Light-Harvesting Nanorods Based on Pheophorbide-Appending

Article pubs.acs.org/Biomac

Light-Harvesting Nanorods Based on Pheophorbide-Appending Cellulose Keita Sakakibara,*,†,‡,§ Mari Granström,∥,⊥ Ilkka Kilpelaï nen,*,∥ Juho Helaja,∥ Santtu Heinilehto,■ Rintaro Inoue,‡ Toshiji Kanaya,‡ Jonathan P. Hill,†,§ Fumiaki Nakatsubo,¶ Yoshinobu Tsujii,‡,§ and Katsuhiko Ariga*,†,§ †

World Premier International (WPI) Center for Materials Nanoarchitectonics (MANA), National Institute for Materials Science (NIMS), 1-1 Namiki, Tsukuba 305-0044, Japan ‡ Institute for Chemical Research (ICR), Kyoto University, Gokasho, Uji, Kyoto 611-0011, Japan § JST, CREST, 1-1 Namiki, Tsukuba 305-0044, Japan ∥ Department of Chemistry, University of Helsinki, P.O. Box 55, FI-00014, Finland ⊥ BASF SE, GCN/R - M311, 67056 Ludwigshafen, Germany ¶ Research Institute for Sustainable Humanosphere (RISH), Kyoto University, Gokasho, Uji, Kyoto 611-0011, Japan ■ Center of Microscopy and Nanotechnology, University of Oulu, Erkki Koiso-Kanttilan katu 3, FI-90570 Oulu, Finland S Supporting Information *

ABSTRACT: In contrast to the success in artificial DNA- and peptide-based nanostructures, the ability of polysaccharides to self-assemble into one-, two-, and three-dimensional nanostructures are limited. Here, we describe a strategy for designing and fabricating nanorods using a regioselectively functionalized cellulose derivative at the air−water interface in a stepwise manner. A semisynthetic chlorophyll derivative, pyro-pheophorbide a, was partially introduced into the C-6 position of the cellulose backbone for the design of materials with specific optical properties. Remarkably, controlled formation of cellulose nanorods can be achieved, producing lightharvesting nanorods that display a larger bathochromic shift than their solution counterparts. The results presented here demonstrate that the self-assembly of functionalized polysaccharides on surfaces could lead the nanostructures mimicking the naturally occurring chloroplasts.

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INTRODUCTION Recent advances in both peptide/DNA syntheses and supramolecular chemistry have triggered new interest in construction of biopolymer-based nanostructures with control over dimensions on the nanoscale, or alternatively nanofabrication, which offers potential for realizing bottom-up nanotechnology through low energy and inexpensive spontaneous processes.1,2 The driving forces of the self-assembly for peptides and DNAs are based on intermolecular interactions, mostly mediated by planar hydrogen bonding between CO and N−H amide groups in folding of small peptide segments or complementary hydrogen bonding among purine and pyrimidine residues in DNA double helices. On the other hand, the third class of biopolymers, polysaccharides, may evolve as another type of molecular building blocks but require novel approaches. One of the prominent features of polysaccharides is the multiplicity of reactive hydroxyl groups that can be covalently functionalized without destroying their inherent regularity and chirality. Only a few nanofabrication methods have been applied to polysaccharides. On this point, most reports highlight solution-based strategies to prepare nanoparticles3 or inclusion © 2013 American Chemical Society

complexes of synthetic polymers with helical polysaccharides including amylose, curdlan, and their branched polymers (starch and schizophyllan).4−7 Recently, Kim and co-workers reported a supramolecular device for artificial photosynthetic mimics based on organized carbohydrate-dye assemblies.8 However, two-dimensional structural systems, especially on surfaces or at interfaces, accurately constructed at the molecular level are far less common, mainly because it is difficult to manipulate hydrogen bonding interaction among monosaccharide building units. We have considered regioselective functionalization of native cellulose, a linear semiflexible polysaccharide consisting of regio- and stereospecific β-1,4-glycosidic linked D-glucose units, as a fundamental building block to obtain advanced cellulosic materials with a novel functionality.9,10 Hitherto, we attached a porphyrin group to some of the primary hydroxyl groups of cellulose selectively at the C-6 position, affording organized Received: June 11, 2013 Revised: July 16, 2013 Published: July 18, 2013 3223

dx.doi.org/10.1021/bm400858v | Biomacromolecules 2013, 14, 3223−3230

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Figure 1. Structure of the regioselectively chlorophyll-appended cellulose acetate (1) and an illustration of the assembly process.

