Systematic Quantification of Electron Transfer in a Bare Phospholipid Membrane Using Nitroxide-Labeled Stearic Acids: Distance Dependence, Kinetics, and Activation Parameters

In this report, we present a method to characterize the kinetics of electron transfer across the bilayer of a unilamellar liposome composed of 1,2-dimyristoyl-sn-glycero-3-phosphocholine. The method utilizes synthetic phospholipids containing noninvasive nitroxide spin labels having the >N–O• moiety at well-defined distances from the outer surface of the liposome to serve as reporters for their local environment and, at the same time, permit measurement of the kinetics of electron transfer. We used 5-doxyl and 16-doxyl stearic acids. The paramagnetic >N–O• moiety is photo-oxidized to the corresponding diamagnetic oxoammonium cation by a ruthenium electron acceptor formed in the solution. Electron transfer is monitored by three independent spectroscopic methods: by both steady-state and time-resolved electron paramagnetic resonance and by optical spectroscopy. These techniques allowed us to differentiate between the electron transfer rates of nitroxides located in the outer leaflet of the phospholipid bilayer and of those located in the inner leaflet. Measurement of electron transfer rates as a function of temperature revealed a low-activation barrier (ΔG‡ ∼ 40 kJ/mol) that supports a tunneling mechanism.


■ INTRODUCTION
Electron transfer (ET) is one of the principal processes involved in harvesting and transferring energy in natural and artificial systems. Indeed, it is an essential process in all living organisms. 1−3 Extensive studies of ET have been performed on proteins, 1,4,5 DNA, 6,7 dendrimers, 8 and artificial photosynthetic centers. 9,10 A consistent effort is devoted to the ET across lipid membranes. 11 Most of studies on ET across biological membranes focus on natural or artificial redox-active complexes embedded in a lipid bilayer and on the effects of the presence of cations, on the processes of electron and proton transfer. 12,13 The study of membranes is relevant to solar energy conversion. 12,14−19 Also, knowledge of ET across membranes is fundamental both for understanding the functioning of natural photosynthetic systems and for creating their artificial analogs. 12,20 The contribution of the fatty-acid chains of the phospholipids to ET is poorly understood. The roles of the main types of chemical bonds, that is, hydrogen-, π-, and σ-bonds in ET are not well understood either. 13,21,22 In studies on membrane systems, unilamellar liposomes are often used as model systems, 23−27 with the donor and acceptor being separated by a lipid bilayer. In general, methods that rely on optical spectroscopies utilize bulky and rigid dyes, inferring perturbations to the system; it is a challenge to identify probes that do not substantially alter the membrane, and, at the same time, provide clear-cut information on the ET phenomena. 12,14,17,19,23−26,28−30 Another challenge for the methods is the flexibility to cover kinetics extending over several orders of magnitude.
Electron paramagnetic resonance (EPR) spectroscopy represents an interesting option for studying reaction kinetics in a wide range of time scales, from microseconds 31 to hours. 32 It requires a paramagnetic center in the system, but, with respect to popular optical spectroscopies, it overcomes the disadvantage of coping with unresolved and overlapping optical absorptions or background signals.
Here, we present a novel and highly efficient methodology for studying ET across membranes based on EPR. We make use of nitroxides as paramagnetic centers that are attached to stearic acids in different positions to localize them at different depths in the membrane. These molecules have been selected as they do not substantially perturb the phospholipid bilayer. 33−36 Photoinduced oxidation of the nitroxides, using a well-established procedure, yields time-dependent EPR signal intensities that can be straightforwardly translated into kinetic data. It is a key advantage that the study of the EPR spectra (eventually in the presence of added relaxants) can provide relevant information both on the positioning of the nitroxide moiety within the bilayer; and on the environment of the probe.
Use of these probes enables us to systematically address the contribution of the phospholipids to ET across a biological membrane. It is important to distinguish this "background" contribution if one wants to focus on ET of active constituents, such as proteins and metal complexes dissolved into the membrane.
Two probes, 5-doxyl stearic acid (5DSA) and 16-doxyl stearic acid (16DSA) (Scheme 1), were introduced into unilamellar liposomes composed of 1,2 dimyristoyl-sn-glycero-3-phosphocholine (DMPC). 37,38 While in 5DSA the nitroxide moiety is located adjacent to the hydrophilic moiety of DMPC, in 16DSA it resides deep within the lipophilic domain. 39 Molecular dynamics simulations predicted 5DSA to be located only ∼ 0.5 nm from the membrane surface. For 16DSA the distance increases to ∼ 1.8 nm. 39 These distinctly differing locations of the probes are, therefore, well suited to obtaining insight into the distance dependence of the ET process.
