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Langmuir 2006, 22, 4408-4414
Bicontinuous Cubic Phase of Monoolein and Water as Medium for Electrophoresis of Both Membrane-Bound Probes and DNA Nils Carlsson,† Nima Sanandaji,† Marina Voinova,‡ and Bjo¨rn Åkerman*,† Department of Chemistry and Bioscience, and Department of Applied Physics, Chalmers UniVersity of Technology, SE-41296 Go¨teborg, Sweden ReceiVed July 29, 2005. In Final Form: January 26, 2006 Porous hydrogels such as agarose are commonly used to analyze DNA and water-soluble proteins by electrophoresis. However, the hydrophilic environment of these gels is not suitable for separation of important amphiphilic molecules such as native membrane proteins. We show that an amphiphilic liquid crystal of the lipid monoolein and water can be used as a medium for electrophoresis of amphiphilic molecules. In fact, both membrane-bound fluorescent probes and water-soluble oligonucleotides can migrate through the same bicontinuous cubic crystal because both the lipid membrane and the aqueous phase are continuous. Both types of analytes exhibit a field-independent electrophoretic mobility, which suggests that the lipid crystal structure is not perturbed by their migration. Diffusion studies with four membrane probes indicate that membrane-bound analytes experience a friction in the cubic phase that increases with increasing size of the hydrophilic headgroup, while the size of the membrane-anchoring part has comparatively small effect on the retardation.
Introduction Electric-field-driven migration in porous gels is commonly used to separate DNA and proteins in order to measure a wide range of properties such as monomer sequence, molecular weight, and isoelectric point.1 Agarose and polyacrylamide gels are useful polymeric networks for such purposes because the electrophoretic transport through their aqueous pores of micrometer to nanometer dimensions causes size-separation by filtration (sieving).2 A large protein or DNA molecule migrates with lower velocity than a smaller one, and molecular mass can be determined by comparison with a set of homologous size standards. However, a severe limitation with the hydrophilic gels (containing typically 99% water) is that they are usually not compatible with important amphiphilic molecules such as native membrane proteins. Protein solubilization by use of detergents is one approach, but the appropriate surfactant has to be found for each protein more or less by trial and error3 and the surfactant tends to affect the gel velocity of the solubilized protein4,5 and hence the apparent mass compared to size standards. One solution may be to run electrophoresis of native proteins in planar lipid membranes, and separation of a peripheral-type of membrane proteins anchored to a surface-supported bilayer has indeed been demonstrated.6 However, for integral membrane proteins that span the membrane, lipid drag on the hydrophobic part is relative insensitive to protein size as predicted theoretically.7 Second, in the planar case, the aqueous drag contributes little to the total friction even when the hydrophilic part constitutes a major part of the protein,8 as expected from the viscosity of the lipid layer * To whom correspondence should be addressed. † Department of Chemistry and Bioscience. ‡ Department of Applied Physics. (1) Westermeier, R. In Electrophoresis in Practice, 3rd ed.; Wiley-VCH: New York, 2001. (2) Viovy, J. L. ReV. Mod. Phys. 2000, 72, 813-872. (3) Luche, S.; Santoni, V.; Rabilloud, T. Proteomics 2003, 3, 249-253. (4) Heuberger, E. H. M. L.; Veenhoff, L. M.; Duurkens, R. H.; Friesen, R. H. E.; Poolman, B. J. Mol. Biol. 2002, 317, 591-600. (5) Werhahn, W.; Braun, H. P. Electrophoresis 2002, 23, 640-646. (6) Groves, J. T.; Wu¨lfring, C.; Boxer, S. G. Biophys. J. 1996, 71, 2716-2723. (7) Vaz, W. L. C.; Criado, M.; Madeira, V. M. C.; Schoellmann, G.; Jovin, T. M. Biochemistry 1982, 21, 5608-5612. (8) Tamm, L. K. Biochemistry 1988, 27, 1450-1457.
