J . Phys. Chem. 1994,98, 1732-1738
1732
Kinetics of the Solubilization of Styryl Dye Aggregates by Lipid Vesicles Athina Zouni, Ronald J. Clarke,' and Josef F. Holzwarth Fritz-Haber-Institut der Max-Planck-Gesellschaft, Faradayweg 4-6, 0-141 95 Berlin, Germany Received: July 8, 1993; In Final Form: November 9, 1993'
The interaction of the potential-sensitive styryl dye R H 4 2 1 with dimyristoylphosphatidylcholinevesicles has been investigated above and below the gel-liquid crystal-phase transition temperature by the fluorescence stopped-flow method. The observed kinetic behavior is consistent with a disaggregation of dye aggregates in the aqueous solution and the incorporation of dye monomers into the vesicle membrane. Upon incorporation the dye undergoes a decrease in its pK, of over 2 p H units. This can be attributed to the change of the electronic environment of the dye. The magnitude of the pKa shift is discussed in terms of the local membrane dielectric constant, the dipole potential, and possible dye reorientation, which may all affect the free energy change of the protonation reaction.
Introduction Potential-sensitive fluorescent styryl dyes are presently being widely employed for the visualization of voltage transients in membrane preparations.' They are particularly useful in work with cells, cell organelles, or lipid vesicles, where the measurement of the membrane potential difference with microelectrodes is not possible because of their small size. The kinetics and mechanism of the dyes' response to voltage changes are, however, still not fully understood. An important step in the elucidation of the mechanism of the voltage response is to discover how the dyes are incorporated into the membrane and where they are localized when bound. In previous publications the absorbance and fluorescence properties of the styryl dye RH421 (see Figure 1) have been investigated in aqueous solution and in the membrane,2 and its dynamics within the membrane have been studied.3 The absorbance and fluorescence emission spectra of the dye when bound to dimyristoylphosphatidylcholine (DMPC) vesicles are shown in Figure 2. It was found that the dye aggregates strongly in aqueous solution. The aggregation causes a dramatic decrease in the fluorescent quantum yield. In the presence of lipid vesicles the fluorescence is greatly enhanced due to the dissolution of the aqueous-phase aggregates. Here we report a fluorescence stoppedflow investigation of the kinetics of interaction of the dye with DMPC vesicles. In combination with pH titrations of bound dye, the results obtained yield information concerning the degree of aggregation of the dye within the membrane and the polarity of its environment. Materials and Methods
N-(4-Sulfobutyl)-4-(4-@-(dipentylamino)phenyl) butadieny1)pyridinium inner salt (RH421) was obtained from Molecular Probes (Eugene, OR) and was used without further purification. It has been previously observed4 that such diene-linked dyes can exist in two stereochemical forms (E,E and E,Z) with different fluorescence and absorbance properties. The fluorescence emission spectrum of our dye in ethanolic solution showed only a single peak with a maximum at 695 nm, which was independent of the excitation wavelength. The longest wavelength absorbance band occurred at a A, of 520 nm. Using a Bruker AMX-500 spectrometer the 1H N M R spectrum of the dye was measured in deuterated chloroform. The coupling constants of the olefinic protons were found to be 15.0 f 0.2 Hz,indicating the E,E (all * To whom correspondence should be addressed. 0
Abstract published in Aduance ACS Abstracts. January 15,
1994.
0022-365419412098- 1732$04.50/0
R /N W R = C H CH CH CH CH
2 2 2 2 3
Figure 1. Structure of RH421. 015
c Ln 3
3F
Abs.
ii
010
0.05
I
Figure 2. Absorbance (A) and fluorescenceemission(F) spectra of RH421 bound to dimyristoylphosphatidylcholine (DMPC) vesicles. To a suspension of vesicles containing 500 pM DMPC (absorbance) or 83 pM (fluorescence)in aqueous buffer (30 mM tris, 1 mM EDTA, 150 mM
NaC1, pH 7.2), 6.4 pM RH421 (absorbance),or 0.63 pM (fluorescence) were added from an ethanolicstock solution; 0.5-cm-path-lengthquartz cuvettes were used, T = 30 OC, bandwidths = 5 nm. The fluorescence wasexcited at 460nm (+GG435 cutoff filter). Thefluorescencespectrum
is in arbitrary units, and its intensityhas been normalized to the maximum absorbance of the absorbance spectrum.
