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Jul 6, 2017 - Separation Yield of Donor−Acceptor Linked Molecules in Biological. Environment via Modulating Their Cationic Moieties. Ning Cai,. †,...
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Strategy to Attain Remarkably High Photoinduced ChargeSeparation Yield of Donor−Acceptor Linked Molecules in Biological Environment via Modulating Their Cationic Moieties Ning Cai,†,‡,¶ Yuta Takano,†,¶ Tomohiro Numata,§,∥ Ryuji Inoue,§ Yasuo Mori,*,∥ Tatsuya Murakami,†,⊥ and Hiroshi Imahori*,†,# †

Institute for Integrated Cell-Material Sciences (WPI-iCeMS), Kyoto University, Sakyo-ku, Kyoto 606-8501, Japan School of Chemical Engineering and Light Industry, Guangdong University of Technology, Guangzhou 510006, China § Department of Physiology, Graduate School of Medical Sciences, Fukuoka University, Nanakuma 7-45-1, Johnan-ku, Fukuoka 814-0180, Japan ∥ Department of Synthetic Chemistry and Biological Chemistry, Graduate School of Engineering, Kyoto University, Nishikyo-ku, Kyoto 615-8510, Japan ⊥ Department of Biotechnology, Faculty of Engineering, Toyama Prefectural University, 5180 Kurokawa, Imizu, Toyama 939-0398, Japan # Department of Molecular Engineering, Graduate School of Engineering, Kyoto University, Nishikyo-ku, Kyoto 615-8510, Japan ‡

S Supporting Information *

ABSTRACT: A series of ferrocene−porphyrin−fullerene linked triads (TC1, TC2, and TC4) possessing different numbers of cationic moieties were designed and prepared to achieve a high photoinduced charge-separation (CS) yield in a biological environment. In a solution, TC1, TC2, and TC4 demonstrated the formation of their nanoaggregates. Among the new triads, TC4 possessing the four cationic moieties exhibited the formation of a long-lived charge-separated state with the highest CS yield (86%) ever reported in cell membrane-like lipid bilayers, which is consistent with the largest change in the cell membrane potential of PC12 cells via the photoinduced CS under green light illumination. The highest CS yield in the biological environment can be rationalized by the well-tailored balance in hydrophobicity and hydrophilicity of TC4. This finding provides a strategy to improve greatly the photoinduced charge-separation yield of donor− acceptor linked molecules in the biological environment and also will be informative for extracting the full potential of the photoinduced charge-separated state toward biological applications.

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INTRODUCTION Photoinduced electron-transfer (PET) reactions are a key process occurring in natural photosynthesis and organic photovoltaics.1−4 PET between an electron donor and an acceptor results in formation of the corresponding donor radical cation and acceptor radical anion which can induce local electric field or drive oxidation and reduction reactions, respectively.5 Recent synthetic efforts have been devoted to establishing highly efficient artificial donor−acceptor (D−A) systems in which PET is optimized in terms of a photoinduced chargeseparation (CS) yield and lifetime toward efficient conversion of light energy into chemical or electrical energy in organic solvents or in solid films.6,7 In comparison, few PET reactions of synthetic compounds were studied in biocompatible mediums, such as highly polar aqueous solutions or lipid bilayers.8−14 Taking into account the importance of PET in a © XXXX American Chemical Society

biological environment, especially photoinduced CS in photosynthetic reaction centers and potential applications of PET to control biological functions,15−17 nanomaterials and organic compounds that utilize PET reactions have recently captured considerable attention.14,18,19 The exclusive use of PET is also expected to reduce undesirable side effects which would diminish the therapeutic effects, e.g., generation of harmful singlet oxygen by photoinduced energy transfer (PEN).20 Nevertheless, the preceding CS yield and lifetime in D−A systems have not yet been sufficiently high and long, respectively, to utilize full potential of PET in artificial and biological membranes, although they have been achieved in solutions.21,22 For instance, to the best of our knowledge, the Received: May 10, 2017 Revised: July 6, 2017

