J. Phys. Chem. 1995, 99, 16149-16154
16149
Kinetics of In-Plane Photoinduced Electron Transfer between Lipid-Functionalized Pyrene and Alkyl-Substituted Dipyridinium Cation in Lipid Vesicles and Langmuir Films L. Li and L. K. Patterson* Radiation Laboratory and Department of Chemistry, University of Notre Dame, Notre Dame, Indiana 46556 Received: March IO, 1995@
Measurements of electron transfer between singlet excited lipid-functionalized pyrene and alkyl-substituted dipyridinium cation have been carried out in the lipid-water interfacial region of both spread monolayers and vesicles. These studies have been conducted to evaluate the effects of limited dimensionality on the kinetics involved. Zwitterionic dioleoylphosphatidylcholine (DOL) and negatively charged dioleoylphosphatidylglycerol (DOPG) have been used as the lipid hosts in this study. The functionalized compounds were 1-(l0-(6(8)-octadecylpyrenyl)decanoyl)-2-hexanoylphosphatidylcholine (OPyPC), utilized as the donor, and an amphiphilic acceptor, 4-(N-hexadecylpyridinium-4-yl)pyridinechloride (CB+). It has been shown by monolayer compression studies that the rates of forward transfer in spread monolayers are sensitive to lipid organization and that the time-resolved quenching of pyrene fluorescence conforms well to Owen’s model for two-dimensional diffusion. The apparent dependence of the diffusion coefficient on surface pressure suggests some repositioning of CB+ in the interfacial region at higher surface pressures. Flash photolysis measurements in vesicles demonstrate that electron transfer from the excited singlet state is more efficient than from the triplet. Further, the determination of back reaction shows both a short- and long-lived component for the reduced acceptor (CB) while the donor product cation (OPyPC+) disappears more quickly. This behavior indicates that the host lipid can act as a sacrificial donor for OPyPC+.
Introduction Photoinduced electron transfer is at the center of processes which generate energy rich intermediates as precursors to the storage of energy both in biological and man-made systems.I.* Both to enhance the initial reaction and to retard back reaction, the reaction components may be incorporated into organized media where the geometry of reaction and the pathways of reactants and products can be influenced if not completely ~ o n t r o l l e d . ~In- ~systems involving lipid organizates (such as vesicles and monolayers), the kinetics of electron transfer will be influenced not only by the geometry of the system but also by the character of the microenvironment (polarity, viscosity, pH, lipid headgroup) in which such components are located. The design of systems which may provide optimal control over reaction rates and product yields will depend on knowing how microenvironmental parameters of heterogeneous media affect the kinetics of reaction. In order to elucidate the influence which such parameters can exert on kinetic behavior, one may benefit by utilization of a model environment in which these parameters may be readily altered under control. The spread lipid monolayer at the airwater interface provides such a model, allowing one to vary widely the lipid used and also to vary the organization of the lipids via compression. While this approach is applicable to investigating processes which may be monitored by luminescence-both steady state and time resolved-it is difficult to measure processes involving nonluminescent intermediates. Such is normally true for back electron transfer in which neither the redox species nor their products fluoresce. In such cases it is necessary to monitor kinetics in complimentary systems (e.g. vesicles). In this study we have examined the spectral and photoinduced electron transfer behavior of a lipid functionalized pyrene donor, 1-(10-(6(8)-octadecylpyrenyl)decanoyl)-2-hexanoylphosphoati@
Abstract published in Advance ACS Abstracts, July 1, 1995.
dylcholine (OPyPC), and an amphiphilic acceptor, 4-(N-hexadecylpyridinium-4-y1)pyridinechloride (CB+). The effects of molecular organization on these behaviors have been monitored in the presence of both lipid monolayers and vesicles. Phospholipid compounds with differing headgroups-zwitterionic dioleoylphosphatidylcholine (DOL) and negatively charged dioleoylphosphatidylglycerol (D0PG)-have been used as the host lipids. The molecular structures of the compounds involved are given in Figure 1. It has been found that the kinetics of forward transfer in these systems may be modeled to a mechanism of in-plane or two-dimensional diffusion control and that the speed of initial transfer may be altered by layer compression. The behavior of the resulting redox pair, determined by flash photolysis in vesicles of the same lipids, shows a competition between back reaction and probable scavenging of the pyrene cation by the lipid itself.