Figure 1. Schematic structures of the porous media used in this work. (a) Diamond cubic phase (Q224), with the aqueous channels (grey circles) surrounded by lipid bilayers. (b) Schematic representation of a porous agarose network of randomly oriented gel fibers.
being about 100 times higher than for water.9 The electrophoretic velocity in a planar membrane can therefore not be expected to be sensitive to overall membrane protein size. In an attempt to retain the possibility of membrane-bound migration but with an enhanced hydrophilic drag compared to planar membranes, we have explored the potential of running electrophoresis in a bicontinuous cubic phase formed by the lipid monoolein and water10 (Figure 1a). This liquid-crystalline structure contains a three-dimensional system of water-filled pores with a diameter of about 5 nm.11 It is created by a continuous and highly curved lipid bilayer, which is able to accommodate at least certain integral membrane proteins.11,12 Our two-part working hypothesis is that if (1) membrane-bound molecules can migrate along the bilayer then (2) the aqueous pores may be of the right size to cause sieving on their hydrophilic parts, analogous to hydrophil electrophoresis in conventional gels. We have taken the approach of investigating these two conditions individually, by separating the membrane-anchoring function and the hydrophilic parts in amphiphilic analytes. Previously, we have used hydrophilic oligonucleotides of well(9) Tsapis, N.; Reiss-Husson, F.; Ober, R.; Genest, M.; Hodgens, R. S.; Urbach, W. Biophys. J. 2001, 81, 1613-1623. (10) Larsson, K. J. Phys. Chem. 1989, 93, 7304-7314. (11) Rummel, G.; Hardmeyer, A.; Widmer, C.; Chiu, M. L.; Nollert, P.; Locher, K. P.; Pedruzzi, I.; Landau, E. M.; Rosenbusch, J. P. J. Struct. Biol. 1998, 121, 82-91. (12) Landau, E. M.; Rosenbusch, J. P. Proc. Natl. Acad. Sci. 1996, 93, 1453214535.
10.1021/la052086l CCC: $33.50 © 2006 American Chemical Society Published on Web 03/29/2006
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Chart 1. Membrane Probes DiI-C18-(5)-Ds (DiI), DiOC18 (DiO), BODIPY-C5-HPA (C5-HPA), and [Ru(phen)2Me2Dppz]2+
Table 1. Properties of the Gels
Chart 2
type of matrix monoolein cubic diamond phase agarose 5%
characterized nanometer sizes13 comparable to hydrophilic parts of membrane proteins and showed14 that the aqueous pores indeed are of suitable size and nature to cause size-selective sieving. Here we study membrane-anchored fluorescent probes (Chart 1) with hydrophilic parts which are small compared to most membrane proteins. With the aim to establish the mode of migration (rather than separation), we show that the continuous nature of the lipid phase allows membrane probes to be transported by electrophoresis in the same cubic phase that causes sieving of oligonucleotides. As control for conventional hydrogels, we use a hydroxyethylated agarose gel (Figure 1b) which has similar average pore size as the monoolein cubic phase.
a
continuous lipid wall randomly oriented fibers
average pore radiusa (nm) 2.5 3.7
For monoolein from ref 11. For 5% metaphor agarose from ref 19.
was used as received, 5′-end-modified with the fluorescent dye Cy5, and dissolved in water. We have earlier14 shown that ss25 forms a nearly symmetric hairpin held together by six base pairs (Chart 2). In particular, the eletrophoretic properties of ss25 in both monoolein and agarose gels is equivalent to that of a 12-mer duplex, and the ss25 hairpin is therefore predicted to behave as a rigid rod with a radius of about 1 nm and is 4 nm long (0.34 nm per base pair).15 We use it here because of its higher base-pairing stability
than a 12-mer duplex, considering the destabilizing acidic conditions of the present monoolein protocol (see below). Preparation of Gels. The details of preparation of slabs of monoolein cubic phase suitable for electrophoresis is described elsewhere.14 Briefly, the paste-like monoolein cubic phase (here referred to as the cubic gel) was formed by submerging a layer of crystalline monoolein under an excess of electrophoresis buffer (0.2 M acetate buffer, pH 4 if nothing else is indicated) at 37 °C for 4-5 days. The resulting swollen phase was optically transparent and isotropic, as expected for the cubic phase, assumed to be the diamondtype (Q224 of symmetry Pn3m) since it is formed in contact with an excess of water.10,16 We have not performed any further structural studies on the monoolein phase, but we note that the swelling protocol we use is similar to that used in some structural studies16 and, second, that it is unlikely that the low (µM) concentrations of analytes we use will perturb the liquid-crystal structure. Agarose gels were cast from solutions prepared by dissolving the metaphore agarose powder in electrophoresis buffer by boiling for 10 min. The properties of the two gels are summarized in Table 1. Electrophoresis. Samples of a total volume of 20 µL in electrophoresis buffer contained either one of the fluorescent membrane probes at concentrations between 0.25 and 2 µM (and a final ethanol concentration of 5% carried over from the stock solution) or the oligonucleotide at concentrations between 0.5 and 2 µM (strands). The samples were loaded in separate wells, and the gels were run in electrophoresis buffer (no ethanol) at constant field strength between 2 and 15 V/cm at 20 °C. Imaging of the dye fluorescence in a Storm or Fluoroimager scanner (100 µm resolution) was used to monitor the position of the sample molecules on the gel. Excitation was 633 nm (650 nm long-pass emission) for DiI and oligonucleotides, and 488 nm excitation (530 ( 15 band-pass emission) for DiO, C5-HPA, and [Ru(phen)2Me2dppz]2+. A charged impurity in the monoolein sample, most likely oleic acid resulting from hydrolysis of monoolein, limited electrophoresis to pH below 5.6.14 The membrane probes and the oligonucleotides themselves
(13) Pecora, R. Science 1991, 251, 893-898. (14) Carlsson, N.; Winge, A. S.; Engstro¨m, S.; A° kerman, B. J. Phys. Chem. 2005, 109, 18628-18636.