A

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The Journal of Physical Chemistry C highest, but moderate CS yield of 67% in lipid bilayer membranes was presented by O’Brien and co-workers in 1994.11 Meanwhile, we reported the amphiphilic D−A linked molecules consisting of ferrocene−zincporphyrin−fullerene (Fc−ZnP−C60) triads,18,19 which were exploited to control the cell membrane potential via the photoinduced chargeseparated state. The triads, however, demonstrated rather low CS yields up to 27% in the artificial lipid bilayer. The other biomimetic system which effectively utilized PET also generated the CS yield of 10% with a light-driven proton pumping for production of ATP by D−A linked molecules.9 In this context, achieving the high CS yield in the biological environment while keeping the sufficiently long lifetime is still a challenge toward utilizing PET in biological systems. In this study, a new series of the Fc−ZnP−C60 triads, TC1, TC2, and TC4, were designed to address the above issues (Figure 1). Multiple trimethylammonium cationic moieties were introduced into the hydrophobic triad core to (i) tune the amphiphilic nature of the molecules, (ii) suppress the undesirable aggregation behavior in the biological environment, and (iii) enhance the binding of the triads to the negatively charged plasma membrane.

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EXPERIMENTAL SECTION General Methods. 1H NMR spectra were measured by a JEOL JNM-EX400 NMR spectrometer or a Bruker AVANCE 500 spectrometer for TC1, TC2, and TC4. Electrospray ionization (ESI) mass spectra were measured via a Fouriertransform mass spectrometer with an electrospray ionization source. Atmospheric pressure chemical ionization (APCI) mass spectra were recorded via a Fourier-transform mass spectrometer with an atmospheric pressure chemical ionization source. Matrix-assisted laser desorption−ionization (MALDI) time-offlight mass spectra were obtained by a Fourier-transform mass spectrometer using a MALDI source with 1,8-dihydroxy9(10H)-anthracenone (DIT) as a matrix. Infrared (IR) spectra were recorded on a JASCO FT/IR-470 Plus spectrometer. Statistical analysis was performed using Igor Pro v6.3 using a one-way ANOVA with a Dunnett’s post hoc test for significance. Ultraviolet−Visible Absorption and Fluorescence Measurements. Static UV−visible absorption and fluorescence spectra were recorded using a PerkinElmer Lambda 900 spectrometer and a HORIBA SPEX Fluoromax-3 spectrofluorometer, respectively. In fluorescence measurements, the excitation wavelength was set at 430 nm. Atomic Force Microscopy (AFM). AFM images were obtained with a MFP-3D-SA atomic force microscope (Asylum Technology) in AC mode. In all AFM measurements, AC240TS micro cantilevers (Olympus) featured a force constant of 1.7 N m−1 and a nominal tip radius of less than 10 nm. The samples were deposited onto a cleaved mica substrate from the DMSO/H2O (1/99, v/v) solution. The excess solution was removed by spin-coating at 1500 rpm for 12 s and 2200 rpm for 6 s, followed by washing with water; the samples were then dried in air. All measurements were performed in air. Particle size was evaluated as follows. Acquired images were analyzed by ImageJ23 with the function of “analyze particle” on a 16-bit monochrome image to determine a mean height and an area of the particles. Then, the size of the particles, which were deemed to be spheres, was calculated from the volume of the particles.

Figure 1. Molecular structures of TC1, TC2, and TC4 and reference molecules Fc-ZnP-C60-ref, TC1-ref, and ZnP-ref.

Theoretical Calculations. Geometries were optimized using the Gaussian 09 (revision D.01) program24 with the B3LYP25−28 functional with the D3 version of Grimme’s dispersion with the original D3 damping function.29 The 631G(d)30,31 basis set was used for H, C, N, and O atoms, and Los Alamos ECP plus DZ32 was used for Fe and Zn atoms. B

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The Journal of Physical Chemistry C Table 1. CR Rate Constants (k) and CS Yields (Φ) of TC1, TC2, TC4, and TC1-refa medium benzonitrile DMSO/H2Od

liposome in DMSO/H2Od

compd. Fc-ZnP-C60-ref TC1-ref TC1 TC2 TC4 TC1-ref TC1 TC2 TC4

k1 (×105 s−1) 1.0 5.3 6.2 4.5 3.6 2.5 5.6 5.0 3.7

b

A1 (%) b

100 53 58 62 60 53 55 62 59

k2 (×104 s−1)

