Spatial Orientation and Electric-Field-Driven Transport of Hypericin

Article pubs.acs.org/JPCB

Spatial Orientation and Electric-Field-Driven Transport of Hypericin Inside of Bilayer Lipid Membranes Alena Strejčková,† Jana Staničová,‡ Daniel Jancura,† Pavol Miškovský,† and Gregor Bánó*,† Department of Biophysics, Pavol Jozef Šafárik University, Jesenná 5, Košice 041 54, Slovak Republic Institute of Biophysics and Biomathematics, University of Veterinary Medicine, Komenského 73, Košice 041 81, Slovak Republic

† ‡

ABSTRACT: Fluorescence experiments were carried out to investigate the interaction of hypericin (Hyp), a natural hydrophobic photosensitizer, with artificial bilayer lipid membranes. The spatial orientation of Hyp monomers incorporated in diphytanoyl phosphatidylcholine (DPhPC) membranes was determined by measuring the dependence of the Hyp fluorescence intensity on the angle of incidence of pand s-polarized excitation laser beams. Inside of the membrane, Hyp monomers are preferentially located in the layers near the membrane/water interface and are oriented with the S1 ← S0 transition dipole moments perpendicular to the membrane surface. Transport of Hyp anions between the two opposite sides of the lipid bilayer was induced by applying rectangular electric field pulses to the membrane. The characteristic time for Hyp transport through the membrane center was evaluated by the analysis of the Hyp fluorescence signal during the voltage pulses. In the zero-voltage limit, the transport time approached 70 ms and gradually decreased with higher voltage applied to the membrane. In addition, our measurements indicated an apparent pKa constant of 8 for Hyp deprotonation in the membrane.

1. INTRODUCTION Hypericin (Hyp) (Figure 1), a natural photosensitizing pigment occurring in the plants of the genus Hypericum, has been

phase, and in most polar aprotic solvents. A pKa value of 1.8 has been reported for the bay-hydroxyl group of Hyp (Figure 1) in aqueous environment.7 There are still open questions regarding Hyp-induced processes in cells. Two of the most important issues are the kinetics and molecular mechanism of Hyp uptake by target cells. Particularly, the transport of Hyp through the cell membrane and its subsequent intracellular localization are of major importance. In general, Hyp is preferentially accumulated in lipid membrane structures of cells.17−19 Depending on the cell line and experimental conditions, Hyp has been reported to localize in different subcellular compartments (endoplasmic reticulum, mitochondria, plasma membrane, nuclear membrane, Golgi apparatus).4,10,17,20−22 Despite numerous publications dealing with the physicochemical properties and intracellular localization of Hyp, many details of Hyp interaction with biological membranes are still to be examined. Comprehensive study of Hyp interaction with GUVs (giant unilamellar vesicles) and lipid monolayers has recently been published.23 Using GUVs of ternary lipid mixtures, it has been shown that Hyp has high affinity to lipid rafts. In addition, it has been proven that Hyp is prone to build up in cholesterol-rich domains of binary mixture (cholesterol and DPPC) Langmuir monolayer systems.23 Work of Losi et al.24 indicated that above

Figure 1. The chemical structure of (a) Hyp and (b) deprotonated Hyp (bay-hypericinate). The arrow pointing toward the bay region indicates the S1 ← S0 transition dipole orientation.

intensively studied because of its cytotoxic, antibacterial, and antiviral properties and its possible application in photodynamic therapy of cancer.1−6 Hyp, due to its hydrophobic character, forms nonfluorescent aggregates in aqueous solutions,7,8 which significantly suppresses its photodynamic activity.9−11 On the other hand, Hyp is readily soluble in many polar organic solvents (ethanol, dimethyl sulfoxide (DMSO), acetone)7,8 and in lipid structures. In these environments, Hyp exists in its biologically active monomer form, exhibiting strong fluorescence and a high quantum yield of singlet oxygen formation.12−16 Typically, Hyp is deprotonated in the aqueous © 2013 American Chemical Society