trans) isomeric form.5 A series of stock solutions were prepared in ethanol. For spectral measurements 5 pL of an ethanolic dye solution was added to a 0.5-cm-path-length quartz cuvette containing 1 mL of aqueous solvent. The final solutions measured thus contained a small and constant percentage of 0.5% ethanol. The molar absorptivity was simply calculated by dividing the measured absorbance by the dye concentration and the cuvette path length. Dimyristoylphosphatidylcholine (DMPC) was obtained from Avanti Polar Lipids (Alabaster, AL). DMPC unilamellar vesicles were prepared according to a modification of the ethanol injection 0 1994 American Chemical Society
Solubilization of Styryl Dye Aggregates method of Batzri and KomG8 A 30 mM solution of DMPC (1 mL) in ethanol was injected slowly over 15 min with continuous stirring into 10 mL of buffer solution a t 30 "C. The final solution contained no detectable trace of ethanol, i.e., [ethanol] I10pM, according to an NADH/alcohol dehydrogenase enzymatic assay (Boehringer, Mannheim). Dialysis tubing was purchased from Medicell International (London, UK). The phospholipid content of the vesicle suspensions was determined by the phospholipid B test from Wako (News, Germany). The vesicles produced were unilamellar with external diameters in the range 50-100 nm, as determined by cryoelectronmicroscopy and confirmed by dynamic light scattering. The majority of kinetic measurements with the vesicles were performed in a buffer containing 30 mM tris, 1 mM EDTA and 150 mM NaC1. The pH of the buffer was adjusted to pH 7.2 with HCl. All solutions were prepared using triply distilled water. The origins of the various reagents used were as follows: tris[ (hydroxymethy1)aminolmethane (99.9%, Sigma), EDTA (99%, Sigma), NaCl (Suprapur, Merck), HCl(O.1 M Titrisol solution, Merck), ethanol (analytical grade, Merck), glycine (99%, Merck), and sodium dodecyl sulfate (99%, Sigma). Absorbance measurements were performed with a Shimadzu UV-2100 UV-visible recording spectrophotometer using a bandwidth of 5 nm. Steady-state fluorescence measurements were recorded with a Shimadzu RF-5000 recording spectrofluorophotometer using bandwidths of 5 nm for both the excitation and emission monochromators. To minimize contributions from scattering of the exciting light and higher order wavelengths a glass cutoff filter was used in front of the excitation monochromator. Measurements of the pH dependence of the dye absorbance and fluorescence were carried out using a series of 0.05 M glycine buffers of varying pH.9 NaCl (1 .O M) was added to the vesicle dialysis medium and to the glycine buffers in order to prevent variations in the ionic strength on either side of the membrane. The determination of the pKa values was carried out by fitting the data obtained from the pH titrations to the HendersonHasselbalch equationlo using the commercially available nonlinear least-squares program ENZFITTER. The program was purchased from Biosoft (Cambridge, UK) and was run on a IBMAT/386 compatible personal computer (mey-Soft, Berlin, Germany). The kinetics of the interaction of RH421 with dimyristoylphosphatidylcholine vesicles were investigated using a homemade fluorescence stopped-flow apparatus.ll A 150-W Xe arc lamp (Osram, Berlin, FRG) was used as the exciting light source. This was powered by a T N X 150 power supply (Heinzinger, Rosenheim, FRG). The dye/vesicle suspensions were excited in the wavelength range 360-560 nm by using a BG18 glass filter (Schott, Mainz, FRG) and a 500-nm (SWP) short-wavelength pass filter (Oriel, Darmstadt, FRG). The fluorescence emission was detected above 590 nm by using an OG590 glass cutoff filter (Schott, Mainz, FRG). The output signal of the photomultiplier (EM1 9558 QA) was displayed on a 7603 oscilloscope with a 7A22 plug-in differential amplifier (Tektronix, Beaverton, OR) and stored in binary form using a Biomation 1010 transient recorder (Gould Inc., Santa Clara, CA). The overall bandwidth for the detection circuit was 1 kHz. The analysis of the data was carried out using a HP9845B computer (Hewlett-Packard, Loveland, CO). At least four experimental traces were averaged for each pair of dye and vesicle solutions. The solutions in the drive syringes were equilibrated to the required temperature prior to each experiment using a KT33 thermostat (Haake, Berlin, FRG). The dead time of the stopped-flow unit was determined to be approximately 6 ms. Computer simulations of the stopped-flow transients were performed using a Digital VAX computer. The initial concentrations of all the dye species were calculated using a combination
The Journal of Physical Chemistry, Vol. 98, No. 6, 1994 1733 of the methods of linear interpolation, extrapolation and bisection within a subroutine of the Numerical Algorithms Group (NAG) Fortran Library. The differential forms of the rate equations were then integrated using backward differentiation formulas12 within a further subroutine of the NAG Library.