Scheme 1. Synthesis of 1a

Reagents and conditions: (a) trityl chloride, pyridine, [amim]Cl, 60 °C, 6 h, 98%; (b) acetic anhydride, pyridine, [amim]Cl, rt, 24 h, 83%; (c) HCl, THF, rt, 3 days, 96%; (d) pyro-pheophorbide a, DCC, DMAP, DMF, r.t., 24 h, 79%.

a

pheophorbide-appended cellulose acetate (2,3-O-diacetyl-6-Opheophorbide-cellulose 1; Figure 1), wherein an air−water interface is used to confine hydrogen bonding network to twodimension. Recent studies have illustrated that the Langmuir− Blodgett (LB) technique is an effective strategy for producing various nanoarchitectures.11 In this paper, we show that the cellulose rod-shaped nanostructures have optical properties as well as photocurrent generation properties, which can become artificial light collectors as similar to the photosynthetic systems consisting of chlorophyll a-protein complexes in chloroplasts of higher plants in nature.

multichromophore arrays for the efficient photocurrent generation thin films.9 In addition, the acyl groups at the secondary hydroxyl groups (i.e., C-2 and C-3) act as solubilizing groups, which are relevant not only for processability but also for insulating hydroxyl groups. The regioselective functionalization avoids the establishment of multiple intermolecular hydrogen bonds,3c though not controlling the direction of hydrogen bonding precisely. We therefore asked whether it is possible to control the hydrogen bond network by view of confined media, such as an air−water interface, where molecules are restricted in a certain space and obstructed from the development of random intermolecular interactions. Herein, we report the fabrication of light-harvesting polysaccharide-based nanorods, based on a regioselectively

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EXPERIMENTAL SECTION

Materials. All reagents and solvents were of the highest grade available from Aldrich, Fluka, and J. T. Baker and used without further 3224