We selected DMPC-based membranes since they display a transition from a gel-like to a liquid crystalline phase at room temperature (296.6 K). 40 This allows us to investigate the influence of the rigidity of the lipid bilayer on ET. In the liquid crystalline state, lateral diffusion of lipids within the vesicle bilayer is rapid, whereas the transverse "flip-flop" motion is extremely slow. 41 Below 296.6 K, the transition to the gel-like phase causes all modes of mobility to decrease by orders of magnitude. 42,43 To monitor the ET kinetics, we needed a reaction sequence that could be triggered. Here, we utilized the well-established photoinduced oxidation of [Ru(bpy) 3 ] 2+ to [Ru(bpy) 3 ] 3+ in the presence of ammonium persulfate ((NH 4 ) 2 S 2 O 8 ). 44−50 The decisive and rate-limiting step is the subsequent oxidation of the nitroxide moiety by ground-state [Ru(bpy) 3 ] 3+ . The reaction produces the EPR-silent oxoammonium cation (OAC), with concomitant regeneration of [Ru(bpy) 3 ] 2+ (Scheme 2). This approach has been used successfully for monitoring ET reactions, 31,44,51 since the photoinduced formation of [Ru(bpy) 3 ] 3+ is faster than its reaction with RNO • ; furthermore, the Ru-complex is highly water-soluble, therefore it does not diffuse into the membrane.
The complex [Ru(bpy) 3 ] 2+ displays strong absorption at 452 nm. Following the decrease in absorption at this wavelength thus provides an additional tool for following the ET kinetics. 46,52,53 To obtain a strong set of experimental data, we followed the kinetics of the reactions displayed in Scheme 2 by steady-state EPR (continuous irradiation), by laser-triggered time-resolved EPR, and by optical absorption spectroscopy.

■ EXPERIMENTAL SECTION
Liposome Preparation. Liposomes were prepared by ultrasonication. 54 Briefly, the phospholipid and the corresponding stable nitroxide radical (SNR) (w/w 50:1) were dissolved in 0.5 mL dichloromethane and dried to a thin film under a nitrogen stream in a test tube at room temperature. After further drying for 1 h under vacuum, the film was hydrated with 1 mL of buffer at neutral pH and then sonicated for 15 min. Residual free SNRs were removed by dialysis, and their complete removal checked by EPR.
Liposome Characterization by Small-Angle X-ray Scattering (SAXS). The measurements utilized a SAXSpoint 2.0 (Anton Paar, Graz, Austria) apparatus containing a Primux 100 micro microfocus X-ray source operating at λ = 0.154 nm (Cu-K α ). Two-dimensional scattering patterns were recorded by a 2D EIGER R Series Hybrid Photon Counting (HPC) detector (Dectris Ltd., Baden-Daettwil, Switzerland). The samples were inserted into a capillary (1 mm diameter) and measured nine times for 300 s. The scattering patterns were averaged and compensated for the cosmic X-ray impacts. All measurements were performed at 20°C. Absolute scale calibration was achieved by using water as a secondary standard. 55 All SAXS data were evaluated by a generalized indirect Fourier transform (GIFT) method to determine the pair distance distribution functions. 56−58 Liposome Characterization by Dynamic Light Scattering. The dynamic light scattering (DLS) equipment consisted of a diode laser (Coherent Verdi V5, λ = 532 nm) and a goniometer with singlemode fiber detection optics (OZ from GMP, Zurich, Switzerland). The data were acquired by an ALV/SO-SIPD/DUAL photomultiplier with pseudocross correlation and an ALV 7004 Digital Multiple Tau Real Time Correlator (ALV, Langen, Germany). The ALV software package was used to record and store the correlation functions. Light scattering was measured five times for 30 s, at a scattering angle of 90°a nd a temperature of 25°C, and the resulting correlation functions were averaged. The hydrodynamic radius was calculated using optimized regulation technique software. 59 Continuous-Wave Electron Paramagnetic Resonance. Continuous wave (cw) EPR spectra were recorded on a Bruker X-band spectrometer (EMX, 100 kHz field modulation) at 275, 283, 303, and 310 K with a 0.15 mT field modulation amplitude. Photolysis was conducted using a Hamamatsu Lightingcure LC4 Xe/Hg lamp. The correlation times were calculated using the MATLAB-based GUI SimLabel. 60 The concentration of the SNR was 2 × 10 −5 M in all measurements. The concentration of [Ru(bpy) 3 ]Cl 2 and (NH 4 ) 2 S 2 O 8 were 1 × 10 −5 and 5 × 10 −4 M, respectively.
