Excited-State Proton Transfer and Geminate Recombination in

ARTICLE pubs.acs.org/JPCC

Excited-State Proton Transfer and Geminate Recombination in Hydrogels Based on Self-Assembled Peptide Nanotubes Sun-Young Park,† Hyeok Jeong,† Hahkjoon Kim,‡ Jin Yong Lee,§ and Du-Jeon Jang*,† †

School of Chemistry, Seoul National University, NS60, Seoul 151-742, Korea Department of Chemistry, Duksung Women’s University, Seoul 132-714, Korea § Department of Chemistry, Sungkyunkwan University, Suwon 440-746, Korea ‡

bS Supporting Information ABSTRACT: The excited-state proton transfer (ESPT) of 7-hydroxyquinoline (7HQ) embedded in hydrogels based on self-assembled peptide nanotubes (PNTs) has been investigated with variation of peptides as well as protic hydrogen isotopes. In the inner cavities of PNT-based hydrogels, 7HQ prefers to exist as an anionic species rather than a normal species or a cationic species already in the ground state because the hydrogels are mildly basic. Upon excitation, ESPT takes place from a water molecule in a PNT hydrogel nanocavity to the imino group of the anionic species to produce a zwitterionic species, and its back reaction, that is, geminate recombination between the proton of the iminium group of the zwitterionic species and the deprotonated water molecule, also occurs because of the extremely viscous environment as well as the spatial constraint of semirigid PNT hydrogels. As a result, an excited-state prototropic equilibrium between the anionic and the zwitterionic species is established in PNT hydrogels. Water nanopools confined in smaller PNTs are denser than those in larger PNTs, so that 7HQ molecules in the smaller PNTs have more chances to form strong hydrogen bonds with water molecules. Consequently, both ESPT and geminate-recombination in the smaller PNTs are faster than the respective ones in the larger PNTs.

1. INTRODUCTION In the past few decades, demands for high-speed electronic devices such as molecular computers have increased rapidly, and correspondingly, many efforts to miniaturize physical components of electronic devices have brought about great advances in molecular nanotechnology. Researchers in the fields of the design and the fabrication of nanomaterials such as carbon nanotubes have drawn considerable inspiration from the way that biological systems construct living units by self-assembling supramolecular structures;1
5 many peptide- or protein-based microtubules in living organisms were found to have subcellular structures formed via self-assembly.6
10 Because the molecular self-assembly process of biological systems was reported to facilitate the fabrication of complicated architectures and machinery, it has often been used to design novel materials that could be applicable to microelectronics and microelectro-mechanical systems (MEMS).11
13 In this regard, many researchers have begun to investigate approaches for the manufacture of intelligent, functional, self-assembling materials,3
10 and peptide-based nanotubes have come up as an issue on that account.14
18 For the first time in 1993, Ghadiri et al. reported an artificially synthesized peptide nanotube (PNT) where all side chains of amino acids were allowed to point outward.19 This triggered extensive studies on PNTs and their potential application in molecular wires, catalysts, ion channels, membrane pores, and drug-delivery vehicles.14
21 r 2011 American Chemical Society

A cyclic peptide having two or more amino acids, which is a molecular building block of a PNT, usually favors a low-energy flat conformation.16,19 When more than two cyclic peptides are stacked neatly, an amide backbone is constructed by hydrogen (H) bonds among cyclic peptides in a direction perpendicular to the plane of the cyclic peptide, like contiguous antiparallel β sheets that are commonly found in natural proteins.22 Therefore, a highly ordered parallel array of nanotubular structure with a fixable internal diameter and a modifiable exterior surface, that is, a PNT, is synthesized via self-assembly of peptides, and hundreds of PNTs are packed tightly to form crystalline fibers.15,16,23 Selfassembled PNTs can be utilized in the development of a hydrogel, which is a heavily hydrated material and consists of dilute networks of assembled peptides.24
26 Hydrogels have been a subject of growing interest in biomaterial fields relevant to tissue engineering, axon regeneration, and drug-delivery control because they offer 3D interwoven scaffolds to support the growth of cells.27,28 In an aqueous solution, cyclic peptides are rapidly selfassembled and organized to form a well-ordered 3D structure once self-assembly is triggered by an appropriate chemical or a suitable medium, generating PNT-based hydrogel eventually.26 Received: July 29, 2011 Revised: October 31, 2011 Published: November 07, 2011 24763

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Scheme 1. Equilibria among the Prototropic Species of 7HQ in Water

Figure 1. Molecular structures of Fmoc-β-(2-naphthyl)-L-alanine (Fmoc-NA) and Fmoc-diphenylalanine (Fmoc-FF).