Received: November 20, 2012 Revised: December 21, 2012 Published: January 3, 2013 1280

dx.doi.org/10.1021/jp3114539 | J. Phys. Chem. B 2013, 117, 1280−1286

The Journal of Physical Chemistry B

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The present study also indicates an apparent pKa value for the Hyp transition from a neutral to monodeprotonated form inside of BLMs.

a certain concentration, Hyp forms nonfluorescent clusters in the gel-phase DPPC membrane (at room temperature), whereas dynamic self-quenching of Hyp fluorescence is predominant in the sol-phase DPPC membrane (above the phase-transition temperature). Production of reactive oxygen species (superoxide anions and/or singlet oxygen) following photoexcitation of Hyp bound to liposomes was reported by several groups,14,24,25 and a binding constant for Hyp partition into DMPC liposomes was determined by Ehrenberg et al.14 The Hyp photodynamic action was found to rigidify DMPC liposomes,19 which is probably a consequence of the membrane lipid peroxidation. Two inspiring publications, reporting on theoretical studies of Hyp interaction with DPPC bilayers, have emerged in the recent years.26,27 Molecular dynamics calculations26 have indicated that neutral Hyp molecules, dissolved in the membrane bilayer, are preferentially located in the densest regions close to the polar head groups of the membrane with the bay area of Hyp pointing toward the water interface. It follows that the dipole moment of the S1 ← S0 transition, which is oriented along the short axis of the Hyp skeleton28 (Figure 1), is predominantly parallel to the membrane normal vector. It was also shown that the presence of cholesterol lowers the membrane permeability for Hyp.27 Most of the previous experiments on Hyp interaction with model membranes have been performed with liposomes or micelle systems.14,19,23−25 In this work, we take advantage of the definite spatial orientation of planar bilayer lipid membrane (BLM) systems to complement the mosaic of previous approaches. We present two sets of fluorescence-based measurements (see the schematic view in Figure 2) that reveal

2. EXPERIMENTAL SECTION 2.1. Materials. Dimethylsulfoxid (DMSO, 99.9%), chloroform (99.8%), and hexadecane (99%) were purchased from Sigma-Aldrich. Hypericin and 98.5% n-hexane were purchased from Invitrogen and Merck, respectively. 1,2-Diphytanoyl-snglycero-3-phosphocholine (DPhPC) powder was obtained from Avanti Polar Lipids. Stock solutions of DPhPC in chloroform (10 mg/mL) and Hyp in DMSO (1 × 10−3 M) were prepared. A solution of DPhPC in n-hexane was prepared by drying the chloroform solution of DPhPC in a glass flask (purging with nitrogen for 3 h) and by adding n-hexane to a final DPhPC concentration of 5 mg/mL. The electrolyte used in the experiments was 140 mM KCl and 10 mM HEPES at pHs ranging from 6 to 9 (adjusted by KOH). 2.2. Membrane Formation. The BLMs were formed in a Teflon cell (Eastern Scientific LLC, BC-20A) consisting of two identical 2 mL chambers. The chambers were separated by a 0.02 mm thick Teflon partition with a 0.1 mm diameter aperture at the center. The aperture edges were prepainted with a 1% v/v solution of hexadecane in n-hexane. BLMs were prepared by the Montall−Mueller method33 using the hexane solution of DPhPC. The quality of membranes was controlled by measuring their electric capacitance with the pulsed method described in the work of Meier et al.34 At the end of the experiments, short 1.5 V voltage pulses were used for membrane rupture. All of the measurements presented were carried out at room temperature (296 K). 2.3. Fluorescence Measurements. The optical arrangement used for fluorescence experiments (Figure 3) consisted of a laser-based excitation system and a detection part. Hyp molecules dissolved in the BLM membrane were excited either with a 594 nm cw laser (Cobolt: Mambo, 50 mW, linear polarization), the wavelength of which fits the absorption maximum of Hyp at about 600 nm, or with a 488 nm laser (Spectra Physics: Cyan, 100 mW). The laser power was weakened by filter F1 down to a level of 20−50 μW reaching the membrane. The stability of the laser power during the experiments was monitored by photodiode PD. The laser beam, after being reflected from the 50/50% beam splitter BS was focused into the membrane by an f = 20 mm aspheric lens L3. The estimated spot size of the beam on the membrane was approximately 25 μm. 2.3.1. Hyp Accumulation in BLM. After membrane formation, Hyp dissolved in DMSO was added to both sides of the BLM cell. The resulting concentration of DMSO in the electrolyte was kept at 0.1% during all experiments. Magnetic stirrers placed at the bottom of the two chambers were applied throughout the experiments (unless otherwise stated) to facilitate the transport of Hyp toward the membrane. After adding Hyp to the system, the increase of the fluorescence intensity (integrated over the spectral region transmitted through band-pass filter F3) from the membrane was monitored by a photomultiplier tube located at the PMT1 position and operated in a photon counting mode. The fluorescence signal was detected every 2 s until reaching steadystate conditions. The time needed for system equilibration was about 300−500 s. During this period, Hyp was first dispersed in water, forming nonfluorescent aggregates. Subsequently, these aggregates diffused35 toward the BLM through the unstirred