(15) Bloomfield, V. A.; Crothers, D. M.; Tinoco, I. In Nucleic Acids, University Science Books: Sausalito, CA, 2000. (16) Qiu, H.; Caffrey, M. Biomaterials 2000, 21, 223-234.
Materials and Methods Monoolein from Danisco (Denmark) and hydroxyethylated Metaphore agarose (FMC) was used as received. The fluorescent membrane probes (Chart 1) DiI-C18-(5)-DS (DiI), DiOC18 (DiO), and BODIPY-C5-HPA (C5-HPA) (Molecular Probes, Eugene, OR) were dissolved at 1 mg/mL in ethanol as stock solutions. The ruthenium complex [Ru(phen)2Me2dppz]2+ was a gift from P. Lincoln, Chalmers University, and was used in its racemic form. A single-stranded oligonucleotide (SGS, Sweden) with the following 25-mer sequence (ss25) 3′-GACCGGCAGCAAAATGTTGCAGCAC-5′
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Figure 2. Fluorescence images of gels after electrophoresis of membrane probe DiI or the 25-mer hairpin oligonucleotide (25mer) in either (to the left) the monoolein-water cubic phase (5 h of electrophoresis) or (to the right) in a 5% hydroxyethylated (Metaphor) agarose gel (5 min). Field strength 5 V/cm and pH ) 4 (acetate buffer) in both gels. Arrows indicate position of sample well, scalebar is 6 mm. Concentration of membrane probe 0.9 (lanes 1, 5), 1.4 (lane 2), and 1.9 µM (lanes 3, 6) and of oligonucleotide strands 0.6 µM (lanes 4, 7). are stable under these conditions, whereas the Cy-5 label of the oligunucleotides undergoes a slow degradation (over hours) with an accompanying loss of fluorescence.14 Diffusion. Diffusion coefficients were measured by introducing a zone of the analyte into the bulk of the gel by electrophoresis. The field was then turned off and the gel was scanned repeatedly in order to record the broadening of the concentration profile with time. Diffusion coefficients were evaluated from fitting the concentration profiles after different durations to the broadening Gaussian zoneshapes predicted for one-dimensional diffusion.17 The diffusion of [Ru(phen)2Me2dppz]2+ was studied by layering the dye sample solution on top of a sample of cubic phase preformed in a spectroscopic cuvette, which was subsequently sealed. The diffusion coefficient was obtained from the rate of motion into the cubic phase of the dye concentration front, measured from fluorescence scanning images (see Supporting Information).
Results 1. Choice of Porous Matrixes. Monoolein and water form a multitude of liquid crystals.10,16 We choose to work with the diamond-type cubic phase (Figure 1a) because it can be in equilibrium with a water-rich phase.10 This phase property was exploited to allow the gel to be in contact with the buffer reservoirs which contain the electrodes and, second, to allow the electrophoresis to be run with the gel submerged under buffer. The latter aspect is an important practical result because this submarine mode of electrophoresis is the optimal approach to control ionic composition and temperature in the gel. We employ a 5% gel of hydroxyethylated agarose as a control for a conventional hydrophilic matrix because it has an average pore size close to that which holds in the cubic phase (Table 1). 2. Cubic Phase. 2.1. Migration of Amphiphilic and Hydrophilic Molecules. The left-hand gel image in Figure 2 shows the positions of the zones of the membrane probe DiI and of the 25-mer hairpin oligonucleotide (see Materials and Methods) after ∼5 h of electrophoresis in the cubic phase at 5 V/cm. It is seen that both types of molecules have entered the monoolein cubic phase from the wells (arrow) and migrate within it as distinct zones. The right-hand part of Figure 2 shows the same samples run for 5 min on the 5% hydroxyethylated agarose gel at the same field strength. The oligonucleotide again migrates as a distinct zone, but no zone of DiI can be observed on the agarose gel. Figure 2 shows that the behavior of the membrane probe is distinctly different in the cubic phase and the agarose gel, in accordance with the monoolein gel being amphiphilic in nature compared to the water-rich hydrogel. The intensity of the oligonucleotide zone is much weaker in the cubic phase compared to the agarose gel. The Cy5 dye is known to be bleached at acidic (17) Atkins, P. W. In Physical Chemistry, 6th ed.; Oxford University Press: New York, 1998.