A2 (%)

n.d. 3.2 4.5 4.0 2.4 2.6 3.6 3.7 3.8

n.d. 47 42 38 40 47 45 38 41

Φ 0.99b 0.57 ± 0.50 ± 0.50 ± 0.51 ± 0.27 ± 0.18 ± 0.38 ± 0.86 ±

0.03c 0.03c 0.03c 0.03c 0.01c 0.01c 0.02c 0.04c

a Data were obtained from the decay profiles of C60•− (λex= 550 nm, λobs = 1000 nm) by a single or biexponential fitting. SE of the data shown here is less than 5%. bData from ref 22. cValues were obtained by comparing the absorbances at 1000 nm under the same conditions with taking account of the different molar absorption coefficients caused by the solvent system used for the TA measurements.19,22 dDMSO/H2O, 1/99, v/v.

washed once with HBSS(+), and filled with HBSS(+). Then, an aqueous solution of the compound was added to the cells to adjust the final concentration to 0.5 μM based on the compound. The cells were incubated for 3 min and washed by HBSS once and filled by HBSS(+). Confocal images were recorded by Zeiss LSM780 (Carl Zeiss Microscopy GmbH, Jena, Germany), which was equipped with a 40× oil objective lens (NA 0.75), GaAsP (32×), array and femtosecond pumped pulsed laser. Images of the compound were obtained by using spectral imaging with linear unmixing. Evaluation on the Amount of Incorporated Triads in a Single Cell. PC12 cells (8 × 105 cells) were seeded on a polyL-lysine (PLL)-coated six-well plate (Corning, MA) and cultured for 24 ± 3 h. After the medium was removed, the cells were washed twice with HBSS(−) and filled with HBSS(−). Then, an aqueous solution of the triads was added to the cells to adjust the final concentration to 0.6 μM based on the triads. The cells were incubated for 3 min and washed by PBS(−) and dissolved by 0.2% TritonX-100 aq and dried in a vacuum freeze-dryer (Eyela, Tokyo, Japan). The residue was extracted by DMSO, and the extract was subjected to the absorption spectrometer. Total number of the incorporated molecules was obtained by the relative integration value of the Soret band intensity to the reference (2.0 μM DMSO solution of the corresponding compound). Then, the number was divided by the number of the cells which were treated similarly by HBSS(−) instead of the aqueous solution of the compound. Detection of Photoinduced Reactive Oxygen Species (ROS). Detection of ROS with singlet oxygen sensor green reagent (SOSGR) for 1O2 (Thermo Fisher Scientific Inc., MA) was performed on the basis of the manufacture’s protocol. Briefly, 1.0 μM triad containing 5.0 μM SOSGR in deionized H2O containing 1.0% DMSO was irradiated in a cuvette for 4 min by Xe lamp (7.0 mW cm−2) with a band-pass filter (545− 555 nm). The resulting solutions were immediately subjected to the spectrofluorometer with an excitation wavelength of 480 nm. The measurements were repeated three times. Detection of Photoinduced Cell Damage. Detection of cell necrosis or apoptosis was carried out by using Annexin VFITC Apoptosis Kit (BioVision, Inc., CA), which contains Annexin V-FITC conjugate for early apoptosis detection and propidium iodide (PI) for late apoptosis and necrosis detection, according to the manufacture’s protocol. Briefly, the cells were seeded at PLL coated glass bottom dish with 1.0 × 105 cells mL−1 and cultured for 1 day. Then, the medium was removed, washed with HBSS(−) twice and filled with HBSS(−). Then, aqueous solution of the compound was added to the cells to the