Figure 2. Scheme of the orientation and transport experiments. Hyp molecules preferentially located in the layers below the membrane surface are represented by arrows pointing toward the bay region. (a) The orientation of Hyp was examined by changing the angle of incidence α of the excitation laser beam (either s- or p-polarized). (b) An electric field was used to induce transport of Hyp anions within the membrane.

the preferential orientation and localization of Hyp within BLMs. First, we provide experimental evidence for Hyp orientation within BLMs by measuring its fluorescence intensity upon excitation with a laser beam at different angles of incidence (Figure 2a). The experimental data obtained are compared with the published results of molecular dynamics calculations of Hyp−BLM complexes.26 Second, we determine the characteristic time for the transport of deprotonated Hyp (bay-hypericinate) molecules (Figure 1) between the two opposite surfaces within the BLM. Hyp anion motion is induced by an electric field applied to the membrane (Figure 2b). Transport experiments of molecular ions inside of BLMs have been reported in a series of previous publications,29−32 in which the ion motion was detected by weak current transients. 1281



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expected to rise as the angle of incidence of the p-polarized laser beam increases and to remain unaffected when tilting the s-polarized beam. Laser beams of the same trajectory and orthogonal s and p polarizations were created in two separate branches of the beam path (delimited with beam samplers BS1 and BS2 in Figure 3) using orthogonal mirror reflections. The core adjustment of the laser angle of incidence on the membrane was achieved by linear translation of mirror TM, as indicated in Figure 3 by the dashed beam trajectory. The 1:1 telescope, composed of L1 and L2 lenses, was used for minor corrections for L3 spherical aberration. Translation of the L1 (which is in a plane conjugate to the L3) did not change the angle of beam incidence on the membrane. For each TM position, the beam spot was set to the membrane center by fine adjustment of the L1. The position of the laser spot on the membrane was viewed through a stereo microscope. The fluorescence signal was measured for each polarization and angle of incidence with PMT1. 2.3.3. Membrane Transport Experiments. In order to detect Hyp motion inside of BLMs, we performed another series of experiments. Hyp present in the membrane is either in a neutral or monodeprotonated state at the pH range from 6 to 9. It was assumed that Hyp anions were distributed and oriented inside of BLMs in a similar way to the prediction for the neutral form of Hyp in molecular dynamics simulations.26 We have applied rectangular electric field pulses to the BLMs to rearrange the distribution of hypericinate molecules between the two layers of the membrane (resulting in a higher Hyp− concentration on the more positive side of the membrane) (Figure 2b). Immediately after applying or changing the electric field, a net flux of Hyp− appeared between the two BLM sides until steady-state conditions were reached. During this period, Hyp anions were transported through the membrane central part, where they lost their preferred orientation. This was reflected in a transient increase of the fluorescence signal (with the excitation beam set perpendicular to the surface), which was detected in a time-resolved fluorescence experiment. The steady-state fluorescence was determined by local concentrations and corresponding Hyp aggregation states in the two layers. The time course of the fluorescence transient provided us with data on the kinetics of Hyp transport through the central part of the membrane. Ag/AgCl electrodes were used for applying repetitive voltage pulses to the membrane (generated by a Tektronix AFG3102 function generator) during the transport experiments. The transient fluorescence signal (following step-like changes in the electric field) was detected with the photomultiplier tube at position PMT2, where the spectral range was limited to 625 ± 15 nm by the spectrograph. The signal from the photomultiplier was acquired in a photon-counting mode by a Stanford Research Systems SR430 multichannel scaler/averager synchronized with the function generator. In order to minimize the noise level of the system, the magnetic stirrers were turned off during the transport measurements. Typically, the fluorescence signal acquired from 1000 consecutive voltage pulses was summed up in each measurement.