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purification if not otherwise stated. Microcrystalline cellulose (Aldrich) was dried in a vacuum oven at 60 °C for 24 h prior to use. Water used for the LB subphase was distilled using an Autostill WG220 (Yamato) and deionized using a Milli-Q Lab (Millipore). Its specific resistance was greater than 18 MΩ cm. Spectroscopic grade chloroform (Kanto Chemicals) was used as the spreading solvent. All reactions were carried out under argon atmosphere. The ionic liquid, 1-allyl-3-methylimidazolium chloride ([amim]Cl), was synthesized according to Zhang et al.12 with slight modification to their procedure; both allyl chloride (Aldrich) and 3-methylimidazole (Aldrich) were distilled prior to use. [Amim]Cl was further purified, to remove trace color, by dissolving the crude [amim]Cl mixture in water and refluxing with activated charcoal (18 h). The solution was filtered through Celite plug and the water was removed by distillation and dried for 2 days in vacuo to yield pure [amim]Cl as a pale yellow crystalline solid with a melting point of 52 °C. The chlorophyll, pyropheophorbide a, was extracted from green leaves and modified according to references.13 Synthetic route for compound 1 is shown in Scheme 1. Synthesis of (2,3-O-Diacetyl-6-O-pheophorbide)cellulose (1). 6-O-Tritylcellulose (DS = 0.90)14 (2.0 g, 5.3 mmol per AGU) was dissolved in [amim]Cl and acetic acid anhydride (2 mL, excess) and pyridine (2 mL, excess) were added. The reaction mixture was stirred at rt for 24 h. The mixture was poured into water and the formed precipitate was filtered off, washed with plenty of water (3 × 50 mL water). The precipitate was dissolved in DMSO from which it was regenerated with water to give 2,3-O-diacetyl-6-O-tritylcellulose (2.1 g, 83%, DSAc = 1.92; DStotal = 2.92). 13C NMR (125 MHz, DMSO-d6): δ = 21 (Ac), 62.5 (C-6), 70−73 (C-2,3,5), 76 (C-4), 86.2 (C-7), 101 (C-1), 112.7 (C-12), 126.4 (C-10), 127.3 (C-11), 134.2 (C-9), 138.4 (C-8), 159.6 (C-13), 171 (CO). CP-MAS NMR (300 MHz): δ = 21 (Ac), 62 (C-6), 73 (C-2,3,5), 83 (C-4), 101 (C-1), 127 (Tr), 148 (Tr), 170 (CO). Then, 2,3-O-diacetyl-6-O-tritylcellulose (1.0 g, 2.1 mmol) was dissolved in THF (50 mL) and conc. HCl (1.5 mL) was added. The reaction was stirred at rt for 3 days, after which the product was regenerated from water to give 2,3-O-diacetylcellulose (0.5 g, 96%, DSAc = 1.92). 13C NMR (125 MHz, DMSO-d6): δ = 17 (Ac), 59 (C-6), 69 (C-2), 70 (C-3), 72 (C-5), 78 (C-4), 99 (C-1), 171 (CO). pyro-Pheophorbide a (30 mg, 0.05 mol) was dissolved in DMF (10 mL) and N,N′-dicyclohexylcarbodiimide (DCC) (7 mg, 0.05 mmol) and N,N′-dimethylaminopyridine (DMAP) (10 mg, 0.05 mmol) were added. The reaction was stirred at rt for 2 h, after which it was added to the solution of 2,3-O-diacetylcellulose (10 mg, 0.04 mmol) in DMF (5 mL). The reaction was stirred at rt for 24 h. The product was regenerated with water and washed with MeOH to give 1 as a black solid (9 mg, 79%, DSchlorophyll = 0.07). 1H NMR (500 MHz, CDCl3): δ = 1.4 (H-82), 1.8 (H-181), 2.0 (OAc), 2.8/2.9 (H-171), 3.3 (H-71), 3.4 (H-5′), 3.7 (H-121), 3.9 (H-81), 4.1 (H-6), 4.2 (H-61), 4.5 (H-1), 4.6 (H-2), 5.3 (H-132), 5.4 (H-3), 6.2 (H-32), 8.0 (H-31), 8.6 (H-20), 9.4 (H-5), 9.6 (H-10). IR (cm−1): 3394 (OH), 3330 (NH), 3110 (aromatic), 1690 (CO), 1466 (C−O), 650 (aromatic). UV− vis (λmax, nm): 416, 669. Preparation of Cellulose Nanorods. The surface pressure versus area (π−A) isotherm measurement and thin-film preparation were carried out using an FSD-300 computer-controlled film balance (USI System, Fukuoka). Diluted solutions of the cellulose derivative (0.05 wt %) in chloroform was prepared for the spreading solution. The water subphase temperature was kept constant at 20 °C by circulating thermostatted water system. Fluctuation of the subphase temperature was within ±0.2 °C. The surface pressure was measured by the Wilhelmy method. The diluted solution was spread onto a water subphase in a Teflon-coated Langmuir trough. The solvent was allowed to evaporate off for 15 min and the π−A isotherm was measured at a constant compression rate of 0.2 mm s−1. The monolayer formed at the air−water interface was transferred onto a substrate by the vertical dipping method with upstroke motion of 0.01 mm sec−1. The substrate was immersed before spreading the diluted cellulose solutions onto the water subphase. A quartz plate (for UV and fluorescence), freshly cleaved mica (for AFM), silicon wafer (for

GIWAXS), and an ITO electrode (for photocurrent measurements) were employed as substrates for the monolayer deposition. Measurements. NMR spectra were recorded on Varian Inova 500 MHz NMR spectrometer in DMSO-d6 or CDCl3 as the solvent if not otherwise stated. Chemical shifts, relative to tetramethylsilane (TMS) as an internal standard, are given in δ values, and coupling constants in hertz. 13C resonances were assigned by means of APT, HMQC, and HMBC spectra. AFM images were obtained with a commercial AFM unit (SPA400-SPI4000, Seiko Instruments Inc., Chiba, Japan). All AFM images were taken in a dynamic force mode (DFM, i.e., tapping mode) at optimal force. Ultraviolet−visible (UV−vis) spectra were recorded on a Shimadzu UV-3600 spectrophotometer, and fluorescence spectra was recorded on a JASCO FP-6500 spectrofluorometer. In-plane and out-of plane grazing incidence wide-angle X-ray scattering (GIWAXS) measurements were performed on the BL03XU instrument15 installed at a synchrotron (SR) X-ray scattering facility, SPring-8, in Nishiharima, Japan and used the scintillation counter as a detector. The incident wavelength and incident angle of X-ray were 1.00 Å and 0.1°, respectively and the scattering angle (2θ) ranged from 5 to 30°. The acquisition time at one point was 800 ms to avoid unwanted damage to sample. Under the above setup, we can cover a q range from 0.55 to 3.25 Å−1, where q is the magnitude of the scattering vector and is defined by