The properties, such as the internal diameter and the external surface, of a PNT can be simply adjusted by choosing a peptide, according to the functionality and the size of the side chain of the peptide.19,29 The flexibility and the hydrophilic interior of PNTs are important features in designing artificial transmembrane ion channels and pores.30
32 Proton transfer is one of the most common and fundamental reactions, and it plays important roles in a wide variety of chemical and biological phenomena.33
57 Because water is the major agent in most of biological processes, biological proton transport is usually mediated by water molecules.50,51 In biological systems, water is often contained in a small pocket of a cell membrane,47
49 and the properties, for example, viscosity and polarity, of such confined water are quite different from the properties of bulk water.42,44 Accordingly, the properties of biological water determine the dynamics of proton transport in biological systems.42
44,47
51 We have previously reported that proton transfer in a water nanopool confined in a reverse micelle or β-cyclodextrin is enormously slow compared with proton transfer in bulk water because the viscosity of confined water is much larger than that of bulk water and the polarity of confined water is much smaller than that of bulk water.38,42 Therefore, to understand the principal mechanisms of biological and chemical processes relevant to vital phenomena, it is indispensable to explore the dynamics of proton transfer mediated by water molecules in confined systems such as the interior of cell membranes. However, it is extremely difficult to investigate proton transport directly in biological systems due to the massiveness and the structural complexity of the systems.35,36 In this regard, PNT-based hydrogels can be good biomimetic systems to study the mechanism and the dynamics of biological proton transport mediated by water molecules in confined systems at the molecular level. Therefore, we have decided to explore excited-state proton transfer (ESPT) in the confined and viscous environment of semirigid PNT hydrogels using 7-hydroxyquinoline (7HQ) as a probe molecule. Hydroxyquinolines, having two prototropic groups of photoacidic enol and photobasic imine, have been extensively explored as good probe melocules to study biological proton-relay processes in diverse biomimetic systems.37
44 The ESPT of 7HQ along H-bonded water molecules takes place in a stepwise manner via forming an anionic species (A), rather than a cationic species (C), resulting in the formation of a zwitterionic species (Z) that is a tautomeric form of 7HQ (Scheme 1).42,43 In bulk water, ESPT is known to occur very quickly (18 MΩ cm) using a Milli-Q system (Millipore). PNT hydrogels having a peptide concentration of 2 mg/mL were prepared by diluting the respective stock solutions of Fmoc-NA and Fmoc-FF in an aqueous solution of 0.1 mM 7HQ. For the measurement of kinetic isotope effects, the protic 1H atoms of 7HQ were exchanged with 2H atoms by dissolving 7HQ in deuterated water (isotropic purity g99.9%, received from Sigma-Aldrich). 24764

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To avoid any preaggregation, we prepared stock solutions freshly prior to each experiment. pH was adjusted by the addition of a dilute HCl or NaOH solution to an aqueous 7HQ solution. 2.2. Measurements. Whereas transmission electron microscopy (TEM) images were obtained with a microscope (JEOL, JEM-2000), absorption spectra were measured with a UV/vis spectrophotometer (Scinco, S3100). Emission spectra were obtained using a home-built fluorometer consisting of a Xe lamp of 75 W (Acton Research, XS432) with a monochromator of 0.15 m (Acton Research, Spectrapro150) and a photomultiplier tube (Acton Research, PD438) attached to a monochromator of 0.30 m (Acton Research, Spectrapro300). Time-resolved fluorescence kinetic profiles with excitation of samples using third-harmonic pulses (355 nm) of a mode-locked Nd:YAG laser of 25 ps (Quantel, Pizzicato) were detected using a streak camera of 10 ps (Hamamatsu, C2830) attached to a CCD detector (Princeton Instruments, RTE128H). Emission wavelengths were selected by combining band-pass and cutoff filters. Fluorescence kinetic constants were extracted by fitting kinetic profiles to computer-simulated exponential curves convoluted with instrument response functions. All of the static and the kinetic measurements were carried out at room temperature.