Figure 3. The optical setup used for fluorescence measurements. F1, F2: neutral density filters; BS1, BS2: beam samplers; S1, S2: manual shutters; MP: out-of-plane mirror pair; PD: photodiode; TM: translating mirror for beam angle adjustment; L1, L2, L3: lenses; BS: beam splitter; PMT1, PMT2: photomultiplier tubes; F3 (used with PMT1): band-pass filter (624 ± 20 nm); F4 (used with 488 nm excitation): long-pass laser-line edge filter; DM (used with PMT1): short-pass 600 nm dichroic mirror.

water layer36 adjacent to the cell walls, and finally, Hyp molecules reaching the membrane were incorporated into the BLM. Hyp molecules were also dissolved in other phospholipid layers deposited on the cell walls, dispersed in the electrolyte, and/or localized at the water/air interface. In all experiments, even at the highest Hyp concentrations, most of Hyp molecules were located in the lipid phase (eventually, some Hyp molecules stuck to the Teflon surface) after reaching steadystate conditions. For the sake of simplicity, the amount of Hyp used is denoted by the bulk concentration cbulk that corresponds to Hyp present in the whole experimental system (water + lipid phase). Fluorescence spectra of Hyp accumulated in the membrane were measured at 488 nm excitation by an Acton Research Corporation SpectraPro-300i spectrograph equipped with Princeton Instruments Spec-10:400 TE cooled CCD camera (Figure 3). 2.3.2. Spatial Orientation Measurements. The spatial orientation of Hyp molecules inside of BLMs was investigated using excitation at 594 nm. The Hyp fluorescence response was measured using s- and p-polarized laser beams with a different angle of incidence to the membrane surface. Figure 2a shows the situation in which Hyp molecules are predominantly oriented with the bay regions pointing out from the membrane. With the laser beam directed perpendicular to the membrane plane, the absorption (and consequently the fluorescence intensity) was minimized due to the dominantly orthogonal orientation of the laser polarization to the dipole moment of the Hyp S1 ← S0 transition. The fluorescence intensity is

3. RESULTS AND DISCUSSION 3.1. Hyp Accumulation in BLM. The fluorescence spectrum of Hyp incorporated in BLM together with the fluorescence of Hyp monomers in DMSO solution is shown in Figure 4a. The two spectra are almost identical, which confirms that the membrane fluorescence is due to Hyp monomers. A 1282



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Figure 5. Fluorescence intensity of Hyp incorporated in BLMs as a function of the angle of incidence of the excitation beam. Experimental data obtained with DPhPC membranes (pH = 7.4; cbulk = 0.75 × 10−6 M; excitation wavelength: 594 nm; detection region: 624 ± 20 nm) are plotted with solid squares and open circles for p- and s-polarized laser light, respectively. Theoretical predictions based on molecular dynamic simulations of Hyp in DPPC membranes26 are also indicated; the dashed and dotted curves represent limiting cases of “motionless” and fully “randomized” Hyp molecules. Presented data were normalized to the zero incident angle limit. Figure 4. (a) Fluorescence spectra of Hyp dissolved in the DPhPC membrane (pH = 7.4; cbulk = 0.75 × 10−6 M) and in DMSO (c = 0.75 × 10−6 M) measured at 488 nm excitation. (b) Steady-state fluorescence intensity of Hyp incorporated into the DPhPC membrane at different bulk Hyp concentrations (pH = 7.4; excitation wavelength: 594 nm; detection band: 624 ± 20 nm).