q=

4π sin(θ) λ

(1)

where 2θ and λ are the scattering angle and wavelength of the incident beam, respectively. X-ray photoelectron spectroscopy (XPS) was used to study the presence of metal impurities at the surface of the samples. Measurements were carried out by using ESCALAB 250Xi spectrometer (Thermo Fisher Scientific) with monochromatic Al Kα (1486.6 eV) X-ray radiation and 900 μm analysis area. Wide energy range survey spectra were acquired at detector pass energy of 150 eV. Narrow energy range scans acquired at 40 eV pass energy were collected individually from Al, Ag, Cd, Co, Cu, Fe, Ga, Hg, In, Ni, Mg, Mn, Pb, Ru, Sn, Ti, and Zn metals to determine their presence more accurately. Measured XPS survey and narrow scan spectra showed that there are no metal impurities at the surface of the samples. The photocurrent measurements were carried out at a constant bias potential using an electrochemical analyzer (ALS650B, BAS) at room temperature (approximately 25 °C) on light irradiation. A 500 W xenon arc short lamp (UXL-500SX, Ushio, Japan) was used as a light source, equipped with a UV and IR cutoff filter (SCF, Ushio, Japan). The monochromatic light was obtained from a xenon lamp filtered with metal interference filters MIF-W type (400−550 nm wavelength, 10 nm fwhm, Vacuum Optics Corporation of Japan). The light intensity at the irradiation substrate surface was measured with a thermopile (MIR-100C, TAZMO, Japan). The LB film on an ITO electrode as a working electrode was contacted with an aqueous solution containing 0.1 M Na2SO4 as an electrolyte and 50 mM hydroquinone (H2Q) as a sacrifice electron donor. An electrode area of 1.54 cm2 was exposed to the electrolyte solution. A saturated calomel electrode (SCE) as a reference electrode and a platinum wire electrode as a counter electrode were used. The electrolyte solution was initially purged with nitrogen for 15 min and then maintained under a flow of nitrogen. The photocurrent quantum yield (Φ) based on the number of photons adsorbed by the chlorophyll moiety in the LB film was calculated according to the following equation i

Φ=

100 e [I(1 − 10−A)]

,

I=

Wλ hc

(2)

where i is the observed photocurrent density, e is the charge of the electron, I is the number of photons per unit area and unit time, A is the absorbance of the monolayer at λ nm, λ is the wavelength of light illumination, W is light power irradiation at λ nm, h is the Planck constant, and c is the light velocity. 3225


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bonding between C-6 hydroxyl groups and the π−π stacking of the chlorin moieties would contribute to the stable formation of the floating monolayer. Next, the monolayer was transferred onto freshly cleaved mica at a surface pressure of 10 mN m−1 by the vertical dipping method. Figure 3 shows representative tapping mode atomic force microscopy (AFM) image of the obtained monolayer film,

RESULTS AND DISCUSSION Formation of Cellulose Nanorods. We selected pyropheophorbide a at the C-6 position of cellulose as a chlorin moiety and acetyl groups at C-2 and C-3 positions as solubilizing moieties. The pheophorbides are metal-free chlorophyll derivatives that are present only in low abundance in natural photosynthetic systems, yet having similar light absorption and emission properties as chlorophylls.16 As depicted in Scheme 1, the target compound 1 was produced via four steps in a 62% overall yield, starting from microcrystalline cellulose according to the homogeneous protection group strategy combined with ionic liquid (1-allyl3-methylimidazolium chloride, [amim]Cl), as reported previously.10 Briefly, the synthesis of 1 was started from regioselective protecting of the C-6 hydroxyl group by a trityl moiety in the reaction between MCC and trityl chloride in the presence of pyridine in [amim]Cl. Then, the obtained 6-Otritylcellulose was acetylated using the common procedure employing acetic anhydride with pyridine in [amim]Cl, yielding 2,3-O-diacetyl-6-O-tritylcellulose with a total degree of substitution (DS) of 2.92 in a 83% yield. Subsequent deprotection of the trityl moiety under acidic condition in THF and the following esterification of chlorophyll (i.e., pyropheophorbide a) using the activation approach by DCC and DMAP gave the target 1 with a DS of 0.07 for the chlorophyll substituents at the C-6 position and a DS of 1.92 for the acetyl groups at the C-2 and C-3 positions. The DS value of the chlorin functionalities was found to be restricted to low values presumably because of the steric hindrance of the pheophorbide moiety in the course of the reaction. As a result, the hydroxyl groups remaining at the C-6 position (DS6(OH) = 0.93) serve as a main hydrogen bond donor and acceptor along the polymer backbone. To prepare a floating Langmuir monolayer, the chloroform solution of 1 was first spread onto a water surface. The surface pressure versus molecular area (π−A) isotherm (Figure 2)