Incorporation of the Triads into Liposome. To an aqueous solution of COATSOME EL-11-A (Lipid composition: POPC:Cholesterol:POPG = 30:40:30 mol %, NOF co., Tokyo, Japan), a DMSO solution of TC1, TC2, TC4, or TC1ref was added at a concentration of 2 mol % relative to the lipids, and the solution was incubated at 37 °C for 3 min. The incorporation was monitored by following the change in the absorbance of the Soret band. Typically, it was completed within 1 h. Nanosecond TA Measurements. Nanosecond timeresolved TA measurements were carried out using the laser system (Unisoku Co., Ltd., Osaka, Japan). Solvents were deaerated by argon bubbling for 5 min before measurements. A solution containing the triads or a mixture of the triads and COATSOME EL-11-A (molar ratio: 1/50) was excited by a Panther OPO pumped by a Nd:YAG laser (Continuum SLII10, 4−6 ns fwhm), and the photodynamics were monitored by continuous exposure to a xenon lamp (150 W) as a probe light and a photomultiplier tube (Hamamatsu 2949) as a detector. The instrument response function (IRF) for this system is less than 10 ns (ca. 9 ns). The CS yields (Φ) in Table 1 were determined by the comparative method, which uses nanosecond TA spectra and the following equation:19,35 Φ = Φref ×

ΔODmax‐sample ΔODmax‐ref

where Φref is the CS yield of the reference compound (0.99 for Fc-ZnP-C60-ref in PhCN),22 ΔODmax‑sample and ΔODmax‑ref are the maximum ΔOD intensities at 1000 nm of the compound of interest and the reference compound, respectively. Difference of the molar extinction coefficients and the influence of the aggregation in the solvent systems are taken into account with the data of the fullerene radical anion obtained in the previous study.19 Standard error (SE) of the data obtained here is less than 5%. PC12 Cell Culture. Rat pheochromocytoma (PC12) cells were cultured in Dulbecco’s modified Eagle’s minimal essential medium (DMEM) with high glucose (Sigma-Aldrich) supplemented with 5% Fetal bovine serum (FBS) (Japan Bioserum, Hiroshima, Japan) and 5% horse serum (HS) (Thermo Fisher Scientific Inc., MA, United States). The cell culture media contained 30 U mL−1 penicillin and 30 μg mL−1 streptomycin (Thermo). The cells were cultured in 5% CO2 and 95% air and passaged every 3−4 days. PC12 cells were used for the following experiments unless stated. Confocal Microscopy. PC12 cells were cultured on PLL coated glass bottom dish. After 1 day, medium was removed, C

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The Journal of Physical Chemistry C final concentration of 1.8 μM based on the compound. The cells were incubated for 3 min at room temperature and washed by HBSS(−) once and filled by HBSS(−), followed by the illumination of the Xe lamp (7.0 mW cm−2, 545−555 nm) for 4 min. After the irradiation, the cells were treated by the Annexin V-FITC/PI solution and then observed by a Fluoview-10i (Olympus). Fluorescent cells were counted as cells in early apoptosis (green) or those in late apoptosis or necrosis (red or yellow). For one experiment, more than 500 cells were counted from randomly chosen three images (more than 150 cells per an image). Assays were performed three times. Patch Clamp Measurements. Whole-cell patch recordings for current clamp were recorded using nystatin-perforated patch technique on PC12 cells at room temperature (22−25 °C) with Axopatch 200B patch-clamp amplifier (Molecular Devices, CA). On the basis of the correlation depicted in Figure S11, an aqueous solution of the triad (0.34 μM for TC1-ref, 0.55 μM for TC1, 0.23 μM for TC2, 0.14 μM for TC4) was added to the cells in the buffer solution. For the whole-cell recordings, the Na+-based bath solution contained (in mM) 140 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, 10 HEPES, and 10 Dglucose (pH adjusted to 7.40 with NaOH, and osmolality adjusted to 320 mosmol kg−1 H2O with D-mannitol). The pipet solution contained (in mM) 55 K2SO4, 20 KCl, 5 MgCl2, 0.2 EGTA, and 5 HEPES (pH adjusted to 7.40 with KOH, and osmolality adjusted to 300 mosmol kg−1 H2O with Dmannitol). Photoinduced Vm was calculated by the following equation; Vm (mV) = Vhν − VCtl, where Vhν and VCtl are membrane potentials recorded after and before illumination, respectively (ultrahigh pressure Hg lamp, 7.0 mW cm−2, 545− 555 nm).