Experimental Section The probe OPyPC was purchased from KSV Chemicals. The lipids DOL and DOPG were obtained from Avanti Biochemicals. Both were used without further purification. The acceptor CB+ was synthesized according to the method of Nishijima et aL5 The structure of the compound was confirmed by NMR and elemental analysis. Materials for deposition at the gaswater interface were dissolved in methanolhenzene (5/95 vh), which was used as the spreading solvent. The apparatus for monolayer optical absorption measurement in these studies has been presented in detail elsewhere.6 The emission spectrum from the water surface was obtained by excitation with a Liconix 4230NB Cd-He Laser at 325 nm and monitored by a EG&G OMA Vision CCD equipped with a fiber optic system. The time-resolved single photon counting equipment used in these experiments has also previously been desribed.’ Here the excitation source employed was a modelocked, Q-switched Quantronix Nd:YAG system which provides a 100 ps pulse with a frequency of 4.7 kHz at the third harmonic,
0022-365419512099-16149$09.00/0 0 1995 American Chemical Society
16150 J. Phys. Chem., Vol. 99, No. 43, 1995
Li and Patterson
DOL
501
OPyPC 1 - ( l o - @ (6)-odadecylpyrenyldecanoyl)-2-hexanoylphosphatidylchoilne
I
DPG
= - $ %I? Dioleoyl phosphatidylglycerol
DOPG
e&-%-
0
u
Dioieoylphosphatidylcholine
DOL
2
4
6
8
i o so
60 70 80 Arealmolecule (Az)
BO
100
110
izo
Figure 2. Force-area isotherms for monolayers of (a) CB+; (b) OPyPC; (c) DOL; and (d) DOPG. 40
CB' 4- (N- hexadecylpyridininium-4-yl)
pyridine chloride
Figure 1. Structures of the probe, acceptor, and host lipid. 354 nm. One may obtain integrated power at this frequency up to 20 mW. All monolayer measurements were carried out on a pure water subphase (Milli-Q, Millipore) under a nitrogen atmosphere at room temperature (25 f 1 "C). Monolayer compression was carried out at 2 A2/(molecule min). For vesicle preparation, lipids with a OPyPChpid mole ratio of 1:75 were dissolved in chloroform/methanol (95/5 v/v). Solvent was removed by a stream of N2 and by subsequent evaporation in a vacuum oven for 2 h at 50 "C. Vesicles were prepared according to Ollmann et al. except that 10 mh4 potassium phosphate buffer (pH 7.4) was used to make up a final lipid concentration of 2.0 mg/m.* The samples were sonicated for 25 min (Ultrasonic Processor (50-W Model), microtip, 30 W) under N2 at 25 "C and centrifuged before use. Laser flash photolysis studies were carried out under an atmosphere of N2 with an apparatus previously de~cribed.~ The excitation source for these studies was a PRA N2 laser providing an excitation wavelength of 337 nm. The output energy per pulse was maintained at about 0.35 mJ to prevent photoionization of pyrene. Measurement of light absorbed by the pyrene chromophore in flash photolysis was determined according to literature methods using as a reference benzophenone carboxylic acid solutions having an optical density equivalent to that of the sample at 337 nm.10-12
Results Force-Area Data. Surface pressure-area isotherms on pure water surfaces for the probe OPyPC and acceptor CB+ were measured to determine relative molecular areas. The results for CB+, OPyPC, DOL, and DOPG are given in Figure 2. The surface pressure-area profiles for the phospholipids correspond to those found in the 1iterat~re.I~ The force-area isotherm of OPyPC yields a limiting area of 115 A2 molecule-I, consistent with values expected from the literature for similar pyrenebearing lipids.I4.l5 However, the isotherm of the acceptor CB+ displays an apparent limiting area per molecule of -4 A2 molecule-'. As the minimum area for a simple fatty acid is on the order of 25 A2 molecule-' and a C22 methylviologen analog of this compound exhibits an area per molecule of -40 %L2 molecule-', such behavior indicates some partitioning between the subphase and the gas-water interface for monolayers of the pure compound. In micelle studies, a c14 viologen surfactant
30 A
E
t E e
*O
Y
10
0 0
50
100
150
200
250
Area I molecule ( A* ) Figure 3. Force-area isotherms for monolayers of pure DOL, pure DOPG, and 1: 1 mixtures of these lipids with CB+. The plots are given in terms of aredipid molecule.
