1 (n ) 3) and 6.2 ± 1.2 (n ) 3) for an SLB with 0.01 wt % GM1 and 0.1 wt % GM1, respectively. Estimating the surface coverage corresponding to these values from the initial coverage of GM1 in the SLB is rather uncertain but results in an upper limit of 2.1 ± 0.3 × 10-9 mol/m2 with 0.01 wt % GM1 in the SLB and 16.2 ± 3.1 × 10-9 mol/m2 with 0.1 wt % GM1 in the SLB (see Supporting Information for details). These values of surface coverage are of the same magnitude as those obtained for SA-Cy3 but, as was also the case for SA-Cy3, they are lower than those for the crystalline structure of the protein.34 Drift Velocity and Protein Separation. When reversing the direction of the bulk flow, the accumulated protein molecules are driven in the opposite direction by the hydrodynamic force. Figure 6 shows micrographs of the fluorescence intensity from SA-Cy3 after a bulk flow of 200 µL/min has been applied from right to left in the figure, driving the accumulated protein molecules away from the edge of the lipid bilayer. (34) Ludwig, D. S.; Ribi, H. O.; Schoolnik, G. K.; Kornberg, R. D. Proc. Natl. Acad. Sci. U.S.A. 1986, 83, 8585–8588.

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Initially, the shape of the accumulated protein molecules followed the curvature of the advancing bilayer front, which remained approximately constant over time, as observed previously.21 After reversal of the bulk flow, the intensity profile became gradually more parabolic as the protein molecules were driven away from the edge of the lipid bilayer. Since the hydrodynamic force driving the protein molecules is highest at the center of the channel, and falls to zero at the walls of the channel,22 the drift velocity of the protein molecules will be highest at the center of the channel, resulting in the gradually more parabolic profile. The magnitude of the intensity peak will decrease rapidly after the bulk flow has been reversed, due to diffusion and intermolecular repulsion of the protein molecules, at the same time broadening the peak (see Figure 7A). The peak will also initially become asymmetric, with a steeper slope on the side closest to the edge of the lipid bilayer. This effect is most pronounced shortly after the flow has been reversed, whereas at longer times the shape approaches that of a Gaussian curve. One possible explanation of this behavior is that the hydrodynamic force driving the protein is affected by the protein concentration in the SLB, which in turn will affect the drift velocity of the protein molecules. This behavior is not unlikely, since it can be expected that the hydrodynamic force will act only on the upper surface of the protein molecules when the molecules are packed close together, resulting in a lower force on each protein molecule than would have been the case if the protein molecules had been spaced more widely apart, in which case the hydrodynamic force could act on the entire protein molecule. This will in turn result in the protein molecules on either side of the peak moving faster than the protein molecules at the center of the intensity peak, resulting in the asymmetric profile. Since the difference in concentration over the peak decreases with time, the hydrodynamic force at longer times will be approximately equal over the entire intensity peak, resulting in the loss of asymmetry of the peak. Figure 7B shows the position of the center of mass of the intensity peak in each image frame after the bulk flow was reversed and the protein molecules were driven away from the edge of the lipid bilayer. Initially, the position of the peak changes slowly but reaches a steady drift velocity after ∼150 s. This supports the hypothesis made earlier that the hydrodynamic force is smallest where the protein concentration is high. Since the protein molecules are close together at flow reversal, the hydrodynamic force on the anchored molecules will be lower than the force on the same molecules in a diluted system, resulting in the slower drift velocity shortly after flow reversal. From the position of the intensity peak it is also possible to calculate the steady-state drift velocity of the protein molecules at different bulk flow rates. The drift velocity was 0.48 ± 0.02 µm/s, 1.24 ± 0.02 µm/s, and 1.81 ± 0.13 µm/s (n ) 2-3), when the bulk flow was 100 µL/min, 200 µL/min, and 300 µL/min, respectively. The drift velocity scales approximately linearly with the bulk flow, especially at the two higher bulk flow rates. A convenient way of representing the drift velocity, v, is therefore to scale it with the applied bulk flow rate, Q, yielding vSA/Q ) 366 ± 19 m-2 (n ) 5) when using the data from the 200 µL/min and 300 µL/min experiments. The drift velocity of the SA-Cy3 molecules when the bulk flow is reversed can be compared to the drift velocity of the advancing

Figure 7. (A) Line profiles of the intensity at the center of the channel at different times after a 200 µL/min bulk flow has been applied in the negative x-direction at t ) 0 s. The intensities are normalized to the peak intensity at t ) 0 s. (B) The position of the center of mass of the SA-Cy3 intensity peak at the center of the channel at different bulk flow rates, Q, as a function of time. The solid lines are linear fits to the data.