Figure 2. UV−visible absorption spectra of (a) TC1 (black, solid), (b) TC2 (blue, solid), and (c) TC4 (red, solid) in DMSO and of (d) TC1 (black, dotted), (e) TC2 (blue, dotted), and (f) TC4 (red, dashed) in DMSO/H2O (1/99, v/v). Inset shows the expanded absorption spectra around the Q-band.

(Figure 2 and Table S1). These suggest that the size of the nanoaggregates decreases in the same order,33,34 which is rationalized by the increase in the hydrophilicity of the triad molecules with increasing the number of the cationic moieties. The nanoaggregates of TC1, TC2, and TC4 were visualized by atomic force microscopy measurements (Figure S6). A large number of particles were observed probably because of the strong interaction between the negatively charged mica surface and the positively charged nanoaggregates of the triads with the cationic moieties. The AFM image analysis shows the nanoaggregate size of 12 ± 5 nm for TC1, 9 ± 4 nm for TC2, and 3 ± 1 nm for TC4 (Figure 3), indicating the domain size order of TC1 > TC2 > TC4, which agrees with the observation in the absorption measurements. On the basis of the structure of TC1 optimized by density functional theory (DFT), the molecular lengths of TC1, TC2, and TC4 are predicted to be ca. 4 nm (Figure S7). The mean size of TC4 (3 nm) is comparable to the predicted length and is much smaller than those (9−10 nm) of the previously reported analogous triads without the multicationic moieties,19 realizing the formation of the small nanoaggregates consisting of a few molecules of TC4. Nanosecond time-resolved transient absorption (TA) measurements were conducted for TC1, TC2, TC4, and TC1-ref in DMSO/H2O under green light excitation (550 nm). Figure 4 displays the TA spectra of TC1, TC2, TC4, and TC1-ref in DMSO/H2O. The fingerprint of C60 radical anion (C60•−) around 1000 nm is observed, while the characteristic absorptions of ZnP•+ at 600−700 nm and Fc+ at 800 nm are not detectable owing to the fast decay of ZnP•+ (∼109 s−1) and small molar absorption coefficient of Fc+,22,37,38 suggesting the selective formation of the final charge-separated state (Fc+− ZnP−C60•−). Namely, initial CS takes place from the singlet excited-state of ZnP (1ZnP*) to C60 to generate Fc−ZnP•+− C60•−, followed by the fast charge-shift from Fc to ZnP•+, yielding Fc+−ZnP−C60•−, as we demonstrated before.22 The charge-recombination (CR) rate constants of TC1, TC2, TC4, and TC1-ref were determined by analyzing the decay profiles at 1000 nm (Table 1). The CS yields were also obtained by the comparative method with Fc−ZnP−C60-ref as the reference (99% in benzonitrile).19,21,22 It is noteworthy here that the calculation of the CS yield based on rate constants acquired from femtosecond−picosecond transient absorption spectro-

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RESULTS AND DISCUSSION The new triads, TC1, TC2, and TC4, were synthesized according to the synthetic route shown in Figure S1 (see the Supporting Information). Elaborately designed aniline derivatives featuring different numbers of bromoalkyl groups were linked to both sides of a zinc porphyrin chromophore with sterically hindered groups via condensation reaction and hydrolysis, affording the formyl Fc-ZnP dyads. These dyads further underwent Prato reaction in a combination of C60 and sarcosine to yield the desired Fc-ZnP-C60 triads, TC1, TC2, and TC4. All the new compounds were characterized by standard methods including 1H NMR (Figures S2−S4), IR spectroscopies, and high-resolution mass spectrometry. The reference molecules, Fc-ZnP-C60-ref,22 TC1-ref,19 and ZnPref,18 were also prepared following literature reports. The UV−visible absorption spectra of TC1, TC2, and TC4 were recorded in DMSO and DMSO/H2O (1/99, v/v). As shown in Figure 2 (see also Table S1), broadening and peakshift of the absorption in DMSO/H2O relative to those in DMSO reflect the formation of their nanoaggregates with an increase of solvent polarity.19,33,34 The strong fluorescence quenching of TC1, TC2, and TC4 in DMSO/H2O in comparison with DMSO (Figure S5) can be ascribed to selfquenching of their singlet excited-states caused by intense aggregation among the ZnP and/or C60 moieties in the highly polar aqueous solvent, intermolecular quenching associated with intermolecular PET in the aggregates, and accelerated intramolecular quenching related to faster intramolecular PET by the change in the solvent polarity.22,35,36 The molar absorption coefficient of the Soret band increases in the order TC1 < TC2 < TC4 in DMSO and in DMSO/H2O D