has been shown to be somewhat water ~ o l u b l e . ' ~By - ' ~contrast, a C22 methylviologen-mentioned above-has been shown by Lee and Bard to give a good monolayer.'* It is to be expected, then, that the interaction of CB+ with the lipid layer will be quite responsive to changes in organization of both the layer and headgroup of the molecule. Figure 3 gives the force-area isotherms for 1:l mixtures of DOL/CB+ and DOPG/CB+ in terms of area per phospholipid molecule. In both cases the addition of CB+ causes expansion of the layer over all surface pressures, indicating association of the acceptor with the layer. For the DOL/CB+ system, the area for the mixed layer approaches that of an ideal mixture, using the C22 methylviologen as a model. By contrast, the addition of CB+ brings about a very marked expansion in the area of the mixed DOPG/ CB+ layer. While one might expect more penetration of acceptor into the layer, due to headgroup charge, one might also expect some offsetting headgroup attraction. The marked effect might be thought to be related to perturbations of interactions involving the glycerol portion of the DOPG headgroups. However, a comparable expansion in the layer was obtained when replacing DOPG in the mixed system by the related negatively charged lipid, dioleoyl-sn-glycero-3-phosphate, which has no glycerol moieties in the headgroup. This suggests that the strong interaction of CB+ with DOPG draws the CB+ rings into a configuration coplanar with the water surface. Absorption and Emission Spectra. The absorption spectrum of a OPyPC monolayer is given in Figure 4 along with
J. Pkys. Ckem., Vol. 99, No. 43, I995 16151
Electron Transfer in Vesicles and Langmuir Films h
IOoo
I\. EI
i
D 5 . 2 1~ 0 . 7 c m ~
X
1
I
I
I
I
1000
I
c
j 3
(nm)
Figure 4. Absorption spectra of OPyPC in benzene (- - -) and in the monolayer (-). Monolayer absorbance multiplied by a factor of 20.
t '
D 4.10 x 10-7cm%-l
0
100
i 200
300
Time (ns) Figure 6. Plots of the Owen relationship for determination of diffusion coefficients from time-resolved fluorescence of OPyPC in mixed OPyPC/CB+/DOLmonolayers (1:6:100) at 30 mN/m and in mixed OPyPC/CB+/DOL vesicles (1:6:100). ,Ier= 354 nm and ,I,, = 400 nm. The solid lines are theoretical curves calculated via eq 1.
350
400
450
500
550
L
I
Q) Q)
OPyPC / DOPG
15000
0
v)
2
5
10000
ii
.-5 2 Q)
5000
first peak which might be expected to change with altered polarity may be seen to be larger for DOPG.4 Such behavior would generally indicate for DOPG layers a somewhat larger polarity in the environment experienced by the pyrene chromophore of OPyPC. This is consistent with some repulsion among the charged headgroups, allowing more opportunity for the chromophore to encounter the aqueous interfacial region. Time-Resolved Quenching Measurements. To determine the dynamics of electron transfer from the pyrene singlet state to CB+, fluorescence decay measurements have been made in OPyPC/DOL and OPyPCDOPG monolayers (1:lOO) in the absence and presence of CB+ at different mole fractions. The observed wavelength of emission was 400 nm. For the ideal diffusion-controlcase, the equation for the time dependent decay of fluorescence intensity for two-dimensional quenching has been derived by Owen and modified by Medhage and Almgren.'9$20It may be given as
n(t) = n(0) exp{-tlz,
0 350
400
450
500
- 7.44[Q]R(tD)"*- 2.28[Q]Dr} (1)
550
Wavelength (nm) Figure 5. Emission spectra of mixtures of DOL or D O E with OPyPC at molar ratio of 1:90 in monolayers. Arrows point in the direction of increasing intensity with increased surface pressure (n),where n = 6, 10, 20, and 30 mN/m.