Figure 3. Scans of fluorescence intensity in the cubic phase along lanes containing a zone of DiI membrane probe after indicated times of electrophoresis. The well is at x ) 0. Field strength 5 V/cm.
Figure 4. Migrated distance vs time for the membrane probe DiI (circles) and the 25-mer hairpin oligonucleotide (squares) undergoing electrophoresis in the monoolein cubic phase at 5 V/cm. Open and closed symbols represent two independently prepared gels. Simple open symbols represent migration starting from the well (first three curves to the left in Figure 3); crossed open symbols represent the distance starting from a point 10 mm into the same gel. Velocities from the slopes on the two gels are 0.62 ( 0.06 and 0.59 ( 0.04 mm/h for the oligonucleotide and 0.37 ( 0.04 and 0.23 ( 0.01 mm/h for the membrane probe. Error bars correspond to deviation between duplicate samples run on the same gel.
pH,14 and this effect is more pronounced in the monoolein gel due to a longer exposure that result from the slower migration. Importantly, the monoolein part of Figure 2 show that DiI is stable at the acidic pH at least for hours, so the absence of observable zones in the agarose gel (lanes 5 and 6) is not due to bleaching of probe fluorescence. Second, the distance migrated by the oligonucleotide is about the same in the two gels despite the much shorter running time for the agarose gel, which shows that its velocity is markedly lower in the monoolein cubic phase. More precisely, at 5 V/cm the 25mer hairpin oligonucleotide has a velocity V of 0.6 ( 0.07 mm/h in the monoolein cubic phase, which is about 50 times lower than in the agarose gel where V ) 29.1 ( 0.4 mm/h (results not shown). The migration of both samples in the cubic phase was monitored by repetitive scanning of the gels. Figure 3 shows fluorescence intensity profiles of the DiI membrane probe recorded by scanning along lane 3 in Figure 2 after different durations of electrophoresis at 5 V/cm. The zone is seen to broaden as it proceeds, but in all instances, there is a well-defined intensity maximum, which was taken as the position of the zone. The open circles in Figure 4 show how this zone-position for DiI varies with electrophoresis time, as measured in two ways. The open circles show migrated distance measured relative to the edge of the well by making use of curves such as the first three ones in Figure 3. By contrast,
Electrophoresis of Membrane-Bound Probes and DNA
Langmuir, Vol. 22, No. 9, 2006 4411 Table 2. Experimental Results on the Membrane Probesa probe µb (10-6 cm2/V‚s) Dd (10-8 cm2/s) fµ /fDf hydrophobic tailsg size of headgrouph (Å)
Figure 5. Electrophoretic velocity in the cubic phase vs electric field strength for membrane probe DiI at indicated concentrations. Straight line is the least-squares fit to all data.
the crossed open circles show migration in the bulk of the gel by plots of the distance migrated relative to a position 10 mm into the gel (fourth curve in Figure 3) that was reached after 21 h of electrophoresis. The hydrophilic 25-mer hairpin oligonucleotide exhibited the same type of progressively moving intensity profiles in the monoolein gel as in Figure 3 (results not shown), in agreement with earlier observations.14 The derived time plots of zone position vs time for ss25 on the same gel as the DiI probe are included in Figure 4 (as squares), both when starting from the well (open squares) and in the bulk of the gel (crossed open squares). The approximate linearity of the plots in Figure 4 shows that both types of biomolecules migrate with a constant velocity over macroscopic distances. Duplicate samples on the same cubic gel show good reproducibility (error bars), and the membrane probe is seen to migrate about 1.7 times slower than the 25-mer oligonucleotide. Notably, the velocities (slopes) near the well and in the bulk of the gel are very similar for DiI, as well as for the oligonucleotide. This observation indicates that there is no marked perturbation of the cubic structure near the well. The plots of distance vs time were also linear for DiO, C5-HPA, and [Ru(phen)2Me2dppz]2+ (data not shown), as in Figure 4. When the experiment was repeated on an independently prepared cubic gel (Figure 4, closed symbols) the velocity of the oligonucleotide (solid squares) was the same as in the first gel within experimental uncertainty, whereas the DiI membrane probe (solid circles) was 30% slower. This latter difference probably reflects the degree of variation between different gel-preparations with the present protocol, and suggests that the membrane probe is more sensitive to variations in the structure of the prepared cubic phase. 