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Figure 4. Nanosecond time-resolved TA spectra of (A) TC1 at 0.46 μs, (C) TC2 at 0.56 μs, (E) TC4 at 0.46 μs, and (G) TC1-ref at 0.34 μs. The decay profiles (black), biexponential fitting curves (red), and the fitting residuals (blue) of (B) TC1, (D) TC2, (F) TC4, and (H) TC1-ref at 1000 nm arising from C60•− in DMSO/H2O (λex= 550 nm).

TC1, TC2, TC4, and TC1-ref in DMSO/H2O consist of two long-lived components, i.e., the fast CR derived from the monomeric state and the slow CR derived from the nanoaggregate state.21,23,39,40 The pre-exponential factors (A1 and A2) would reflect the relative amplitudes of the two states, and those of TC1, TC2, and TC4 in DMSO/H2O showed values similar to those in liposome, respectively. The CS yields of TC1, TC2, and TC4 are virtually the same (50−51%), and the CR rate constants and the amplitude ratios are largely similar. The CS yield of TC1-ref being higher than that of TC1 indicates that suppression of the local aggregation around the porphyrin units by the hydrophobic long alkyl chains is of importance to achieve the high CS yield in the aqueous solvent (vide supra). Unexpectedly, the number of the cationic moieties has little impact on the CS yields and CR rate constants. The local aggregation would not be affected by the electrostatic interaction (i.e., the cationic moieties) introduced at the distal bridges between the porphyrin and the ferrocene as well as the porphyrin and the C60 units in the highly polar aqueous solvent. To get insight into the behavior of the Fc−ZnP−C60 triad molecules in the biological cell membrane, they were examined in an artificial lipid bilayer system (COATSOME EL-11-A, see the Experimental Section). First, the UV−visible absorption spectra were recorded to clarify the influence of the incorporation into the liposome (Figure S9, Table S1). The absorption spectra of TC1, TC2, and TC4 in the liposome

Figure 3. Size distribution of the nanoaggregates of (A) TC1, (B) TC2, and (C) TC4 on a mica substrate. The size was determined by the AFM image analysis on Figure S6. The size distribution of TC4 was obtained by using the diluted spin-coating solution (0.5 μM) on the mica substrate.

scopic measurements is unlikely to be reliable.21 Because the formation of the Fc+ cation is difficult to monitor, the estimation of the charge-shift yield from Fc to ZnP•+ requires the subtraction from the decay rate constant of ZnP•+ to the CR rate constant in the corresponding ZnP−C60 reference dyad. Because the aggregation behavior of the dyad is anticipated to be significantly different from that of the Fc− ZnP−C60 triad as a consequence of the different molecular structures, the plausible difference in the CR rate constants of the dyad and triad would cause a large error in the estimation. Moreover, the complex kinetics of multicomponents arising from the different states of the aggregates in the present environment may make the accurate analysis and interpretation impossible. Unlike Fc−ZnP−C60-ref showing a single exponential decay at 1000 nm in benzonitrile (Figure S8), the decay profiles of E