Theory The pKa's of membrane-bound groups are often different from the corresponding value for the isolated group in aqueous s o l ~ t i o n . ~ ~ , From ~ ~ , ~the ~ difference - ~ ~ ~ ~ Oin the pKa values in water and in the membrane the energy difference for the deprotonation reaction can be calculated as follows: AGO, - AGO, = 2 . 3 0 3 R T ( p c - p c ) where AGO, and AGO, are the standard free energies for deprotonation in the membrane and in water, respectively, and p c and p c are the corresponding values of pKa in the membrane and in water. Several reasons could account for a difference in the energetics of the deprotonation reaction in the two environments. First, a membrane boundary potential, Ub, could lead to an attraction or a repulsion of H30+ions. Second, the difference in the dielectric constant of the aqueous and membrane phases could cause a difference in the Born energy of H30+ions in the two phases. The electrostatic energy, Eel, due to the presence of a membrane boundary potential is given by
where z is the valence of the ion (+1 for HsO+) and F i s Faraday's constant. here represents the total boundary potential from the position within the membrane where the protonation occurs to the bulk solution, Le., the diffuse double-layer region adjacent to the membrane is included. The Born energy due to the difference in dielectric constant of the medium is given by
(3) where eo is the charge on an electron, L is Avogadro's constant, eo is the permittivity of a vacuum, r is the radius of the ion, e, is the local dielectric constant of the membrane, and e, is the dielectric constant of water. Considering at this stage the energies of the H30+ions alone, the energy difference AGO, - AGO, can be equated to the negative amount of energy required for the transfer of a H30+ ion into the membrane. Combining the electrostatic and Born energy contributions and substitution into eq 1, it can be shown that the pKa shift is given by
For H30+ ions r = 0.17 X m13J5and e, = 78. From experimental measurements of the pKa shift eq 4 thus allows the calculation of t, if the value of ub is known or alternatively the calculation of ub if the value of e, is known. An additional contribution to the energy of an ion within a lipid membrane which has not been included in eq 4 is the image energy due to polarization forces at the interface. Flewelling and Hubbell20 have derived an equation for the combined Born and image energies based on a previous solution of Neumcke and Lauger.21 The calculation requires a knowledge of the distance of the ion from the membrane-aqueous solution interface. Since for small ions the Born energy contribution is far in excess of the image energy22 the error introduced by neglecting the image energy is minimal. So far the pKa shift has been interpreted solely in terms of the energy of transfer of H30+ions into the membrane. A further contribution to the pKa shift could arise because of the anisotropic
1734 The Journal of Physical Chemistry, Vol. 98, No. 6, 1994 I
I
e
0 0
2
4
6
e
l
10
PH
Figure 3. Variation of the molar absorptivity, e, of RH421 at 500 nm in the presence of 500 fiM of dimyristoylphosphatidylcholinein the form of unilamellar vesiclesand 1.0 M NaCl asa function of pH. The titration was performed in 0.05 M glycine/HCI buffer solutions. The solid curve represents a nonlinear fit of the data to the Henderson-Hasselbalch equation. [RH4211 = 6.4 pM,bandwidth = 5 nm, T = 30 "C.
nature of the membrane. Any dye reorientation within the membrane occurring concomitantly with deprotonation may result in a variation in the apparent pKa because of the difference in energies of the orientations of the protonated and deprotonated forms. This effect is not included in eq 4. Nevertheless, as will be shown later, eq 4 still allows a limiting value of the membrane dielectric constant to be calculated.