2.2. Effect of Field Strength and Probe Concentration on the Electrophoretic Velocity. Figure 5 shows the electrophoretic velocity, V, of the DiI probe in the cubic phase as a function of field strength, measured at four different concentrations of the probe. The field dependence is approximately linear, i.e., the electrophoretic mobility µ ) V/E is nearly constant. Careful comparison of the data points at a given field strength show that the velocity exhibits no systematic dependence on the probe concentration. In fact, the pattern of scattering around the linear field trend is similar for all concentrations. Since each fieldstrength experiment was run on a separate gel (containing samples at all four probe concentrations), the deviation from a linear trend reflects the degree of reproducibility in the preparation of the monoolein cubic phase. The field behavior of DiO and [Ru(phen)2Me2dppz]2+ was also linear (data not shown), and the corresponding electrophoretic mobilities, µ (obtained from the slopes), are summarized in Table 2. The values for DiI at pH
C5-HPA
[Ru(phen)2Me2dppz]2+
DiI
DiO
2.5 ( 0.4 (1.6 ( 0.4) 7.6 ( 0.1 (6 ( 0.7) 1.2 (1.5) 2C18 19
1.8 ( 0.3
n.a.c
3.9 ( 0.8
14 ( 0.5
17 ( 1
20.8 ( 1.3e
3.0
n.a.c
4.2
2C18 12
1C17 10
a Error estimates are variations between duplicate samples on the same gel. pH ) 5 except values in parentheses which are at pH 4 b Electrophoretic mobility from slopes as in Figure 5. c C5-HPA migrated toward the negative electrode despite the formal negative charge, so its mobility was not included. d (Macroscopic) diffusion coefficient from concentration profiles as in Figure 6. The microscopic diffusion coefficient D0 for motion along the bilayer itself is somewhat higher, D0 ) D/β where the “obstruction” factor, β, due to the nonplanar membrane geometry is β ) 0.74 for the diamond cubic phase.9 e Measured from spreading of a concentration front rather than a concentration peak (see Methods). f Ratio of translation friction coefficients derived from mobility fµ ) Q/µ and from diffusion fD ) kT/D. g Number and length of hydrophobic tails. For the ruthenium complex, the tail corresponds to the Me2dppz ligand 18 h Estimated as longest dimension of chromophoric headgroup.
4 and 5 differ by 35%, but since these measurements were performed on independently prepared gels we cannot conclude that the difference reflects an effect of pH. The membrane probe C5-HPA also exhibited a linear field dependence but unexpectedly migrated toward the negative electrode despite its formal negative charge. Presently, we do not have an explanation for this behavior, although complex formation with buffer ions is a possibility, and C5-HPA was therefore only used in diffusion studies. 2.3. Diffusion. Diffusion of the membrane probes was investigated by monitoring the spreading of a narrow sample zone (in absence of field) after it had been introduced into the cubic phase by electrophoresis and the field had been turned off. Figure 6a shows the intensity profiles for DiI obtained from repetitive scanning of the gel and the fits to the intensity profiles I(x,t) predicted by one-dimensional diffusion17
I(x,t) ) A e-x /w 2
2
(8)
where w2 ) 4Dt and A ) Io/(πDt)1/2 with diffusion coefficient, D, and integrated intensity, Io. Plots of the fitted values of w2 vs time are linear (Figure 6b), and a diffusion coefficient, D ) (6 ( 0.7) × 10-8 cm2/s (at pH 4), was evaluated from the slopes measured for duplicate samples on the same gel. At pH 5, a higher value of (7.6 ( 0.1) × 10-8 cm2/s was obtained. Again, we cannot assign this to an effect of pH since the difference is of the same magnitude as the observed variation between independently prepared gels (as inferred from the mobility experiments). Diffusion experiments were also performed with DiO and C5-HPA, and the diffusion data are summarized in Table 2. The diffusion of [Ru(phen)2Me2dppz]2+ was studied by monitoring the spreading of the dye into the cubic phase from an overlayered sample solution (see Materials and Methods). The intention was to probe potential binding to the membrane by measuring how the dye partitions between the cubic crystal and the overlayered aqueous phase, but the equilibration was too slow under present conditions to achieve this goal. However, we found that the ruthenium complex exhibits a strongly enhanced
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Figure 6. Measurements of diffusion coefficients of membrane probes. (a) Concentration profiles after indicated durations (hours) of diffusional spreading of a zone of DiI in the cubic phase. Dashed lines are fits to eq 7 which gives the value of w2 for the indicated times. The left third of the profile (end of dashed curves) was left out of the fit because of the asymmetric shape due to an optical perturbation from the well edge at x ) -2. (b). Plots of fitted w2 values from panel (a) vs diffusion time. Slope ) 4D where D is diffusion coefficient. (c) Diffusion of [Ru(phen)2Me2dppz]2+ from an aqueous solution into monoolein cubic phase. Position L of concentration front is plotted vs diffusion time (see Supporting Information for images). Inset shows that L2 is linear in time as expected for diffusion process. The diffusion coefficient is obtained as slope/2 for 1D diffusion.