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corroborating the formation of the same long-lived chargeseparated state (i.e., Fc+−ZnP−C60•−). The decay profiles consisting of the fast and slow components at 1000 nm in the liposome are also similar to those in DMSO/H2O. The CS yields in the liposome exhibit remarkable dependence on the number of the cationic moieties in the triads. With an increase in the number of the cationic moieties, the CS yield is increased dramatically, reaching the highest value of 86 ± 4% in TC4. The highest value is much higher than that of 27 ± 1% in TC1ref.19 As supported by the absorbance change shown in Figure S9, the large aggregates of TC1 and TC2 would cause unfavorable cancellation of the charge-separated states by intermolecular ET and quenching of the porphyrin singlet excited-state.19 Moreover, the four cationic moieties of TC4 would be beneficial to increase the dispersibility in the lipid membrane by the electrostatic interaction between the cationic moieties and the anioic membrane surface, suppressing unfavorable aggregation, as seen in the microscopic image in Figure 6 (vide infra).

maintain the characteristic absorption peaks of the ZnP moieties in DMSO/H2O, whereas observable variation in the absorption decreases (TC1 > TC2 > TC4) as a result of their different aggregation behavior in the cell membrane-like environment.33,34 The large decrease in the absorbance and the molar absorption coefficient during the incorporation indicates the formation of their large aggregates.19 Thus, this trend reflects the aggregate size in the lipid bilayer (TC1 > TC2 > TC4). It is noticeable that the steady-state fluorescence of the porphyrin moiety of TC4 in the liposome is remarkably enhanced relative to that in DMSO/H2O, whereas those of TC1 and TC2 in the liposome are slightly increased (Figure S10). These striking contrasts of TC4 with TC1 or TC2 imply the remarkable suppression of the unfavorable aggregation behavior of TC4 in the biological bilayer membrane by the introduction of multicationic moieties into the triad core, in particular the self-quenching of the singlet excited state. The introduction of multicationic moieties would be preferable for the use of the excited state toward potential biological applications. The nanosecond time-resolved TA spectra of TC1, TC2, TC4, and TC1-ref in the liposome were recorded for assessing the CS properties (Figure 5, Table 1). The TA spectra in the liposome are virtually identical to those in DMSO/H2O,

Figure 6. Confocal microscopy images of PC12 cells treated with WGA-Alexa Flour 488 conjugate and (A) TC1, (B) TC2, and (C) TC4.

These results exemplify that the introduction of the multiple cationic moieties into the Fc−ZnP−C60 molecule is an effective strategy to attain the high CS yield in the cell membrane-like environment, keeping the long lifetime of the charge-separated state. It should be emphasized that the CS yield in the lipid bilayer is the highest value ever reported for those using noncovalent or covalent D−A systems including the highest CS yield (67%) of the tosylate/porphyrin/cyanine in lipid bilayers (DOPC/DOPA, 9/1, w/w) reported by O’Brien et al.11 The finding on the highest CS yield in the lipid bilayer encouraged us to further examine the effects of the cationic moieties in the Fc−ZnP−C60 molecule on the depolarization of the membrane potential in living cells. The amount of the incorporated triad molecules per a single PC12 cell was evaluated via spectrophotometric determination.18,19 There is a linear correlation between the amount of the incorporated triad molecules and the initial concentration of the triad solution used for the loading (Figure S11). It is noteworthy that the amount of the incorporated triad molecules per the single cell

Figure 5. Nanosecond time-resolved TA spectra of (A) TC1 at 0.46 μs, (C) TC2 at 0.64 μs, (E) TC4 at 0.60 μs, and (G) TC1-ref at 0.46 μs in the liposome (λex= 550 nm). The decay profiles (black), biexponential fitting curves (red), and the fitting residuals (blue) of (B) TC1, (D) TC2, (F) TC4, and (H) TC1-ref at 1000 nm arising from C60•− in the liposome (λex= 550 nm). F

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of TC1 (+7 mV), TC2 (+4 mV), and even TC1-ref (+4 mV) (Figure 7). The observed excellent depolarization performance of TC4 agrees with the highest CS yield of TC4 in the liposome together with the most homogeneous distribution of TC4 in the cell membrane. Consequently, it was concluded that well-balanced hydrophobicity and hydrophilicity in the D− A linked molecules lead to the highest CS yield in the artificial bilayer, which is consistent with the largest depolarization caused by the charge-separated state in the biological bilayer.