fluorescence intensity. Changing α for the s polarization does not change the orientation of the light wave’s electric field, thus leaving the fluorescence intensity constant. The present experimental data obtained from Hyp−DPhPC complexes were compared with the results of molecular dynamic simulations carried out for Hyp incorporated in DPPC bilayers by Eriksson et al.26 For the purpose of obtaining the variation of the absorbance of light in the membrane with the angle of incidence of the p-polarized laser beam, the computed distribution of the molecular axis tilt angle was integrated for each azimuthal angle, taking into account the mutual orientation of the light polarization and Hyp transition dipole moment vectors. As a next step, the emission of the excited Hyp molecules into the spatial angle determined by the detection system of our apparatus was calculated. Here, we assumed that the quantum yield of Hyp fluorescence was independent of the molecule’s orientation within the membrane. In principle, there are two limiting cases. In the first case, Hyp molecules were assumed to be motionless during the lifetime of the singlet excited state, which is on the order of a few nanoseconds.37 In the second limiting situation, the orientation of excited Hyp molecules was fully randomized before the emission event. The theoretical predictions of the (relative) Hyp fluorescence intensity based on the computed tilt angle distribution26 are plotted in Figure 5, with dashed and dotted lines standing for the motionless and fully randomized limiting cases, respectively. The measured data correspond well with the calculated dependence of slowly rotating (motionless) Hyp molecules. In general, the calculations predict a more pronounced increase of the fluorescence intensity toward the higher angle of incidence of the excitation laser beam, which indicates a more narrow distribution of the molecular axis tilt angle of Hyp molecules in DPPC membranes as compared to that of DPhPC bilayers. 3.3. Electric-Field-Driven Hyp Transport Inside of the Membrane. During the transport experiments, deprotonated Hyp molecules were redistributed inside of BLMs by means of an electric field applied to the membrane. In the first series of experiments, the electric field was periodically switched

4−5 nm blue shift of Hyp fluorescence maxima in the membrane is observed as compared to the maxima in DMSO solution. The same observation was reported for Hyp dissolved in the lipid part of LDL particles.9 The steady-state fluorescence intensity of Hyp incorporated in BLM as a function of bulk Hyp concentration at pH 7.4 is shown in Figure 4b. The fluorescence intensity reaches a maximum and then decreases with the increase of Hyp concentration. This finding is attributed to the dynamic selfquenching, and/or Hyp self-aggregation in the membrane at high local Hyp concentrations. Similar behavior was observed for Hyp accumulation inside of LDL molecules.9 Because only the monomeric form of Hyp is active in photodynamic applications, it can be hypothesized that the highest fluorescence intensity also indicates optimal conditions for the maximal production of singlet oxygen by Hyp.15 This is why all of the following experiments have been carried out with cbulk = 0.75 × 10−6 M, which corresponds to the most intense steady-state fluorescence observed from the Hyp−BLM complex. 3.2. Orientation of Hyp Monomers in the Membrane. The fluorescence signal from the BLM−Hyp complex plotted against the angle of incidence α of the excitation beam is shown in Figure 5. The measured fluorescence intensity has been multiplied for both s and p polarizations by cos(α) to account for the increase of the illuminated membrane area at higher α. The fluorescence signal increases with the increasing α for the p-polarized beam and is independent of α for the s polarization. As the short molecular axis, which corresponds to the dipole moment of the S1 ← S0 transition of Hyp, is predominantly perpendicular to the membrane surface, a higher angle of incidence corresponds to higher absorption probability for the p-polarized beam. This is reflected in the increase of the 1283