fluorescence in the monoolein cubic phase compared to the monoolein-free control (see Supporting Information), which supports membrane binding (see below). We also found that the concentration front moving into the cubic phase was too sharp to fit eq 8, probably due to the concomitant adsorption to the membrane.18 However, plots of the front position, L (Figure 6c), showed the square-root time dependence characteristic for diffusion, as evidenced by the linear L2 plot (inset) from which a diffusion coefficient of 20.8 × 10-8 cm2/s is derived.
1. Monoolein Cubic Phase as a Medium for Electrophoresis of Amphiphilc Molecules. Our main aim was to investigate if the monoolein cubic phase can be used as a medium for electrophoresis of membrane-bound molecules and to compare it in this respect to a conventional hydrogel of similar pore size. The type of agarose we use here is modified in order to obtain comparable pore size to the cubic phase and is well-suited as hydrogel control because it follows19 the Rodbard-ChrambachOgston model commonly used for conventional agarose gels. The average pore radius in the 5% gel used here (3.7 nm) is somewhat larger than the 2.5 nm of the cubic phase, but reproducing the cubic-phase value would require about 12% agarose gel, and such solutions are too viscous to handle. From Figure 2, it is clear that the monoolein cubic phase can be used for electrophoresis of amphiphilic molecules, whereas
the agarose hydrogel cannot. The monoolein cubic phase thus constitutes an important complement to conventional gels in the case of membrane-bound analytes. In fact, our observation of concomitant migration of membrane probes and oligonucleotides strongly support its bicontinuous structure. Furthermore, Figure 2 reveals a strong retardation of oligonucleotides compared to a conventional gel with similar pore size, which indicates that the electrophoretic conditions in the cubic phase are also unusual for hydrophilic molecules. It is therefore important to understand the mechanism by which both types of molecules navigate its maze of aqueous pores and curved membranes. One important piece of information on the mechanism of migration is the effect of field strength on the electrophoretic velocity. We have earlier shown that the velocity of DNA and DNA-protein complexes in porous matrixes sometimes exhibit nonlinear field dependence, due to perturbations of either the matrix structure20 or of the biomolecules,21,22 as a consequence of the strong electric forces that arise at elevated fields. The fact that the field-dependence is approximately linear for the membrane probes (Figure 5) and for the ss25 hairpin oligonucleotide14 therefore indicates that the monoolein cubic structure is not perturbed by their migration. Furthermore the membraneprobe mobility is independent of probe concentration (Figure 5), which shows that the molecules migrate independently of each other, as has already been shown for the oligonucleotides.14 Taken together, these results allow us to discuss the migration mechanism
(18) Ardhammar, M.; Lincoln, P.; Norde´n, B. J. Phys. Chem. B 2001, 105, 11363-11368. (19) Sanandaji, N.; Carlsson, N.; Voinova, M.; Åkerman, B. Electrophoresis 2006, in press.
(20) Svingen, R.; Åkerman, B. J. Phys. Chem. B 2004, 108, 2735-2743. (21) A° kerman. B. Electrophoresis 1996, 17, 1027-1036. (22) Svingen, R.; Takahashi, M.; A° kerman, B. J. Phys. Chem. B 2001, 105, 12879-12893.