increases with increasing the number of the cationic moieties (e.g., TC1: (1.9 ± 0.0) × 106 < TC2: (3.2 ± 0.1) × 106 < TC4: (3.4 ± 0.1) × 106 at the initial concentration of 0.5 μM).41 This is consistent with the intrinsic nature of living cells with the negatively charged outer cell membrane. Moreover, confocal microscopic observation directly visualizes the incorporation of TC1, TC2, and TC4 into the cell membrane (Figure 6). This difference may be caused by the different size of the nanoaggregates as observed in the AFM measurements as mentioned above. Although the fluorescence of TC1, TC2, and TC4 is very weak, it is detectable by a highly sensitive GaAsP detector. The confocal microscopic observation directly visualized the incorporation of TC1, TC2, and TC4 into the cell membrane. Both TC1 and TC2 are localized, whereas TC4 is rather homogeneously distributed in the cell membrane. These behaviors of TC4 were expected to lead to a large photoresponse in the light induced depolarization of cell membrane potentials. Detection of singlet oxygen (1O2) by SOSG and apoptosis−necrosis assay using Annexin V-FITC/PI dyes was conducted under the green light illumination (545− 555 nm). These assays ensure that any optical responses of PC12 cells are not caused by the photocytotoxicity of the molecules of interest.42 Detection of singlet oxygen (1O2) showed that all the triad molecules do not exhibit significant phototoxicity (Figure S12A). The apoptosis−necrosis assays also did not reveal any considerable induction of cell death by the illumination (Figure S12B). The low photocytotoxicity is in marked contrast to the severe cell damage caused by the positive control, ZnP-ref, under the same conditions. The importance and effectiveness of the high CS yield in the cell membrane were demonstrated in photoinduced change of the cell membrane potential under green light illumination (Figure 7). The amounts of the incorporated different triad molecules in the PC12 cells were adjusted to be identical (1.9 × 106 molecules/cell) (Figure S11). As illustrated in Figure 7A− D, all the triads, TC1, TC2, TC4, and TC1-ref, induced the depolarization in the cell membrane potential under the green light illumination. Remarkably, TC4 afforded the largest depolarization (+16 mV), which was much larger than those

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CONCLUSIONS In this study, a series of ferrocene−porphyrin−fullerene linked triads possessing different numbers of the ammonium moieties were synthesized to explore their structure−function relationship under biological environments. With an increase of the hydrophilic cationic moieties, the size of their nanoaggregates was decreased, and the aggregate dispersibility in aqueous solutions was improved. In particular, TC4 with the largest number (i.e., four) of cationic moieties exhibited the highest CS yield (86%) ever reported for donor−acceptor systems in artificial and biological lipid bilayers, the most homogeneous distribution in the cell membrane, and the largest photoinduced depolarization among all the donor−acceptor molecules. Considering the similarity in the results of the liposome and the cell membrane, the present methodology would be universal to modulate the CS yield in living cell membranes. Accordingly, further exploration will be possible by altering the number and the position of the cationic moieties in D−A linked molecules, realizing efficient photocontrol and artificial redox reactions in cellular systems.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpcc.7b04466. Synthesis of the triads; Figures S1−S12 (PDF)

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. ORCID

Tatsuya Murakami: 0000-0001-5199-500X Hiroshi Imahori: 0000-0003-3506-5608 Author Contributions ¶

N.C. and Y.T. contributed equally.

Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work is supported by Grants-in-Aid for Scientific Research (S) (JP25220801) and (C) (JP15K05563), MEXT, Japan. The iCeMS is supported by the World Premier International Research Center Initiative. We are grateful to the CeMI for microscopic measurements.

Figure 7. Representative photoinduced depolarization traces of PC12 cells incorporated with (A) TC1, (B) TC2, (C) TC4, and (D) TC1ref and (E) their amplitudes of the photoinduced depolarization. Conditions: Xe lamp wavelength, 545−555 nm; input intensity, 7 mW cm−2; illumination time, 8 min. The error bars depict SD. Statistically significant differences between the negative control and each compound are illustrated with asterisks (***P < 0.001).

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