Results Acid-Base Properties of RH421. The dye possesses two nitrogen atoms, one of which could be protonated. Protonation causes a blue shift of the absorption spectrum of approximately 100 nm.2 Similarly, a blue shift of the fluorescence emission spectrum also occurs. The variation of the molar absorptivity of the dye at 500 nm in the presence of DMPC vesicles with pH is shown in Figure 3. The observation wavelength was chosen so as to minimize interference from light scattering. The data have been fitted to the Henderson-Hasselbalch equation as described in the Materials and Methods section and a pK, of 2.1 f 0.1 has been obtained. The titration could not be extended to pH values below 1.8 because of protonation of the phosphate headgroup of the lipid and the consequent phase transition of the bilayer,*6 which causes a significant increase in the turbidity of the suspension. pH titration using fluorescence detection resulted in an identical value of the pKa. It can thus be assumed that no changein the protonationstateofthedyeoccurs within thelifetime of the excited state. Protonation of the sulfonate group of the dye would be expected at the pH of approximately 1.5,23but because it is not part of the conjugated system no significant spectral changes are likely. For RH421 in aqueous solution the pK, has previously been determined2 to be 4.9. Thus, there is a decrease in the pKa of 2.2 pH units on incorporation into the DMPC membrane. Since DMPC is a zwitterionic lipid, it is likely that any electrostatic surface potential is very small. The high ionic strength in the aqueous solution would in addition effectively screen any surface charges." An electrostatic repulsion of H30+ions away from the membrane surface can, therefore, be excluded. The shift in pKa must be due to effects within the membrane. The fact that the pKa decreases in the membrane means that the protonated form of the dye has a higher energy within the membrane than the deprotonated form. Considering eq 4 in the Theory section, one can think of two possible causes of the pK, shift. First, the membrane undoubtedly has a lower dielectric constant than the aqueous solution. The Born energy for the transfer of H3O+ into the membrane would, therefore, cause a decrease in the pKa.
Zouni et al. Second, a boundary potential may be present which also increases the energy of H30+ions within the membrane. As stated above, nosurface potential can be present. Theboundarypotentialwould have to exist totally within the membrane. By considering the interaction of hydrophobic ions with lipid membranes Flewelling and HubbePo have calculated that phosphatidylcholine bilayers should possess a dipole potential of approximately 240 mV between the center of the membrane and each membrane-solution interface. The dipole potential is presumed to arise from the orientation of the ester groups which link the two fatty acid chains to the glycerol backbone. The positive value of the dipole potential would, therefore, hinder the penetration of H30+ ions into the membrane. In actual fact, it is likely that both effects, Born energy and dipole potential, contribute to the pK,shift. A specific effect due to a localization of the protonatable group close to the discrete positive charge of the nitrogen of the phosphatidylcholiae headgroup is a further possibility. Finally, a change in the pK, could also occur if the dye undergoes a reorientation within the membrane upon protonation. A decrease in pK, would beobserved if the orientation of the deprotonated form has a lower energy than that of the protonated form. This is a distinct possibility, since the protonated form of the dye is more highly charged than the deprotonated form. The loss of a proton may, therefore, allow increased penetration of the dye into the membrane and stabilization via hydrophobic interactions. The fact that the absorbance spectrum of deprotonated dye is not greatly shifted on transfer from water into the membrane argues, however, against penetration into the apolar interior of the membrane. The value of in eq 4 is given by the portion of the dipole potential between the position in the membrane where the protonation occurs and the interface, Le., it is probably significantly less than 240 mV. Since its exact value is not known, we shall set ub equal to zero in eq 4, so that a lower limit of the local membrane dielectric constant, em, can be calculated from the pK, shift. For the