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between +120 and −120 mV, as shown in Figure 6a. The corresponding Hyp fluorescence signal (cbulk = 0.75 × 10−6 M)

pulse, as shown in Figure 6b, and avgAB is the overall average of the fluorescence signal. F is plotted in Figure 6c as a function of pH. Despite of data scattering, there is an indication that Hyp is dominantly deprotonated above pH 8 and neutral (not affected by the electric field) at lower pH values. It means that the apparent pKa constant for Hyp deprotonation in the DPhPC membrane is around 8. This finding is in relatively good correlation with the previous results of Jardon et al.38,39 They found that in micellar dispersions, the first deprotonation of Hyp has an apparent pKa value of 7. However, the pKa constant for Hyp deprotonation in the bay region has been reported to be 1.8 in aqueous environment.7 The second set of transport experiments has been carried out at pH 8.5 using 200 mV positive voltage pulses with a 200 ms duration followed by 500 ms zero-field periods (see Figure 7a).

Figure 6. (a) Time dependence of the voltage applied periodically to the membrane. (b) The transient fluorescence intensity of Hyp (pH = 8.4). A and B denote the first and second half periods of the voltage pulse. (c) The value of the fluorescence signal modulation function F during the transient (see the text) as a function of the electrolyte pH.

is shown in Figure 6b. The time needed for charging the membrane was about 1 μs, as determined by the transient current in the system (not shown). It was observed that after changing the electric field polarity, the fluorescence signal first increased, reached a maximum (at about 25 ms), and then relaxed back to a steady-state value. This observation can be explained as follows. After changing the electric field polarity, Hyp anions are forced to rearrange between the two sides of the membrane, resulting in their higher concentration at the positive side. There are two possible mechanisms that may contribute to the fluorescence transient during this transport process. On one hand, Hyp anions passing through the central part of the membrane lose their preferential orientation, which is reflected in higher absorption and higher fluorescence efficiency. On the other hand, because of using a Hyp concentration of cbulk = 0.75 × 10−6 M, the highest steadystate fluorescence intensity corresponds to the situation in which Hyp molecules are distributed evenly between the two sides of BLM. Due to the applied electric field, the Hyp concentration is decreased at the negative side and increased at the positive one, both resulting in a lower steady-state fluorescence intensity (see Figure 4b). On the contrary, during the transient period, the Hyp concentration is (at a given moment) equal on both sides of the membrane, which leads to the maximum fluorescence intensity. More pronounced fluorescence modulation was observed at higher pH values of the electrolyte solution. The relative modulation has been characterized by the ratio of F = (avgA − avgB)/avgAB, where avgA and avgB are the average fluorescence intensities during the first and the second halves of the voltage

Figure 7. (a) Time dependence of the voltage applied periodically to the membrane. (b) The transient fluorescence intensity (pH = 8.5). τ1 and τ2 indicate the characteristic time of exponential fits (solid lines) in the intervals A−B and C−D, respectively. (c) The characteristic exponential decay times shown as a function of the applied voltage.

At the end of the period with no electric field applied, Hyp anions are distributed evenly between the two sides of the membrane, and the fluorescence is stabilized at a constant I0 value; see the dashed line in Figure 7b. After applying the positive voltage pulse (at t = 0), the Hyp fluorescence intensity almost immediately jumps higher (point A) and then relaxes to a new steady-state value below the original level (point B). The early increase of the intensity is presumably due to Hyp moving through the membrane center, where Hyp molecules are not oriented with bay regions preferentially pointing toward the aqueous surrounding. The subsequent decrease of the fluorescent signal reflects the process of Hyp aggregation and/or self-quenching of Hyp fluorescence at the positive side of the membrane combined with the decrease of Hyp concentration at the negative side. When the voltage is turned off, the fluorescence intensity almost instantaneously increases 1284



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to a value above I0 (point C) and then gradually falls back to the original level (point D). By fitting the signal in the intervals A−B and C−D by single-exponential decays, the characteristic times of Hyp transport through the membrane center were determined with (τ1) and without (τ2) the electric field (Figure 7c). It should be noted that the obtained values may reflect Hyp membrane transport in combination with formation/decay of Hyp aggregates. The characteristic time τ2 belonging to the zero-voltage situation is 70 ms. This is an upper limit for the transport of Hyp molecules through the membrane center, which gets faster as the voltage increases (see τ1 in Figure 7c). The shape of the τ1 curve does not extrapolates to τ2 in the zero-voltage limit, which is in a good qualitative agreement with the theoretical predictions and experimental data presented by Anderssen et al.30

11. This work was also supported by the projects CEVA (26220120040) (30%), SEPO-II (26220120039) (30%), and NanoBioSens (26220220107) (30%) of the Operation Programme Research and Development funded by the European Regional Development Fund.