Discussion
Electrophoresis of Membrane-Bound Probes and DNA
of both types of molecules in terms of individual analytes migrating in an unperturbed cubic phase. Electroosmosis can probably be neglected because the monoolein membrane is nonionic. 2. Nature of Membrane-Probe Migration. 2.1. Membrane Probe DiI Migrates Inserted into the Monoolein Membrane. The behavior of DiI strongly supports that it is inserted into the monoolein membrane while undergoing electrophoretic transport in the cubic phase. The alternative possibility, that DiI forms micelles or other water-soluble aggregates and migrates as such in the aqueous pores of the cubic phase, is unlikely on the basis of the following arguments. The membrane probe does not form a zone on the agarose hydrogel (Figure 2, right). If DiI did migrate as water-soluble aggregates in the cubic phase, it most likely should be able to do so in the agarose gel too because its average pore radius (3.7 nm) is larger but similar to the 2.5 nm radius pores of the monoolein cubic phase. Furthermore, micelles of DiI are not likely to form under our conditions because the Kraft temperature for similar surfactants, such as dioctadecyldimethylammonium chloride,23 is well above our experimental temperature. Second, the diffusion coefficients between 6.0 × 10-8 and 7.6 × 10-8 cm2/s we measured for DiI (in different gels at pH 4 and 5, respectively; Table 2) are close to the value (6.0 ( 0.5) × 10-8 cm2/s measured for two similar probes undergoing diffusion in a monoolein cubic phase membrane using a photobleaching approach.9 The agreement supports that the DiI probe is inserted in the monoolein membrane also when it has been introduced into the cubic phase by electrophoresis (at least after the field has been turned off). Indeed, closely related probes insert spontaneously into artificial membranes24 and bind to cell membranes rapidly (minutes) and with high affinity.25 Third, the environment of the dye during the electrophoretic motion can be probed by converting the mobility, µ, into the corresponding friction coefficient fµ ) Q/µ, where Q is the analyte charge. This friction coefficient during migration can be compared to the friction coefficient, fµ, derived from the diffusion coefficient, fD ) kT/D. If the underlying friction during migration is the same as during diffusion, then fµ ) fD. For DiI, the ratio fµ/fD equals 1.2 and 1.5 for the two gels at different pH’s (Table 2). The difference is likely to reflect the variation between different gel preparations (see above), so taken together, the values suggest that our best estimate is fµ/fD ) 1.35 ( 0.15. The ratio being close to 1 indicates that if the dye is inserted in the membrane during diffusion, as argued above, it is likely to also be the case during the electrophoretic migration. In particular, if the membrane probe did migrate in the aqueous phase the ratio should be on the order of 1:100 because the viscositiy of water is about a 100-fold lower than in the membrane.9 The possibility of migration in the aqueous phase is therefore all but ruled out by the fact that the experimental fµ/fD is larger than 1, if anything, and with an estimated uncertainty of about 0.15. 2.2. Mode of Migration of the Other Membrane Probes. The membrane probe DiO has identical hydrophobic tails as DiI and a similar fluorophore headgroup, so it seems likely that this molecule also migrate inserted the membrane. The ratio fµ/fD for DiO is even higher than for DiI (Table 2) and thus deviates even more from the value 1:100 that is expected if migration occurs in water but diffusion occurs in the membrane. Regarding C5HPA, we cannot calculate Dµ because the actual value of the (23) Jo¨nsson, B.; Lindman, B.; Holmberg, K.; Kronberg, B. In Surfactants and Polymers in Aqueous Solution; Wiley: New York, 1998; Chapter 3. (24) Klausner, R. D.; Wolf, D. E. Biochemistry 1980, 19, 6199-6203. (25) Sims, P. J.; Waggoner, A. S.; Wang, C. H.; Hoffman, J. F. Biochemistry 1974, 13, 3315-3330.
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charge, q, in the monoolein environment is not known (since the probe migrates in the opposite direction than expected). Ultimate support for membrane-bound migration of single-tail amphiphilic molecules will therefore require further investigations. However, our results indicate that the comparatively small dppz ligand of [Ru(phen)2Me2dppz]2+ is sufficient as hydrophobic anchor because the data supports that this dye migrates inserted in the membrane. The strongly enhanced fluorescence emission of this dye in the monoolein/water cubic phase compared to the monoolein-free control (see Supporting Information) strongly indicates that this dye is inserted into the monoolein membrane. Protective environments such as membranes are known18 to inhibit the proton-induced quenching of the [Ru(phen)2Me2dppz]2+ emission which occurs in water. Second, spectroscopic studies have shown that [Ru(phen)2Me2dppz]2+ binds to liposomes by inserting the dppz ligand into the membrane,18 so this is a likely binding mode also to the monoolein membrane, at least in absence of field. Third, the ratio fµ/fD is even larger than that for DiI and DiO, again supporting strongly that migration does not occur in the aqueous phase. 