purposes of the calculation the possible effect of dye reorientation is likewise ignored. Accordingly, one finds that t, L 23. Such a value is consistent with the localization of the chromophore in the headgroup region of the membrane.20 Stopped-FlowKinetics. When dye is rapidly mixed with vesicles in the stopped-flow apparatus, as described in the Materials and Methods section, an overall increase in fluorescence is observed. The form of the observed kinetic trace is very dependent on the dye concentration and also on the temperature a t which the experiment is performed (see Figure 4). At low dye concentrations there is a rapid increase in fluorescence which is followed by a decrease. Such kinetic behavior has also been observed for the binding of an oxonol dye to lipid ve~icles.~4 It has been attributed to the binding of dye to the membrane, which leads initially to an increase in fluorescence, but as time goes on and more dye binds, a fluorescence quenching occurs due to the interaction between membrane-bound dye molecules. The quenching may be due to an inner filter effect, Le., self-absorption of the fluorescence emission, or to radiationless resonance energy transfer. At higher dye concentrations a further kinetic phase appears, which is characterized by a slow increase in fluorescence. The relative amplitudes of the fast and slow phases depends on the dye concentration and on the temperature. The amplitude of the slow phase increases a t higher dye concentrations and below the lipid phase transition temperature of 23 'C it is more pronounced than above. The amplitude of the slow phase is also significantly enhanced by an increase in the ionic strength (see Figure 5). The effect of salt is to increase the degree of aggregation of the dye in the aqueous solution.2 Dye aggregates have a much lower fluorescence than dye monomers.2.3 The slow kinetic phase can, thus, be attributed to the disaggregation of the dye in theaqueous phase. This is driven by the removal of dye monomers from the solution by their binding to the membrane.
The Journal of Physical Chemistry, Vol. 98, No. 6, 1994 1735
Solubilization of Styryl Dye Aggregates
4
3.0 1
FlmV FlmV 1.5
1
0.0
0.4
os
I
1.2
tlsec
- 3.0
1.6
7
t Ok
OD
0.8
1.2
tlsec
1.6
I
I 0
2
4
6
8 t/sec
10
c /
- 120
t
I
Figure 4. Experimental fluorescence stopped-flow traces. The final lipid concentration after mixing was 155 pM in all cases. The dye concentrations after mixing and the temperatures were (A) 0.064 pM,21 O C , (B) 1.28 pM,21 "C, (C) 12.78 pM,21 OC,(D)0.064 pM,30 O C , (E)1.28 pM,30 O C , and (F) 12.78 pM,30 OC. The reaction between the dye and the vesicles begins at the point marked by the arrow. The portion of the trace before the arrow is the pretrigger region.
Very similar kinetic behaviour involving two kinetic phases was also observed on mixing the dye with sodium dodecyl sulfate micelles ([SDS] = 5 X 10-3 M). This shows that the slow process is unlikely to be related to a reaction within the membrane. Vesicles and micelles are both capable of removing dye monomers from solution and inducing dye disaggregation in the aqueous phase. Spectral changes due to micelle-induced dye disaggregation have been reported for cyanine and rhodamine dyes by various auth0rs.~5 It was found that the time course of the slow process in the presence of a constant concentration of DMPC vesicles was best described by a sum of two exponential functions. Irrespective of the dye concentration, the major component (approximately 90% of the total amplitude) has a relaxation time of 0.8 f 0.1 s-l at
21 OC and 2.3 f 0.2 s-1 a t 30 OC. The minor component had a reciprocal relaxation time of 0.18 f 0.07 s-l at 21 "C and 0.37 f 0.08 s-1 at 30 OC. The fact that the slow phase is multiexponential indicates that the disaggregation process involvesseveral reaction steps. Apart from an increase in the rate of the slow phase at the higher temperature, the amplitude of the fast phase increases significantly in comparison to the slow phase above the phase transition temperature of 23 OC (see Figure 4). To test the validity of the proposed mechanism, Le., disaggregation followed by incorporation of dye monomers into the membrane, computer simulations have been carried out. It is possible that very large aggregates are formed. For the purpose of the simulation, however, the aggregation has been limited to the formation of tetramers. The following reaction scheme has
Zouni et al.