Power Saturation Experiments. The power saturation experiments for determining the position of the nitroxide moieties within the bilayer of the DMPC liposomes were performed on an E580 ELEXSYS Bruker X-band spectrometer, equipped with a roomtemperature dielectric resonator, ER4123D. All spectra were obtained using the following parameters: modulation amplitude 0. 15  The experimental protocol for the insertion depth measurements was as follows: approximately 5 μL samples were loaded into a gas permeable TPX capillary (L&M EPR Supplies, Inc., Milwaukee, WI, USA), and three saturation experiments were performed. The first experiment was done on the sample in equilibrium with air, to saturate the membrane with oxygen. The second experiment was performed on the same sample after deoxygenation under dry nitrogen flow for 20 min. The third experiment was carried out on a new sample to which a nickel complex with ethylenediaminediacetic acid (NiEDDA) had been added to a final concentration of 20 mM; the sample was then deoxygenated as mentioned above. The three power saturation curves (intensity vs microwave power) were obtained using a home-written program in Matlab that extracts the peak-to-peak amplitudes of the central line of the EPR spectra of the above experiments. The saturation curves were fitted using the standard equation: 27,61,62 y A x where p 1/2 is the saturation parameter, namely, the power at which the first derivative amplitude is reduced to half of its unsaturated value, h is the homogeneity parameter, indicating the homogeneity of saturation of the resonance line (ranging from h = 1.5 for a fully homogeneous line to h = 0.5 for a fully inhomogeneous line), and A is a scaling factor that accounts for the absolute signal intensity. The dimensionless parameter (Φ) that is related to the immersion depth of the nitroxide inside the membrane can be directly calculated from the ratio of the p 1/2 parameters obtained from the fitting of the data from the three experiments described above: Time-Resolved Electron Paramagnetic Resonance. Timeresolved EPR (TR-EPR) signals were collected using the instrument lock-in amplifier connected to a fast digitizer on an E580 ELEXSYS Bruker X-band spectrometer. The modulation frequency was 100 kHz, and the modulation amplitude was 0.2 mT. The modulation of the magnetic field was set to a frequency of 100 kHz so that the time resolution was regulated by the time constant value (TC) to a maximum value of ∼ 10 μs. The experimental procedure involved a single laser pulse to photoexcite [Ru(bpy) 3 Cl 2 ], thus initiating the ET process as depicted in Scheme 2 followed by monitoring the decay of the EPR signal. Photoexcitation was performed using a Quantel Rainbow Nd:YAG laser (1064 nm) mounted with second and third harmonic modules and an optical parameteric oscillator. The final irradiation wavelength was set to 436 nm. The concentration of the SNR was 6.4 × 10 −4 M in all measurements.
Time-Resolved UV−visible Spectroscopy. TR-UV-vis spectra were recorded on a UV-vis spectrometer equipped with optical fibers and a 1024-pixel diode-array detector (J&M Analytik AG, Essingen, Germany). Standard fluorescence quartz cuvettes were used for all measurements. Excitation of the samples was performed using a Hamamatsu Lightingcure LC4 Xe/Hg lamp. The concentration of the  48 It is reduced to [Ru(bpy) 3 ] 2+ by the nitroxide groups (oxidized to OAC) in a bimolecular process acting as the bottleneck of the ET process, eqs (1) and (2), and eqs (3) and (4)  Kinetic Analysis. Kinetic analysis was conducted using the kinetic reaction shown in Scheme 3 and explained in greater detail in the Supporting Information. The rate constants from the literature data were used where possible (Supporting Information, Schemes S1 and S2). The rate constants for the other reactions were determined by fitting the experimental curves with COPASI, a free software. 63 Two second-order distance-dependent reactions were used for the limitingstep electron transfer reaction for both 5DSA and 16DSA.