the spectrum in benzene solution. The spectra are similar except that the red edge of the monolayer spectrum extends -2 nm probably due to solvent effects. Since the acceptor CB+ only absorbs light well below 340 nm, the laser beam at 354 nm will excite only pyrene chromophore in the quenching process. There is no spectral evidence for ground state aggregation from the absorbance data. The emission spectra of OPyPC in both DOL and DOPG monolayers are given in Figure 5 and demonstrate that the chromophore emits primarily from the monomer state, with very little contribution from the excimer. In the quenching studies below, we monitor the quenching of the singlet pyrene monomer. The relative enhancement of the
where TO is the fluorescence lifetime in the absence of the quencher, [Q] the quencher concentration in molecules per unit area, R the radius of interaction between fluorophore and quencher which results in efficient quenching, and D the lateral diffusion ~ o e f f i c i e n t . ~ 'In - ~recent ~ studies, a value of 1.0 nm has been used for R. Time-resolved fluorescence decay curves collected by single photon counting from DOL monolayers have been fitted to eq 1 to determine both D and n(0) via least-square fitting methods (Figure 6, upper curve). The values are given in Table 1 for DOL systems. The results for both DOL and DOPG monolayers as functions of surface pressure are given in Figure 7. We may see that the lateral diffusion coefficient decreases significantly with increasing surface pressure. To compare the diffusion characteristics exhibited in monolayers to those found in vesicles of the same lipids, fluorescence decay measurements have also been made in DOL vesicles in which the ratios of OPyPC/CB+/lipid have been maintained constant. These decays have also been analyzed using eq 1 (Figure 6,
16152 J. Phys. Chem., Vol. 99, No. 43, 1995
Li and Patterson
TABLE 1: Diffusion Coefficients for Components of Electron Transfer from OmPC to CB in DOL surface pressure (mN/m)
surface conc x (molecule/cm2)
diffusion coefficient x 10' (cm2 S-I)
3.0 5.0 6.2 3.0 5.0 6.2 3 .O 5.0 6.2 3.0 5.0 6.2
3.20 4.90 6.5 1 3.59 5.49 7.29 4.06 6.22 8.27 4.5 1 6.9 1 9.18
9.8 10.1 10.5 6.5
CB (vesicles)
3.0 6.0
4.1 4.1
multiple CB per OPyPC molecule
6 6 6 12 12 12 20 20 20 30 30 30
30
3
8.7
400
4.9
1o.oy
2ol 10
0
7.6 7.9 6.0 4.2 6.3 5.2
1 500
600
700
Wavelength, nm
h
a
20
v
a
8
10
0 r A
in
Wavelength, nm
i
Figure 8. Spectra from laser flash photolysis of mixed OPyPC/CB+/ lipid vesicles (1:6:100): (top) lipid is DOL; (bottom) lipid is DOPG. Curves: 0, just after laser flash; 0, 90 p s after the flash.
a1.o
5 X
0
0.0010
l l I5 0.1
10
15 20 25 Surface Pressure (mNlm)
30
35
O.WO8
Figure 7. Plots of diffusion coefficient as a function of surface pressure for various concentrations of CB+ in mixed OPyPC/CB+flipid monolayers (l:CB+/100) where, for DOPG systems: 0, CB+ = 3; 0 , CB+ = 7.5. For DOL systems: ,. CB+ = 3; CB+ = 5 ; 0, CB+ = 6.2.
+,
lower curve). Values of the diffusion coefficients obtained have been included in Table 1 and may be seen to be quite consistent with those found in the monolayer systems. Flash Photolysis Measurements. It is impractical, with the techniques at hand, to study the kinetics of donor cation and acceptor anion which follow their formation by photoinduced electron transfer in spread monolayers. In order to determine the behavior of these species, we have measured their transient behavior by flash photolysis in lipid vesicles with the OPyPC/ CB+/lipid ratios utilized in the monolayer studies. The spectra obtained just after completion of the forward electron transfer in vesicles of OPyPC/CB+/lipid (1:6:100) are given in Figure 8. One may see peaks at about 420, 475, and 600 nm, corresponding to the OPyPC triplet, the cation OPyPC+, and the reduced form of CB+ (CB). A second spectrum taken 90 ,us after the flash is also given in the figure. Here one may see that the triplet and most of the cation have disappeared with the triplet half-life being 25-30 ,us (not shown). However, about half the CB persists, indicating that the back reaction was much less than complete. This lack of complete removal of CB is due to competing reactions of OPyPC+, probably some reaction with the lipid. The behavior of these competing reactions is reported elsewhere.24 The transient decay for CB in DOL vesicles is given in Figure 9. Because of the competing side reaction of OPyPC+ and the triplet contribution, a best fit value for the second-order back reaction rate constant was determined numerically using the calculated concentrations of CB and total OPyPC+ for each of the 100 points of the experimental decay data. The scatter in the data is due to the very low iaser p o i V m e 3 here to avoid photoionization. The
0
25
50
75
100
125
t(C(s)
Figure 9. Decay of CB transient in Figure 8 (top) at 590 nm. The solid curve is a numerical determination of k(back reaction) from the 100 values of [CB] and [OPyPC+] which comprise the decay curve.