1736 The Journal of Physical Chemistry, Vol. 98, No. 6, 1994
dN3/dr = k3N1N2c*, - k4N3- ksNlN3c*,
+ k6N4
(9) N1, N2, N3, and N4 represent the number of dye monomers, dimers, trimers, and tetramers, respectively, in the aqueous solution per vesicle. r represents the number of occupied binding sites per vesicle, and C*Vis the total concentration ofvesicles. A differential equation for N4 is not required, since it can be calculated by utilizing the mass conservation law: -15
-
c*D/c*V = N,
(10) wherec*D is the totaldyeconcentration. The initialconcentrations of all the individual dye species prior to the addition of vesicles for eachvalue of c*D were calculated as described in the Materials and Methods section by solving the following polynomial for c1, the concentration of monomers:
-30 0
0.4
0.2
0.8
0.6
+ 2N2 + 3N3 + 4N4 + r
tlsec
+ + 2K,cI2 + 3K,K2cI3+ 4K,K2K3cl4= 0
- c * ~ c1
(1 1)
K1, K2, and K3 represent the equilibrium constants for the aggregation steps, Le., K1 = kl/k2, K2 = kJk4 and K3 = k~/k6. The concentrations of dimers, trimers, and tetramers are then given by
WmV
- 10 -20
c3 = K,K2cI3
(13)
c4 = K,K2K3cI4
(14)
-30
-40
i
0
Figure 5. Experimental fluorescence stopped-flow traces for dye and vesicle solutions in (A) water and (B) in 2 M NaCl solution. The dye and lipid concentrationsafter mixing were 1.28 and 155 pM,respectively; T = 21 oc. been used as a basis:
D,+ V
nk+
DV
The values of N1, N2,N3, and N4 are simply obtained by dividing c1, c2, c3, and c4 by C*V. The results of such simulations of fluorescence stopped-flow experiments for a range of dye concentrations are shown in Figure 6. The complication of fluorescence quenching in the membrane phase has been treated as described elsewhere27 by introducing a factor z which is the number of binding sites adjacent to a bound dye molecule which must be free in order to prevent quenching. The fluorescence, F, in arbitrary units was calculated at any point in time according to the following equations:
k-
1,
k3
D,+ D2+ D, k4
ks
D,+ D, e D4 k6
D1, D2, D3,and D4 represent dye monomers, dimers, trimers and tetramers in the aqueous solution. V represents a vesicle with n binding sites for dye monomers. The aggregation has been assumed to be a stepwise process with the continual addition of dye monomers. This is consistent with the formation of dye stacks.26 The rate constants k+ and k- refer to interaction of the dye with a single dye binding site. The kinetic theory of the binding reaction has been treated in detail elsewhere.2' The overall reaction scheme can be described by the following series of coupled differential rate equations: dr/dr = nk+Nlc*, - (k+Nlc*,
+
+ kJr
(6)
dN,/dt = -nk,N,c*, (k,Nlc*, + kJr- 2k,NI2c*, + 2k2N2 - k3NINz~*v k4N3 - ksN,N3~*, k6N4 (7)
+
dN2/dt = kINl2c*, - k2N2- k3N1N2c*,
+
+ k4N3
(8)
wherefi, f and f 7 are the values of the fluorescence intensity per mole of dye in the monomer form in aqueous solution, in isolated form in the membrane, and in quenched form in the membrane, respectively. Pi is the probability that a dye molecule in the membrane is not quenched. The fluorescence intensities of aggregated forms of the dye in aqueous solution have been assumed to be zero. The derivations of eqs 15 and 16 have been carried out in an analogous fashion to those of eqs 20 and 21 in ref 24. The values of the parameters required for the simulations were varied in order to obtain the best agreement with the experimental traces. The equilibrium constants used, however, are all consistent with previously determined experimental values.2J It can be seen that the form of the experimental traces is reproduced qualitatively by the simulations. An extension of the model to dye aggregates larger than a tetramer may provide a better agreement, but the fact that the dye concentration dependence of the curve form can be simulated supports the basic validity of the proposed mechanism.
Discussion A mechanism has been proposed for the interaction of the dye RH421 with lipidvesicles. It involves the binding of dye monomers to the membrane and the breakup of dye aggregates in the aqueous solution. Both binding and disaggregation contribute to the increase in fluorescence on addition of vesicles. The relative
The Journal of Physical Chemistry, Vol. 98, No. 6,1994 1737
Solubilization of Styryl Dye Aggregates F1
F1
-
1.5
2.1
-
2.0
.-
I.'