bilayer front which was vfront/Q ) 69 ± 3 m-2 (n ) 6). The average drift velocity of the upper leaflet of the SLB can then be estimated to 138 ± 6 m-2, if it is assumed that the bilayer front moves with half the average velocity of the upper leaflet.21 From this value, and the parabolic velocity profile of the lipid molecules over the width of the channel,22 the velocity of the lipids at the center of the channel can be estimated to be vlipids/Q ) 184 ± 8 m-2 (see Supporting Information for details). The SA-Cy3 molecules thus move roughly twice as fast as the lipids in the upper leaflet of the SLB. These values can also be compared to the number of SA-Cy3 molecules arriving at the bilayer front per unit time. From the initial slope of the curve in Figure 4B, the average velocity of the accumulating protein molecules, in the center of the channel, can be determined to be vacc/Q ) 270 ± 20 m-2 (see Supporting Information for details). This value is 74% of the drift velocity obtained for SA-Cy3 at the center of the channel when driving the protein molecules away from the edge of the bilayer. When driving CT-FITC away from the edge of the lipid bilayer, the intensity peak was observed to divide into multiple peaks, with different drift velocities (see Figure 8). The peak divides into two mobile fractions relatively quickly, whereas a third fraction is observed to separate from the slow fraction at longer times (see the inset in Figure 9A). The distance traveled by the center of mass position of each of the intensity peaks in Figure 9A is given in Figure 9B, from which the drift velocity can be obtained by a linear fit to the data points. Only images where the protein peaks could be separated from each other were used in the analysis. The time ∆t ) 0 in Figure 9B is therefore chosen after the protein peaks had separated. Note also that ∆t * t, where t is the time after flow reversal in Figure 9A. In addition, the distance traveled at ∆t ) 0 has been set to zero for all of the intensity peaks. As for SA-Cy3, the drift velocities scaled linearly with the applied bulk flow, making it appropriate to normalize the drift velocity to the bulk flow rate, resulting in the three CT fractions having the drift velocities: vCT1/Q ) 121 ± 2 m-2, vCT2/Q ) 310 ± 8 m-2, and vCT3/Q ) 645 ± 20 m-2 (n ) 2-5). Thus, the fast fraction moves 5.3 times faster than the slow fraction, and 2.1 times faster than the intermediate fraction. The different drift velocities can be interpreted as heterogeneity among the CTFITC molecules, possibly due to fractions with different

Figure 8. Micrographs of the fluorescence intensity from CT-FITC at (A) t ) 30 s, (B) t ) 70 s, and (C) t ) 110 s after a bulk flow of 200 µL/min was applied from right to left in the figure. The dashed line shows the position of the edge of the bilayer at t ) 0.

numbers of GM1 anchors to the SLB, where the fast-moving fraction would have the least number of anchors. Interestingly, the slowest of the protein fractions was observed to have a drift velocity lower than that of the lipids in the upper leaflet of the SLB. However, the proteins were still moving faster than the drift velocity of the bilayer front, which is a prerequisite for the accumulation of the protein molecules at the edge of the bilayer. This indicates that this protein fraction experiences a Analytical Chemistry, Vol. 83, No. 2, January 15, 2011

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Figure 9. (A) Line profiles of the intensity of CT-FITC at the center of the channel at different times after a 200 µL/min bulk flow has been applied in the negative x-direction at t ) 0 s. The intensities are normalized to the peak intensity at t ) 0 s. The inset shows the splitting of the slow peak into two peaks at t ) 220 s. (B) Distance traveled by the different intensity peaks in Figure 9A as a function of time. The time ∆t ) 0 is chosen after the protein peaks had separated. The solid lines are linear fits to the data.

significant drag force, not only from the lipids in the upper leaflet of the SLB, but also from the lower leaflet of the SLB, and possibly the supporting substrate, thus reducing the drift velocity of the protein molecules. Upon increasing the amount of GM1 in the SLB from 0.01 wt % to 0.1 wt % the size of the fast fraction decreased, whereas the slow fraction increased in size (see Supporting Information for details). If it is assumed that the fast CT-FITC fraction has fewer GM1 anchors to the SLB than that of the slow CT-FITC fraction, this could be interpreted as an increase in the average number of GM1 anchors per CT-FITC molecule. A decrease in mobility of the proteins in the SLB when increasing the amount of GM1 in the lipid bilayer was also observed when measuring the diffusivity of CT-FITC by fluorescence recovery after photobleaching.35 The diffusivity decreased from 0.39 ± 0.04 µm2/s (n ) 8) to 0.31 ± 0.03 µm2/s (n ) 5) when increasing the amount of GM1 in the SLB from 0.01 wt % to 0.1 wt %. SA-Cy3, on the other hand, showed similar values of diffusivity (∼1.4 µm2/s,