■ RESULTS AND DISCUSSION
Structure of the Liposomes and Properties of the Spin Probes. It was first necessary to establish that the presence of the spin-labeled fatty acids does not affect the integrity of the membrane. DLS measurements reveal that both the unmodified liposomes and those containing either 5DSA or 16DSA have a narrow size distribution, with an average hydrodynamic radius of 53 nm (see Supporting Information, Figure S5). SAXS shows that the thicknesses of the liposomes (ca. 5.1 nm, a result corroborated by molecular dynamics simulations), 62 is insignificantly altered when either of the probes resides in the membrane (see Supporting Information, Figure S4). We conclude that insertion of the phospholipids containing the SNRs does not significantly perturb the membrane structure.
Besides being a pivotal reactant in the ET reaction, the RNO • group also serves as an efficient probe of the local environment. Figure 1 shows the temperature-dependent EPR spectra of 16DSA and 5DSA. Significantly, the signals detected for 16DSA display narrower spectral lines than those of 5DSA.
This is a direct indication that the NO • group of the 16DSA resides in an environment whose density/viscosity is substantially lower than that of the surrounding 5DSA. Above the transition temperature, in the liquid-crystal phase (>296.6 K), for both probes the lines become narrower, revealing higher mobility (much shorter rotational correlation times, see Supporting Information Table S1). These temperature-dependent lineshape changes are reversible. The data obtained are in line with 16DSA being positioned in the rather flexible, lipophilic environment toward the center of the bilayer, while 5DSA resides in a more ordered, rigid, and hydrophilic region close to the polar head group. 38,64−67 Our experimental data correspond well with the MD simulations referred to in the introduction. 39 Simulations evidence that all the EPR signals measured represent single species. This demonstrates that each label occupies a well-defined position within the membrane (on the EPR time scale) and confirms that no labels are detectable in the aqueous solution outside the membrane.
It has been reported that the fatty-acid-based chains bearing the labels may fold back within the bilayer. 36 Such a process would substantially affect the evaluation of the distance dependence. We, therefore, used power saturation EPR to experimentally address this issue. This method provides the depth parameter Φ (see the Experimental Section, and also Supporting Information Figure S8 and Table S2), which is a measure of the immersion depth of the spin label. 61 Φ values of 1.5 and 2.9 were obtained for 5DSA and 16DSA, respectively. These values are very similar to those reported for structurally related 5DPC (1-palmitoyl-2-stearoyl-(5doxyl)-sn-glycero-3-phosphocholine, Φ = 1.4) and 14DPC (1-palmitoyl-2-stearoyl-(14-doxyl)-sn-glycero-3-phosphocholine, Φ = 3.0) in comparable POPC membranes (Table S2). 68 This strengthens our contention that the nitroxide moiety in 16DSA is positioned substantially deeper within the bilayer than 5DSA, and that back-folding does not make a substantial contribution, in agreement with earlier findings. 69 Kinetics. After preparing and characterizing the unilamellar phospholipid vesicles and their nitroxide-labeled derivatives, they were added to a solution containing [Ru(bpy) 3 Cl 2 ] and (NH 4 ) 2 S 2 O 8 . As shown in Scheme 2, irradiation converts the Ru(II) complex (present in the external aqueous phase) to Ru(III), which then oxidizes the paramagnetic nitroxide moiety to the EPR-silent oxoammonium cation. Accordingly, the Ru complex in the outer aqueous phase can interact with the nitroxide residing in the adjacent outer leaflet and a more distant one in the inner leaflet. This has to be accounted for in the kinetic equations, which are represented in Scheme 3. Consequently, the oxidation of the nitroxide moiety has to be split into two components: one accounting for the shorter (k outer ) and one for the longer (k inner ) distance. In terms of the overall concentration of the spin labels, we assume that they are evenly distributed between the inner and outer leaflet (eqs 3 and 4 in Scheme 3). The overall reaction scheme is therefore the combination of Schemes 2, and 3, and it is shown in Schemes S1 and S2 (See the Supporting Information) for the two cases of symmetric and nonsymmetric liposomes. The numerical solution of the overall equation scheme as described in the Experimental Section allows the determination of the concentration of all species and the fitting of all the experimental traces.
EPR time traces obtained under steady-state irradiation of nonsymmetrically prepared liposomes are displayed in Figure  2a−d. These curves reveal two reaction regimes, one dominating at short times, the second emerging after about 10 s. This is consistent with the presence of reactions 1 and 2 in Scheme 3, since ruthenium resides only in the solution outside the liposome (see Scheme S1). Our hypothesis has been verified using symmetric liposomes (Figure 2e−h, see  below). The presence of two distinct rates for the two leaflets is possible since the nitroxides do not quickly exchange between the two leaflets in the time frame of the experiments,  Table  S1. Note that the experimental spectra do not contain components corresponding to spin labels in the aqueous phase outside the lipid bilayer.