extinction coefficients used in this calculation were ECB = 0.8 x lo4 M-' cm-' 25 and ~ o p ~ p = c +3.8 x lo4 M-' cm-'.26 The rate constant values obtained in bulk were k(DOL)~p~pc-+c~ = 6.6 x 10" M-I s-' and k(DOPG)0py~++c~ = 5.2 x 10'' M-' s-1.
Discussion Force area measurements reported here give molecular areas for OPyPC consistent with those previously reported for lipidfunctionalized pyrene probes and somewhat smaller than those for the dioleoyl matrix lipids u ~ e d . ' The ~ , ~fluorescence ~ spectra in Figure 5 indicate that-with the absence of excimer formation-OPyPC is homogeneously distributed in the lipid layer over a wide range of surface pressures and provides a good system for study of monomeric chromophore behavior. This is further substantiated in the absorption spectra of Figure 4,where pure OPyPC monolayers show only a slight shift in profile compared to benzene solution. The differences in the surface density of the two host lipids seen in the isotherms at
J. Phys. Chem., Vol. 99, No. 43, 1995 16153
Electron Transfer in Vesicles and Langmuir Films comparable surface pressures would also be expected to affect both the dynamics of the pyrene chromophore and its interaction with the water surface. Again, the differences in relative peak heights found in the emission spectra suggest that the average environment experienced by the chromophore is more polar in DOPG monolayers than in DOL. The very low apparent molecular area observed from isotherms for pure layers of the acceptor CB+ may best be interpreted in terms of limited hydr~phobicity;'~ hence, its interaction with phospholipid layers would be expected to be influenced by headgroup charge. The measurements presented for mixed systems in Figure 3 verify that CB+ does associate with both lipids and, indeed, brings about a rather marked expansion of the layer in CBDOPG mixtures. This would be best explained by a model in which the dipyridinium group laie coplanar to the water surface; and this geometry as well as expansion of the layer would be expected to effect the subsequent photoinduced charge transfer. The singlet excited state of pyrene undergoes efficient quenching by CB+, as demonstrated by the time-resolved decay shown above. Thermodynamically, this is as expected from the redox potentials of the components involved: { E I I ~ ( P ~ + * / Py*)= - 1.81 V NHE (calculated from literature values2'); E1/2(CB+'/CB) -0.90 V25,28}.From the data in Figure 6, it may be seen also that the fluorescence quenching data can be fit well by Owen's equation for two-dimensional diffusion. The values of D found in DOPG systems are higher than those observed for DOL, which probably reflects the differences in lipid density and environment of the donor as well as acceptor described above. The diffusion coefficients may be seen to decrease approximately logarithmically with surface pressure, although the decrease is rather more pronounced for DOPG than for DOL. This can reflect changes in contributions to the organization of the layer from those factors enumerated above: (a) lipid density; (b) effects of surface charge on location as well as orientation of the acceptor; (c) interactions of the donor with the interface. Because of lipid and acceptor headgroup charge, it is difficult to make literature comparisons for reaction dynamics in the DOPG systems studied here. However, there are precedents for DOL based on determinations of diffusion coefficients. The values here are very close to those reported in measurements of D by pyrene excimer formation using 1-(6pyrenylhexanoyl)-2-palmitoylphosphatidylcholine as a probe.29 They are slightly higher than those found with DOL in timeresolved studies by Caruso et al. using a pyrenylsulfonyl moiety attached to the headgroup as donor;2' in that study D values of 1-8 x lov7cm2 s-' were reported over the same composition range. Finally, one may suggest from the reasonable agreement of D in these studies-given the variety of probes used and including processes that do not depend on electron transferthat the forward electron transfer in these systems is essentially diffusion controlled. Table 1 shows that the diffusion coefficients D measured in DOL monolayers at higher pressures are comparable to that observed in DOL vesicles. This suggests that information taken from vesicles can provide insight into the behavior of monolayers under certain conditions and vice versa. Comparison with other vesicle systems may be noted from steady state quenching of pyrene probes; Kano et al. reported an effective diffusion coefficient -3-5 x cm2 s-l in bilayers of dipalmitoylphosphatidylcholine (DPPC) above the phase transition temperature (Tc).30 It may be seen from monolayer studies that the aredmolecule values for DPPC above TC3land DOL at higher surface pressures are comparable. Barenholz et al. report values of D between 4 x lo-' and 9 x lo-' cm2 s-' for