1.8
1.0 1.6
1
0
2.1
2
4
6
a
10
tlsec
t / -
I
4.0
3.5
3.0 0
1
4
6
8
2
1
1
4
1
1
6
1
1
B
1
1
10
l
I
12
I
I
I4
I
I
16
I
l
18
I
!
20
t/sec
Figure 7. Computer simulationsof fluorescence stopped-flow traces at a lipid concentration of 155 pM after mixing and a dye concentration after mixing of 1.28 pM. The number of binding sites per vesicle used for the calculationwere 1 X 104 (above) and 3 X lo4(below). The values of the other parameters required for the simulations were as given in Figure 5.
1.5 0
1
10
tlsec
Figure 6. Computer simulations of fluorescence stopped-flow traces at a constant lipid concentration after mixing of 155 pM and dye concentrationsafter mixing of (a) 0.064pM, (b) 1.28 pM, and (c) 12.78 pM. The simulationswere carried out using eqs 6-16. The values of the parameters used were as follows: number of lipid molecules per vesicle = 4 X 105, k+ = 4 X lo6 M-I s-I, k- = 5 s-I, kl = ks = kg = 2 X lo7 M-I s-l, k2 = kd = k6 = 0.9 s-l, n = 1 X lo4 binding sites per vesicle,fi = 4 X 10' (arbitrary units) M-I,fli = 2 X lo8 (arbitrary units) M-I,fP = 0 (arbitrary units) M-I and z = 200.
contributions depend on the dye concentration and on the temperature. At high dye concentrations, Le., in the micromolar range, the disaggregation becomes more prominent because of the initial presence of large numbers of aggregates in the aqueous phase. Above the gel-liquid crystal phase transition temperature the fluorescence change due to the binding becomes more significant. Previously it has been found that the binding affinity, nK, of the dye increases above the phase transition t e m p e r a t ~ r e . ~ This could be due either to an increase in the binding constant, K, or to an increase in the number of binding sites per vesicle, n. Computer simulations, however, show that thechange in kinetic behavior above the phase transition temperature is best explained by an increase in the number of binding sites (see Figure 7 and cf. Figure 4). An increase in K causes an increase in the fluorescence change due to the disaggregation step, which disagrees with the observed experimental behavior. On the basis of the pK, shift of membrane-bound dye a dielectric constant 2 2 3 has been calculated. This suggests a localization of the protonated chromophore in the lipid headgroup region. The hydrocarbon interior of the membrane is expectedZoto have a dielectric constant of approximately 2-3. The pK, shift of the
dye when bound to DOPC vesicles has been investigated previously.2 There it was found that the pK, decreases by 0.8 pH units, i.e., a much smaller shift than that observed in the case of DMPC. The pK, shift corresponds to a dielectric constant 141. Such a value is alsoconsistent with a localization in the headgroup region of the membrane, but the fact that the pK, shifts are different in the two lipids indicates that the dye experiences a different electronic environment. This could be due to a number of reasons. First, the depth of penetration of the dye into the membrane may be greater for DMPC than DOPC. Second, if the penetration depths are the same, the local polarity of the membrane may be greater for DOPC than DMPC. A third explanation could be that the dipole potential is less for DOPC than DMPC. A higher polarity or a decreased dipole potential could both be due to the greater fluidity of the DOPC membrane. The greater fluidity would randomize the orientation of the ester groups and perhaps allow greater access of water into the membrane. If the dye undergoes reorientation upon protonation, one can imagine a final possibility, i.e., the orientation of the deprotonated form is energetically more strongly favored over that of the protonated form in DMPC than DOPC. This