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(1) Theodossiou, T. A.; Hothersall, J. S.; De Witte, P. A.; Pantos, A.; Agostinis, P. Mol. Pharm. 2009, 6, 1775−1789. (2) Kiesslich, T.; Krammer, B.; Plaetzer, K. Curr. Med. Chem. 2006, 13, 2189−2204. (3) Karioti, A.; Bilia, A. R. Int. J. Mol. Sci. 2010, 11, 562−594. (4) Agostinis, P.; Vantieghem, A.; Merievede, W.; De Witte, P. A. Int. J. Biochem. Cell Biol. 2002, 34, 221−241. (5) Miskovsky, P. Curr. Drug Targets 2002, 3, 55−84. (6) Kramer, B.; Verwanger, T. Current Med. Chem. 2012, 19, 793− 798. (7) Falk, H. Angew. Chem., Int. Ed. 1999, 38, 3116−3136. (8) Falk, H.; Meyer, J. Monatsch. Chem. 1994, 125, 753−762. (9) Kascakova, S.; Refregieurs, M.; Jancura, D.; Sureau, F.; Maurizot, J. C.; Miskovsky, P. Photochem. Photobiol. 2005, 81, 1395−1403. (10) Huntosova, V.; Nadova, Z.; Dzurova, L.; Jakusova, V.; Sureau, F.; Miskovsky, P. Photochem. Photobiol. Sci. 2012, 11, 1428−1436. (11) van de Putte, M.; Roskans, T.; Bormans, G.; Verbruggen, A.; de Witte, P. A. Int. J. Oncol. 2006, 28, 655−660. (12) Bourig, H.; Eloy, D.; Jardon, P. J. Chim. Phys. 1992, 89, 1391− 1411. (13) Roslianec, M.; Weitman, H.; Freeman, D.; Mazur, Y.; Ehrenberg, B. J. Photochem. Photobiol., B 2000, 57, 149−158. (14) Ehrenberg, B.; Anderson, J. L.; Foote, Ch. S. Photochem. Photobiol. 1998, 68, 135−140. (15) Gbur, P.; Dedic, R.; Chorvat, D., Jr.; Miskovsky, P.; Hala, J.; Jancura, D. Photochem. Photobiol. 2009, 85, 816−823. (16) Gbur, P.; Dedic, R.; Jancura, D.; Miskovsky, P.; Hala, J. J. Lumin. 2008, 128, 765−767. (17) Miskovsky, P.; Sureau, F.; Chinsky, L.; Turpin, P. Y. Photochem. Photobiol. 1995, 62, 546−549. (18) Siboni, G.; Weitman, H.; Freeman, D.; Mazur, Y.; Malik, Z.; Ehrenberg, B. Photochem. Photobiol. Sci. 2002, 1, 483−491. (19) Chaloupka, R.; Obsil, T.; Plasek, J.; Sureau, F. Biochim. Biophys. Acta 1999, 1418, 39−47. (20) Theodossiou, T. A.; Noronha-Dutra, A.; Hothersall, J. S. Int. J. Biochem. Cell Biol. 2006, 38, 1946−1956. (21) Ali, S. M.; Olivo, M. Int. J. Oncol. 2002, 21, 531−540. (22) Vandenbogaerte, A. L.; Cuveele, J. F.; Himpens, P. B.; Merlewede, W. J.; de Witte, P. A. J. Photochem. Photobiol., B 1997, 38, 136−142. (23) Ho, Y. F.; Wu, M. H.; Cheng, B. H.; Chen, Y. W.; Shih, M. Ch. Biochim. Biophys. Acta 2009, 1788, 1287−1295. (24) Losi, A. Photochem. Photobiol. 1997, 65, 791−801. (25) Hadjur, C.; Jardon, P. J. Photochem. Photobiol. 1995, 29, 147− 156. (26) Eriksson, E. S.; Dos Santos, D. J.; Guedes, R. C.; Eriksson, L. A. J. Chem. Theory Comput. 2009, 5, 3139−3149. (27) Eriksson, E. S.; Eriksson, L. A. J. Chem. Theory Comput. 2011, 7, 560−574. (28) Yamazaki, T.; Ohta, N.; Yamazaki, I.; Song, P. J. Phys. Chem. 1993, 97, 7870−7875. (29) Ketterer, B.; Neumcke, B.; Läuger, P. J. Membr. Biol. 1971, 5, 225−245. (30) Anderssen, O.; Fuchs, M. Biophys. J. 1975, 15, 795−830. (31) Wulf, J.; Benz, R.; Pohl, W. G. Biochim. Biophys. Acta 1977, 465, 429−442. (32) Waggoner, A. S.; Wang, C. H.; Tolles, R. L. J. Membr. Biol. 1977, 33, 109−140. (33) Montal, M.; Mueller, P. Proc. Natl. Acad. Sci. U.S.A. 1972, 69, 3561−3566.