-
functionalized pyrene quenching in soybean PC.32 These literature values of D are quite close to those determined in this study. The flash photolysis data from vesicle measurements provide some insight into the effects of the lipid environment on the back reaction of this particular system. In Figure 8, it may be seen that there is complete, rather rapid decay of the pyrene cation and some CB, followed by a persistent level of CB. This suggests a pathway for cation degradation altemate to back electron transfer. The double bonds in DOL or the headgroup could provide a sacrificial source of electrons which can compete with back reaction from CB. Additionally, the change in charge on CB may cause it to move into a position and geometry other than that occupied by CB+. The second-order rate constants obtained in bulk and given above were k(DOL)opy~++c~ = 6.6 x 10" M-' s-' and k(DOPG)opypc++c~= 5.2 x 10" M-' S-I. To compare these values with the forward reaction rates, these values may be recalculated in terms of a two-dimensional system by estimating the lipid area in which the back reaction takes place. Because of the agreement between forward reactions in vesicles and monolayers at higher surface pressures, values for the aredipid molecule used in this calculation were taken from the force-area isotherms at 30 mN/m. Following this, one may use the approximation of Galla and Sackman to arrive at a diffusion ~ o e f f i c i e n t .The ~ ~ estimated values in two dimensions become D(DOL)OP~PC++CB = 4 x lo-' cm-2 s-' and D(DOPG)0pypc++c~ =3 x cm-2 s-l. These values are comparable to the rates obtained for forward reaction and appear to be-within the assumptions of the calculation-diffusion controlled.
Conclusion This system-with a lipid-attached donor in the monolayer and a functionalized acceptorprovides a useful method by which to study, in a controlled way, those interfacial parameters which influence the dynamics of photoinduced electron transfer in organized systems. Interestingly, when these results are compared to those from the literature of functionalized pyrene reactions in DOL as a lipid matrix, two points are evident: (a) in terms of diffusion coefficient under the same conditions of molecular organization, the dynamics for electron transfer as well as excimer formation fall into a fairly narrow range for differing pyrene probes and electron acceptors; (b) diffusion coefficients found in monolayers are comparable to those in vesicles under the same phase conditions. The latter observation can, with care, make useful the combination of studies from monolayers and vesicles. However, when different lipids are used, the relative dependence of D on monolayer compression can vary from lipid to lipid (e.g. DOL and DOPG), reflecting the complex interactions of lipid moiety, headgroup, and water surface which comprise the reaction environment. Finally, while a system such as this provides an excellent means for studying the effects on electron transfer of changing the organization and character of the interfacial environment, it is not suggested that this alone will provide a good system for the storage of light energy from energy rich charged intermediates. To that end it is useful to have a means for stabilizing charge such as, for example, introduction of a secondary acceptor which can utilize the character of the headgroup to maintain charge separation. Such studies are in progress.24
Acknowledgment. The research described herein was supported by the Office of Basic Energy Sciences of the U.S.
16154 J. Phys. Chem., Vol. 99, No. 43, 1995
Department of Energy. This is Document No. 3811 from the Notre Dame Radiation Laboratory.