Langmuir pubs.acs.org/Langmuir Article as the "flip-flop" motion is known to be extremely slow. Additionally, as proved by the lineshapes of the cw-EPR spectra and by the power saturation EPR experiments on symmetric liposomes, 5DSA and 16DSA occupy markedly distinct positions in the bilayer and therefore should give rise to different electron transfer rates. Following these observations, we solved the kinetic scheme and fitted the different time traces. From the solution, we determined that eqs 1 and 2 of Scheme 3, characterized by k outer and k inner, are the rate-limiting steps. For 5DSA, k outer increases from 3.6 to 6.2 × 10 5 M −1 s −1 increasing the temperature from 275 to 310 K ( Table 1). The k inner rate constant is two orders of magnitude smaller and increases from 4.7 to 15.0 × 10 3 M −1 s −1 in the same temperature range. Analogously, for 16DSA, the rate constants increase from 1.6 to 3.8 × 10 5 M −1 s −1 and from 6.8 to 33.0 × 10 3 M −1 s −1 , respectively ( Table 1). The observation that the rate constant k outer for 5DSA is larger than that of 16DSA by a factor of two (at all temperatures) let us conclude that the nitroxide moiety of 16DSA is approximately 1 nm more distant from the water/ membrane interface (containing the Ru(III) complex) than that of 5DSA (see also below). 39 Markedly, k inner is bigger for 16DSA (closer to the outer liposome surface) and smaller for 5DSA. The rates, which we assign to the oxidation of the nitroxides in the inner leaflet, k inner , are ca. two orders of magnitude lower than k outer .
To verify that the origin of two electron transfer rates have been correctly assessed, we designed an experiment in which the signal decay depends only on the electron transfer rate of the outer leaflet, the fast electron transfer process (k outer ). We prepared liposomes that contained [Ru(bpy) 3 ]Cl 2 / (NH 4 ) 2 S 2 O 8 in the external and in the internal aqueous phase (symmetric liposomes), and we employed time-resolved EPR coupled to pulsed-laser irradiation to focus on the microsecond range. Indeed, the decay curves so obtained for the EPR signals of 16DSA and 5DSA (Figure 2e−h and Supporting Information Figure S3) display only one component. The corresponding temperature-dependent rate constants are therefore obtained from solution of Scheme S2. They are identical (within the experimental error, Table 1) to those assigned to k outer determined in the steady-state experiment. This clearly underpins the assignment of the two distinctly different time regimes to the ET to the inner and outer leaflets.
In a parallel set of experiments, we used optical spectroscopy to follow the concentration of the oxidant, [Ru(bpy) 3 ] 3+ , present only in the external aqueous phase, to obtain the absolute rate constant for its formation and to establish its steady-state concentration under our experimental conditions. Upon irradiation of [Ru(bpy) 3 ]Cl 2 in the presence of (NH 4 ) 2 S 2 O 8 , the band at 452 nm of [Ru(bpy) 3 ]Cl 2 decreases   (Figure 3a). 53,70,71 Finally, to confirm the kinetics of the slow process, we took advantage of UV-vis spectroscopy and followed the reaction with a low time resolution >10 s. A cuvette containing the photoredox system was irradiated for 30 s before each measurement, to convert all the [Ru(bpy) 3 ] 2+ to its oxidized form. After injection of the liposomes containing the SNRs, the absorbance at 452 nm was monitored. Indeed, the reappearance of the 452 nm band, after immediate addition of the SNR in the dark, allows us to follow the much slower and ratedetermining reduction of [Ru(bpy) 3 ] 3+ by SNRs of the inner leaflet, while the one from the outer leaflet is too fast to be detected. Figure 3b displays the concentration of [Ru(bpy) 3 ] 3+ as a function of time. The corresponding rate constants at 295 K are 3.5 × 10 3 M −1 s −1 for 5DSA and 8.0 × 10 3 M −1 s −1 for 16DSA; they are close to the values for k inner obtained by cw-EPR.