Figure 3. Representative AFM image of the LB monolayer film deposited onto freshly cleaved mica at π = 10 mN m−1 and the height profile along the black line.

showing nanorod structures on surfaces. They tend to pack together over an area of 5 μm2 (Supporting Information, Figure S1). These nanorod structures were observed above the surface pressure of 10 mN m−1, as evidenced by the AFM images of monolayers prepared at 5 and 20 mN m−1 (Supporting Information, Figure S2), suggesting that the nanostructures were formed during compression. These nanostructures were also observed on silicon wafer. It should be noted that this structure was not obtained when the solution of compound 1 was drop-cast onto the substrate (Supporting Information, Figure S3), nor with cellulose triacetate. The cross-sectional diagram (Figure 3) shows size of the individual nanorod-shaped structure: ca. 0.4−0.5 nm in height with the width of 180 nm and the length of around 500 nm. The height corresponds to the cross-sectional thickness of a glucose ring, so that the glucose units would lie nearly flat on the substrate surface.17 Apparently, the height can be confined to be monomolecular level owing to the nature of interfacial confined space. We then employed grazing-incidence wide-angle X-ray scattering (GIWAXS) in order to understand the aggregation state in the LB film. Figure 4 shows the one-dimensional GIWAXS profiles (λ = 1.00 Å, rt) of the 49-layer LB films transferred onto a silicon wafer at a surface pressure of 10 mN m−1. The out-of-plane GIWAXS pattern revealed the existence of a broad scattering at 4.6 Å (q = 0.84 Å−1) that could coincide with the interlayer spacing. This value is in agreement with the height estimated by AFM (Figure 3). By contrast, the in-plane GIWAXS pattern shows two sharp peaks at 7.44 Å (q = 0.84 Å−1) and 3.57 Å (q = 1.76 Å−1) with a broad band at 5.2 Å (q = 1.22 Å−1), suggesting that the rod-shaped nanostructures have a certain degree of order. In order to rule out the possibility that some metal contaminations could produce the observed

Figure 2. Surface pressure versus area (π−A) isotherm for 1 at the air−water interface at 20 °C.

shows a well-condensed phase with a collapse pressure of ∼50 mN m−1 and a limiting area of 0.59 nm2 per anhydroglucose unit (AGU), which corresponds to that of cellulose triacetate (0.54 nm2 per AGU).17 Thus, the limiting area is almost equal to the cross-sectional area of the glucopyranose ring. The collapse pressure was higher than that of cellulose triacetate (below 30 mN m−1), suggesting that both the hydrogen 3226