4. CONCLUSIONS Hyp interaction with DPhPC membranes has been investigated in a series of fluorescence measurements. Fluorescence spectra of BLM−Hyp complexes have revealed that, at low concentration, Hyp is present in the membrane in its monomer form. Above a critical Hyp concentration, the intensity of Hyp fluorescence gradually decreases as a consequence of Hyp aggregation and/or self-quenching inside of the lipid membrane. It has been shown that Hyp monomers incorporated into DPhPC bilayers are predominantly oriented with the S1 ← S0 transition dipole moments perpendicular to the membrane surface. This observation is in good qualitative agreement with the predictions of molecular dynamics simulations carried out by Eriksson et al. for DPPC bilayers.26 According to molecular dynamics calculations,26 Hyp molecules are localized within DPPC BLMs in two opposite layers close to the membrane/ water interface. Transport of Hyp anions between these two layers of the DPhPC membrane was induced in our experiments by electric field pulses. It has been proven that time-resolved fluorescence measurements are suitable for monitoring the motion of molecules during the transport process. The characteristic time for Hyp transport through the membrane center was 70 ms at the zero-voltage limit. The transport of Hyp anions became faster with higher voltage applied to the membrane. Further modeling of Hyp anions would be desirable for detailed description of the observed electric-field-induced transport phenomena. In conclusion, the results of this work may provide useful information for the elucidation of the transmembrane transportation mechanism and intracellular localization of Hyp. The experimental methods presented can be also used for investigation of other hydrophobic molecules/ions inside of BLMs.

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REFERENCES

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Phone: +421-55 234 2595. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors would like to thank K. Stroffekova, G. Fabriciova, and J. Ulicny for helpful discussions. This work was supported by the grants of the Slovak Ministry of Education VEGA 1/ 1246/12, VEGA 1/1154/11, LPP-0290-09, and APVV-02421285



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(34) Meier, W.; Graff, A.; Diederich, A.; Winterhalter, M. Phys. Chem. Chem. Phys. 2000, 2, 4559−4562. (35) Bano, G.; Stanicova, J.; Jancura, D.; Marek, J.; Bano, M.; Ulicny, J.; Strejckova, A.; Miskovsky, P. J. Phys. Chem. B. 2011, 115, 2417− 2423. (36) Barry, P. H.; Diamond, J. M. Physiol. Rev. 1984, 64, 763−872. (37) Mukherjee, P.; Adhikary, R.; Halder, M.; Petrich, J. W.; Miskovsky, P. Photochem. Photobiol. 2008, 84, 706−712. (38) Jardon, P.; Gautron, R. J. Chim. Phys. 1989, 86, 2173−2190. (39) Eloy, D.; Le Pellec, A.; Jardon, P. J. Chim. Phys. 1996, 93, 442− 457.

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Spatial Orientation and Electric-Field-Driven Transport of Hypericin

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