In order to obtain more information on the kinetics, and in particular on the activation parameters, we have determined the temperature dependence of the k inner and k outer using Eyring theory (see Supporting Information Figure S6); here, the rather small but systematic changes of k inner and k outer (Table 1) afford alike and small free-activation energies, ΔG ‡ , of ca. 41 kJ/mol for both 5DSA and 16DSA (Table 2). Finally, it is necessary to discuss the ET mechanism in the light of our experimental data. It has been shown that tunneling, or superexchange of electrons between sites, 72,73 is a plausible mechanism for ET in phospholipid membranes. 19,20,30,74 Our data confirm the low energy barriers that were both predicted and experimentally validated. 1−3 For tunneling to occur, the donor and acceptor pairs need to interact via an electronic coupling matrix element. Calculation of the electronic coupling elements is possible, on the basis of Marcus theory. 2 A numerical estimation of these elements is provided under the Supporting Information. Naturally, these elements are dependent on the distance and on the medium, following an exponential function. In our case, a distance-decay parameter of 0.5 ± 0.1 nm −1 was obtained, a value somewhat similar to others reported for membranes. 74−76 We think that the main reason for the disagreement is the different definition of the distance. We considered geometric distances, whereas in the literature the measurements are conducted electrochemically, thus the technique is likely more sensitive to the apolar part than to the polar one. 75 Other mechanisms that may be considered for ET in liposomes, such as adiabatic ET, have been ruled out. In fact, adiabatic ET is often characterized by a sensitivity of the ET process to solvent properties, that is, reorganization, whether in an isotropic environment 77 or in a microheterogeneous medium. 78 In our system, the alkyl chains of DMPC can be regarded as a solvent with very slow reorganization capabilities. 78 It is thus likely that the dynamics of the alkyl chains (equivalent to solvent relaxation) would dominate the kinetics of ET and slow down the ET rates, which would be then much lower than in an aqueous solution. 31 However, our data do not display such a decrease in ET rates. Moreover, the temperature dependence does not support an adiabatic mechanism. Finally, involvement of a hopping mechanism appears to be unlikely, since no electroactive component (e.g., an aromatic group) is located between the donor and the acceptor. 79

■ SUMMARY AND CONCLUSIONS
In the present study it was demonstrated, for the first time, that it is possible to study the kinetics of ET through a lipid membrane bilayer from an electron donor (nitroxide radicals) located within the membrane to a photogenerated Ru(III) in the aqueous solution. Introduction of the spin-label at different positions on the fatty-acid chain permitted construction of a molecular "ruler" for monitoring ET in natural and artificial membrane systems by precise control of the donor/acceptor distance.
We have used this experimental system to determine both the rate constants and activation energies for ET between [Ru(bpy) 3 ] 3+ and the nitroxide moieties in 5DSA and 16DSA embedded in unilamellar DMPC liposomes. By using both EPR and optical methods, we obtained significant information: (i) We could unequivocally show that the spin labels do not affect the structure of the phospholipid membrane; (ii) we could quantify the penetration depth and the local mobility of the labels 5DSA and 16DSA.  Our results create a consistent picture for ET through the phospholipid bilayer. Low values of activation energy for ET (ΔG ‡ values of 40 ± 4 kJ/mol for 5DSA, and 42 ± 5 kJ/mol for 16DSA), as well as the absence of a pronounced effect of the membrane phase transition on the kinetics of ET, provide experimental evidence that ET within the phospholipid bilayer occurs via a tunneling mechanism. We determined a distance dependence of k ET with a β-parameter of 0.5 ± 0.1 nm −1 .
The "molecular ruler" methodology that we have developed can be applied not only to phospholipid bilayers, but also to biological membranes, to artificial membranes, and to proteins and protein complexes, thus providing direct experimental information concerning the dynamics of their local environments. It can thus be used, for example, in investigating the involvement of π-electrons and/or water bridges in long-range ET. Since water is always present in membranes (the so called "hydropathy plot" 80 ), and EPR signals of nitroxides are sensitive to the polarity of the environment, our approach provides a unique opportunity to studying the roles of water and hydrogen bonds in ET in membranes and proteins (dehydration methods are well known).
EPR spectra of some stable nitroxide radicals (imidazoline and imidazolidine types) are pH-sensitive. 44,51,81 Use of such probes seems extremely promising for simultaneous examination of electron and proton transfer in membrane systems by the EPR approach described here. Their use can provide a unique method for quantitative characterization of protoncoupled ET reactions 82,83 in many important natural and artificial catalytic systems. 84