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chlorines. The absorption edge is extended close to 750 nm. These bathochromic shifts might be attributed to the J-type excitonic coupling between the S0−S1 transition dipole moments of the chlorin Qy bands.18 In fact, the chlorin Qyband shift correlates rather well with the values that have been reported for pheophorbide ester aggregates in ethanol (695 nm) and water−acetone mixtures (698 nm).19 It should be noted that there is indeed some contribution of monomers, dimers, some other oligomeric assemblies, and possibly even some amount of H-aggregates. Nevertheless, qualitatively the spectra in Figure 5 suggest that there is dominating amount of J-aggregates that causes bathochromic shifts at the Qy and Soret band. This behavior is qualitatively similar to that found for chlorosomal chlorophyll J-aggregates.20 In our systems, however, the shifts are less pronounced and not as uniform. Since the chlorosomal chlorophylls are more precisely packed by metal-hydroxyl-carbonyl bond network,21 these defined coordinative bond interactions explain well the highly uniform chlorosome self-assemblies.22 In this comparison, our system has more obscure driving force for the assembly, though there are evidently clear signs of J-aggregation appeared in Figure 5. This is indeed surprising when the cellulose scaffold could be expected to dictate predominantly the chlorin orientations, yet the chromophores organize with each other to notable degree of order. Next stage in our studies will be to attach chlorosomal-like metalated chlorins in cellulose to study in more detail the nature of assembly. It has been demonstrated in NMR studies that the pheophorbides have a tendency to form head-to-head aggregates, that is, ester and ketone groups of chlorin ring pointing on opposite directions in stacking chlorins, in acetone and chloroform.19b,23 The enthalpy of aggregation has been explained with an electrostatic model, in which aggregation arises from repulsive π−π and attractive π−σ interactions between aromatic chlorin macrocycles.24 Additionally, for hydrophobic porphyrins, it has been concluded that solvophobic effects plays an essential role in aggregate formation in polar media, indicating that entropic factor has a contribution in the molecular organization.25 Taking this into all account, the cellulose derivative may be in principle favored to form aggregates by both enthalpy and entropy aspects. Nonetheless, the evident appearance of the aggregates was unexpected, while the DS of 0.07 denotes that in average only every fourteenth AGU is equipped with the chlorin moiety. In turn, this implies wide spacing between the pheophorbides, which makes the chlorin−chlorin macrocycle interaction improbable within the single cellulose chain. In contrast to this, in the LB monolayer film, the head-to-head chlorin ring interactions could take place between the pheophorbides that are attached on neighboring cellulose chains (Figure 1). To this end, it may be well that the hydrophobic chlorin−chlorin interactions is an inherent and driving force for the selfassembly of the cellulose nanostructures at the air−water interface. Another important feature of the absorption spectra is the difference in the relative intensities at the Soret and Qy absorption maxima (ISoret/IQy),26 which are 2.38, 2.78, and 1.11 for those in solution, drop-cast film, and LB film, respectively. It is said that the ratio ISoret/IQy of chloroplasts in higher plants are from 1.22 to 1.35.27 Thus, the pheophorbide-appended cellulose aggregates have spectral characteristics closer to those of the naturally occurring chloroplasts, whereas compound 1 in solution and in drop-

Figure 4. In-plane (red) and out-of-plane (blue) one-dimensional GIWAXS profiles of the 49-layer LB films transferred onto a silicon wafer substrate. The in-plane profile has been shifted vertically and 3fold magnified.

patterns, XPS measurements of surfaces were run to confirm that this was not the case. In consequence, it seems reasonable to say that the rod-shaped structure that appeared in the AFM image (Figure 3) is not a patchy monolayer stripe but a selfassembled nanostructure. It should be emphasized that to the best of our knowledge this is the first report on the nanorod structures from polysaccharides and their derivatives. Optical and Photoelectrochemical Properties. UV−vis spectroscopy is an efficient tool in investigation of molecular interactions between porphyrin molecules. Figure 5 shows the

Figure 5. Normalized UV−vis absorption of compound 1 in chloroform solution (13.1 μM per AGU) (black broken curve) in the drop-cast film (blue curve), and the LB film (red curve) deposited at π = 10 mN m−1.

comparison of the UV−vis absorption spectra of compound 1 in the LB monolayer film (red curve) with the randomly distributed state in chloroform solution (black broken curve) and in the drop-cast thin film (blue curve). Figure 5 clearly shows differences in the absorption spectra. The spectrum of compound 1 in the solution displays the intense Soret band at 416 nm and four Q-bands at 510, 540, 611, and 669 nm, which are characteristics for the corresponding pheophorbide monomer in chloroform.13b In the case of the drop-cast thin film, there is little change in the spectrum where the Soret band at 414 nm and one detectable slightly red-shifted Qy-band at 675 nm are observed. In contrast, the spectrum of the LB monolayer film exhibits the remarkable red-shifted absorption peaks at 441 and 704 nm, descriptive of the aggregation of 3227


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fluorescence result suggests the existence of intermolecularly interacting chlorin-moieties attached to the cellulose backbone. One potential application of the pheophorbide-appended cellulose nanorods lies in light-harvesting photocurrent generation thin film systems. We prepared the monolayer film of 1 on an ITO electrode as a photoanode in an electrochemical cell. Figure 7 shows a steady-state anodic photocurrent appeared during light irradiation around the Soret (blue plot) and Q-band (red plot), demonstrating a good on/ off photoresponse (ca. 90 and 50 nA cm−2 upon irradiation at 420 and 690 nm, respectively, in the presence of hydroquinone (H2Q) under 0 mV bias). The pheophorbide-based film has wide-range photoresponse up to 760 nm (Figure 7B). The film was relatively stable under light illumination (Supporting Information, Figure S4), even though the natural pigment was used. The photocurrent quantum efficiency, calculated by eq 2, was 0.56 and 0.35% at 420 and 690 nm, respectively, which were similar to those of the LB film prepared at 20 mN m−1 and those of the spin-coated film (Supporting Information, Figure S5 and Table S1). This is presumably because the cellulose nanorod LB film in the present study has some of the energy traps, so that the photocurrent property of the nanorods was almost comparable to that of the less-ordered spin-casted film. Despite the low efficiency as compared to the other artificial photosynthetic systems,9,29 appropriate variation of the donor−acceptor systems, especially increasing the chlorin DS, as well as film structure, could improve the device performance.