could also be a consequence of the greater fluidity of the DOPC membrane. If the exact location of such dyes could be determined, the results demonstrate the possibility of using the pK, shifts of probe molecules to determine local membrane electrical properties. In particular, the relationship between membrane fluidity and the dipole potential could be of significance for the mechanism and regulation of ion transport across cell membranes. With respect to the voltage sensitivity of the dye, it is possible that under certain concentration conditions the aggregation in the aqueous solution plays a role. Initially it was thought that such dyes responded to changes in the membrane potential via an electrochromic mechanism. It has since been found, however, that other mechanisms must be involved.2-28 A mechanism involving a potential-dependent change in the degree of dye aggregation in the aqueous phase has been proposed for other dye molecules.29 Acknowledgment. We thank Frau Britta Kryszak and Herrn Uwe Marx for assistance with the p H titrations and preliminary stopped-flow measurements, Dr. Andreas Schafer for the measurement of N M R spectra, and Dr. Paul Fletcher and Herrn
1738 The Journal of Physical Chemistry, Vol. 98, No. 6, 1994
Joachim Frank for valuable discussions. A.Z. is grateful for financial support from the Max-Planck-Gesellschaft. R.J.C. acknowledges with gratitude financial support from the Stipendien-Fonds der Chemischen Ind. We are grateful to a reviewer for drawing to our attention the possibility of dye reorientation as a contributing factor to the pKa shift. References and Notes (1) (a) Grinvald, A.; Salzberg, B. M.; Lev-Ram, V.; Hildesheim, R. Eiophys. J . 1987, 51, 643-651. (b) Miiller, W.; Windisch, H.; Tritthart, H. A. Eur. Eiophys. J . 1986, 14, 103-111. (c) Chien, C.-B.; Pine, J. Eiophys. J . 1991,60,697-711. (d) Ehrenberg, B.; Meiri, Z.; Loew, L. M. Photochem. Photobiol. 1984,39,199-205. (e) Manis, P. B.; Freeman, J. A. J . Neurosci. 1988,8, 383-394. (f) Klodos, I.; Forbush, B. I11 J . Gen. Physiol. 1988,92, 46a (abstr). (9) Biihler, R.; Stiirmer, W.; Apell, H.-J.;Liuger, P. J . Membr. Biol. 1991,121,141-161. (h) Stiirmer, W.; Biihler, R.;Apell, H.-J.;Liuger, P. J . Membr. Biol. 1991,121, 163-176. (i) Heberle, J.; Dencher, N. A. Proc. Natl. Acad. Sci. U.S.A.1992,89,5996-6000. (j) Fromherz, P.; Dambacher, K. H.; Ephardt, H.; Lambacher, A,; Miiller, C. 0.;Neigl, R.; Schaden, H.; Vetter, T. Ber. Bunsen-Ges. Phys. Chem. 1991,95, 1333-1345. Schenk, 0.; (2) Clarke, R. J.; Schrimpf, P.; Schdneich, M. Eiochim. Eiophys. Acta 1992, 1112, 142-152. (3) Zouni,A.;Clarke,R. J.;Visser,A.J. W.G.;Visser,N.V.;Holzwarth, J. F. Eiochim. Eiophys. Acta, in press. (4) (a) Hassner, A.; Birnbaum, D.; Loew, L. M. J . Org. Chem. 1984,49, 2546-2551. (b) Biihler, R.; Stiirmer, W.; Apell, H.-J.; Liuger, P. J . Membr. Biol. 1991, 121, 141-161. (5) Streitweiser, A.; Heathcock, C. H.; Kosower, E. M. Introduction to Organic Chemistry, 4th ed.; Macmillan: New York, 1992; pp 353-354. (6) Batzri, S.; Korn, E. D. Biochim. Biophys. Acta 1973, 298, 10151019. (7) Kremer, J. M. H.; Esker, M. W. J. v. D.; Pathmamanoharan, C.; Wiersema, P. H. Biochemistry 1977, 16, 3932-3935. (8) Holzwarth, J. F. Faraday Discuss. Chem. SOC.1986, 81, 74-76. (9) Dawson, R. M. C.; Elliott, D. C.; Elliott, W. H.;Jones, K. M. InDara for Biochemical Research, 3rd ed.; Oxford University Press: Oxford, 1986; p 426. (10) Stryer, L. Biochemistry, 3rd ed.; W. H. Freeman: San Francisco, 1988; pp 41-42. (1 1) (a) Frank, J. Diploma Thesis; Technical University, Berlin, 1991, pp 40-42. (b) Westerhausen, J. Diploma Thesis; Free University, Berlin, 1979.
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