References and Notes (1) Photoinduced Electron Transfer, PART D ; Fox, M. A,, Chanon, M., Ed.; Elsevier: Amsterdam, 1988; pp 241-441. (2) Kavamos, G. J. Fundamentals of Photoinduced Electron Transfer; VCH: New York, 1993; pp 185-286. (3) Kalyanasundaram, K. Photochemistry in Microheterogeneous Systems; Academic Press: New York, 1988. (4) Photochemistry in Organized and Constrained Media, 1st ed.; Ramamurthy, V., Ed.; VCH Publishers, Inc.: New York, 1991; pp 39-77 and references therein. (5) Nishijima, T.; Nagamura, T.; Matsuo, T. J . Polym. Sci., Polym. Lett. Ed. 1981, 19, 65. (6) Wang, Y. C.; Wang, Y. M.; Patterson, L. K. Rev. Sci. Insrrum. 1994, 65, 2781. (7) Subramanian, R.; Patterson, L. K. J . Phys. Chem. 1985,89, 1202. (8) Ollman, M.; Schwarzmann, G.;Sandhoff, K.; Galla, H. J. Biochemistry 1987, 26, 5943. (9) Nagarajan, V.; Fessenden, R. W. J. Phys. Chem. 1985, 89, 2330. (10) Lutz, H.; Breheret, E.; Lindqvist, L. J. Phys. Chem. 1973, 77, 1758. (11) Inbar, S.; Linschitz, H.; Cohen, S. G. J . Am. Chem. Soc. 1981, 103, 7323. (12) Carmichael, I.; Hug, G. L. J. Phys. Chem. Ref. Data 1986, 15, 155. (13) Mingotaud, A. F.; Mingotaud, C.; Patterson, L. K. Handbook of Monolayers; Academic Press: New York, 1993; pp 484. (14) Kinnunen, P. K.; Virtanen, J. A,; Tulkki, A. P.; Ahuja, R. C.; Mobius, D. Thin Solid Films 1985, 132, 193. (15) Somerharju, P. J.; Virtanen, J. A,; Eklund, K. K.; Vainio, P.; Kinnunen, P. K. Biochemistry 1985, 24, 2773.
Li and Patterson (16) El Torki, F. M.; Schmehl, R. H.; Reed, W. F. J. Chem. SOC., Faraday Trans. 1 1989, 85, 349. (17) Brugger, P.; Gratzel, M. J. J. Am. Chem. Soc. 1980, 102, 2461. (18) Lee, C.-W.; Bard, A. J. Chem. Phys. Lett. 1990, 170, 57. (19) Owen, C. S. J . Chem. Phys. 1975, 62, 3204. (20) Medhage, B.; Almgren, M. J. Fluoresc. 1992, 2, 7-21. (21) Caruso, F.; Grieser, F.; Murphy, A.; Thistlethwaite, P.; Urquhart, R.; Almgren, M.; Wistus, E. J. Am. Chem. SOC. 1991, 113, 4838-43. (22) Caruso, F.; Grieser, F.; Thistlethwaite, P. J.; Almgren, M. Langmuir 1993, 9, 3142-8. (23) Caruso, F.; Grieser, F.; Thistlethwaite, P. J.; Almgren, M. Biophys. J. 1993, 65, 2493-503. (24) Li, L.; Patterson, L. K. Photochem. Photobiol., in press. (25) Hammarstrom, L.; Almgren, M.; Lind, J.; MenCnyi, G.; Norrby, T. J. Phys. Chem. 1993, 97, 10083. (26) Slama-Schwok, A,; Ottolenghi, M.: Avnir, D. Nature 1992, 355, 240. (27) Murov, S. L.; Carmichael, I.: Hug, G.L. Handbook of Photochemistry; Marcel Dekker, Inc.: New York, 1993. (28) Roullier, L.; Laviron, E. Electrochim. Acta 1978, 23, 773. (29) Bohorquez, M.; Patterson, L. K. J. Phys. Chem. 1988, 92, 1835. (30) Kano, K.; Kawazumi, H.; Ogawa, T. J . Phys. Chem. 1981,85,2204. (31) Hunt, R. D.; Mitchell, M. L.; Dluhy, R. A. J . Mol. Struct. 1989, 214, 93. (32) Barenholz, Y.; Cohen, T.; Korenstein, R.; Ottolenghi, M. Biophys. J. 1991, 59 (l), 110. (33) Galla, H.-J.; Sackmann, E. Ber. Bunsen-Ges. Phys. Chem. 1974, 78, 949. JP950701W