cast films has spectral properties much closer to the randomly distributed chlorin in solutions. For many applications including photocurrent generation systems, a red shift and intensification of the Qy transition are desirable, as discussed below. Next, the steady-state fluorescence spectroscopy was conducted to observe the interaction of the chlorin chromophores in the films (Figure 6). The fluorescent

Figure 6. Fluorescence spectra of compound 1 in chloroform solution (13.1 μM per AGU) (black broken curve) in the drop-cast film (blue curve), and the LB film (red curve) deposited at π = 10 mN m−1.

spectrum of compound 1 in the solution, excited at the wavelength of the most intense Soret band, exhibits the emission peak of the chlorin moiety around 678 nm with a shoulder around 720 nm. The band at 678 nm is assigned to the monomer fluorescence whereas that at 710 - 720 nm can be assumed to derive from an aggregate with higher number of molecules included or a different dimer conformation than the one which forms excimers.28 Because the pheophorbideappended cellulose was well diluted in chloroform for the fluorescence measurement, the band at 720 nm is thought to be derived from the intramolecular interaction between pheophorbide moieties attached along the cellulose backbone. The monomer fluorescence at 676 nm was also observed for the drop-cast film, though the excimer band disappeared because the dimer conformation was suppressed in the film. In comparison, the LB film was found to be almost nonfluorescent. This severe quenching is probably because there exist some energy traps of the chromophores such as the mixtures of J-aggregation and some other dimeric and/or oligomeric assemblies. It is understood that the systems contained a relatively low degree of order, which was also suggested by the absorption spectra. At present, how exactly the quenching source affects is unclear. Nonetheless, the

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CONCLUSIONS In this work, we demonstrate the protocol to prepare twodimensional nanorods of the metal-free pheophorbidesappended cellulose, by taking advantage of both regioselective functionalization of cellulose and an air−water interface as a confined space. The formed nanorod has ca. 0.4−0.5 nm in height with the width of 180 nm and the length of around 500 nm. The height corresponds to the cross-sectional thickness of a glucose ring, indicating that the height can be confined to be monomolecular level owing to the nature of interfacial confined space. The in-plane GIWAXS profile of the nanorod-shaped structure of 1 has two sharp diffraction peaks, suggesting a certain degree of order. The spectrum of the LB monolayer film exhibits the red-shifted absorption peaks as compared to that in solution and of the drop-casted film. This bathochromic shift may be attributed to the J-type excitonic coupling between the S0−S1 transition dipole moments of the chlorin Qy bands,

Figure 7. (A) Photocurrent response with illumination at 420 nm (blue) and 690 nm (red) and (B) action spectrum of the monolayer film of 1 on an ITO electrode. Electrolyte solution: a 0.1 M Na2SO4 aqueous solution containing H2Q (50 mM) as a sacrifice donor. Input power: 1.5 mW cm−2. Bias potential: 0 V vs SCE. 3228


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resulting from the interacting pheophorbides. The photofunctional utilization of the cellulose nanorods was also preliminarily demonstrated by the conventional photocurrent measurements. The approach as developed should lead to generate a wide range of cellulose- and polysaccharide-based nanostructures.

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ASSOCIATED CONTENT

S Supporting Information *

AFM and photocurrent studies. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected] (K.S.); ilkka.kilpelainen@ helsinki.fi (I.K.); [email protected] (K.A.). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Dr. J. Kavakka (University of Helsinki) for the chlorophylls, Professor T. Takano (Kyoto University) for support of photocurrent measurement, and Ms. M. Akada (NIMS) for support of AFM observation. This work was partly supported by KAKENHI (24750218) of MEXT, WPI Initiative of MEXT, CREST program of JST, and research grants from the University of Helsinki and Academy of Finland (Grants 122534, 113317 and 132150).

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REFERENCES

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