2.3. Contributions to Friction from Headgroup and Hydrophobic Anchor. It is noteworthy that the ratio fµ/fD increases systematically in the order DiI, DiO, and [Ru(phen)2Me2dppz]2+, i.e., in the order of increasing size of the headgroup (Table 2). This trend suggests that headgroup size is important, but the deviation in the ratio from 1 indicates that the analysis of the migration needs further development. For instance, it is counterintuitive that DiO has a higher mobility than DiI, although the two analytes have indentical membrane-anchoring parts and the hydrophilic part of DiO is smaller than that of DiI. This observation, together with the unexpected migration direction of C5-HPA, led us to use the diffusion data rather than mobility when discussing the effect of the headgroups on the friction experienced by membrane-bound probes. Notably, DiI is slower than DiO by a factor of 2, although both have identical hydrophobic anchors (two C18 chains). On the other hand, DiO, C5-HPA, and [Ru(phen)2Me2dppz]2+ differ substantially in number and size of the hydrophobic tails and still their diffusion coefficients differ by less than 35%. These observations suggest that the size of the hydrophobic part plays a comparatively small role for the underlying friction. By contrast, diffusion rates decrease with increasing size of the headgroup, being smallest for DiI and largest for [Ru(phen)2Me2dppz]2+ (Table 2). This observation indicates that the size of the hydrophilic part is more important for the total friction than is the size of the hydrophobic anchor. One interesting possibility is that steric or hydrodynamic interactions between the headgroups and the walls of the aqueous pores is the dominating size effect for analyte friction during motion of membrane-bound molecules through the cubic phase. 3. Nature of Oligonucleotide Migration. The possibility of a size-dependent friction between hydrophilic parts of membranebound molecules and the monoolein cubic matrix begs the question how water-soluble molecules interact with the lipid membrane. Experimentally, the issue of hydrofil-friction can be addressed by using water-soluble oligonucleotides of different sizes as probes. The strong retardation of the ss25 hairpin in the monoolein cubic phase compared to the conventional gel (Figure 2) of similar pore size indicates that water-soluble analytes are retarded by different mechanisms in the two gels. This is not surprising given the different natures of the confining matrix, i.e., a continuous lipid wall in the cubic phase, compared to a network of discrete fibers in the agarose gel. Recent modeling of the DNA-matrix interactions in these two types of structures19
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supports that a continuous porous maze confers much stronger retardation to a oligonucleotide of nanometer size than does a fibrous matrix of the of the same average pore size. Our study of oligonucleotides of various sizes and secondary structure14 showed a marked effect of oligonucleotide size (and flexibility) on the electrophoretic velocity. Furthermore, single-stranded forms migrated faster than the corresponding double-stranded form, which suggests that hydrophobic interactions (for instance, between membrane and exposed bases) are weak even for the single-stranded form.
Concluding Remarks By studying amphiphilic and hydrophilic molecules separately, we have demonstrated that the bicontinuous nature of the monoolein cubic phase provides both a membrane in which membrane-bound molecules can migrate and aqueous pores with size-separation capability for hydrophilic molecules of nanometer dimensions.14 Cubic phases may therefore offer a larger spectrum of separation motifs for membrane-bound analytes than planar immobilized membranes6,26 because sieving on the hydrophilic parts in the aqueous pores is also likely to affect analyte friction. However, the broad zones and limitation to acidic pH in the present monoolein system have to be overcome before efficient separation can be realized. There is also the issue of polycristallinity. Studies in a cubic phase formed by the amphiphilic polymer Pluronic F127 and (26) van Oudenaarden, A.; Boxer, S. G. Science 1999, 285, 1046-1048.
Carlsson et al.
water show that DNA migration in polycrystalline liquid crystals may occur along grain boundaries rather than through the actual crystallites.20 The very strong retardation of the oligonucleotides in the monoolein cubic phase (compared to a conventional polymeric gel) supports that they do migrate in the actual pores of the cubic crystal, and that grain-boundary migration is limited to ferrying between grains. The low water solubility of the membrane probes may render them more sensitive to discontinuities in the membrane at grain boundaries. This may explain the lower degree of reproducibility in their transport properties between independently prepared cubic phases. It is also noteworthy that the diffusion coefficient we measure for the membrane probe DiI over millimeter distances agrees with values measured over micrometer distances by spectroscopic techniques.9 This suggests that if heterogeneities do exist in the cubic gels, for instance, due to polycristallinity, they are either smaller than a micrometer or larger than a millimeter. Acknowledgment. We thank Peter Nollert and Staffan Wall for helpful discussions, Per Lincoln for the kind gift of [Ru(phen)2Me2dppz]2+, and the Swedish Research Council for financial support. Supporting Information Available: Diffusion of [Ru(phen)2Me2dppz]2+ into the cubic phase as a function of time, and theoretical distribution of pore radius of the aqueous channels in the diamond-type cubic phase of monoolein. This material is available free of charge via the Internet at http://pubs.acs.org. LA052086L