3. RESULTS AND DISCUSSION 3.1. Preparation and Characterization of Hydrogels. Two types of Fmoc-peptides shown in Figure 1 (Fmoc-NA and FmocFF) were dissolved in DMSO individually to prepare stock solutions of 100 mg/mL; then each stock solution was diluted with an aqueous solution of 0.1 mM 7HQ to a final peptide concentration of 2 mg/mL. Then, phase transformation from DMSO to water induced self-assembly of peptides, resulting in the formation of PNT-based hydrogels.33 Gazit and coworkers have reported that in an aqueous solution under mild conditions simple and short-sized building blocks of Fmoc-FF generate a novel self-assembled PNT hydrogel with remarkable mechanical rigidity;26 the rigidity of the self-assembled PNT hydrogel originates from the aromatic moiety of the peptide building blocks. Interactions among aromatic groups play a key role in the formation of tubular structures because they contribute to the order and the directionality of self-assembly as well as to the formation energy of a PNT hydrogel.25 A PNT hydrogel can be shaped in accordance to a vessel where peptides are injected and self-assembled, and produced PNTs are known to be stable even under extreme conditions.26 These properties of PNT hydrogels allow their utilization in diverse technological applications.26 To gain insight into the self-assembled structure of PNT hydrogels, we have obtained TEM images of Fmoc-NA and Fmoc-FF hydrogels and observed their time-dependent optical transition images. The TEM image of an Fmoc-NA hydrogel shows branching and flexible fibrous PNT networks (Figure 2a). Huppert and coworkers have reported that water molecules form a nanopool in the inner cavity of a PNT and that the radius of the water nanopool varies depending on the concentration as well as the type of the employed peptide.33 The average diameter of water nanopools decreases with the concentration of the peptide, and the average size of PNT hydrogel cavities is smaller in an FmocNA hydrogel than in an Fmoc-FF hydrogel; at a peptide concentration of 0.5 mg/mL, the outer radii of waterpools in Fmoc-NA and Fmoc-FF hydrogels are found to be 13.5 and 15.0 nm, respectively, whereas at a peptide concentration of 4 mg/mL, those in Fmoc-NA and Fmoc-FF hydrogels are estimated to be

Figure 2. (a) TEM image of an Fmoc-NA hydrogel at 5 min after initiation of self-assembly. (b) Optical-transition kinetic images of Fmoc-NA (left) and Fmoc-FF (right) hydrogels.

9.0 and 12.0 nm, respectively.33 We consider that because of an additional amino acid of phenylalanine in Fmoc-FF, a Fmoc-FF hydrogel has larger cavities of PNTs than a Fmoc-NA hydrogel does. The locations as well as the number of aromatic groups in a cyclic peptide, which is the building block of a PNT, have a direct influence on the structure of the PNT.25 In addition to the Fmoc moiety, Fmoc-FF contains two phenyl groups, whereas FmocNA contains one naphthyl group. Of note is that Fmoc moieties are cleaved from peptides under mildly basic conditions to form cyclic peptide chains. From a rheological point of view, the number of aromatic groups and their locations in the peptide backbone affect the properties, such as optical transition kinetics, of PNT-based hydrogels (Figure 2b). 3.2. Absorption Spectra. Whereas ground-state 7HQ in pure water shows N absorption at 327 nm and remarkable Z absorption at 405 nm as well,42 7HQ embedded in a PNT hydrogel does not show N absorption but does show only A absorption at 356 nm in an Fmoc-NA hydrogel and at 351 nm in an Fmoc-FF hydrogel (Figure 3). In bulk water, water molecules can hydrate protons well by forming Eigen and Zundel cations, and fast interconversion between the Eigen and the Zundel cations along the systematically well-organized H-bond network of water molecules results in the anomalously high mobility of a proton.54
57 Accordingly, the ESPT of aqueous 7HQ to form Z takes place rapidly via forming A.42,43 PNT hydrogels are also aqueous media having water nanopools, and two types of peptides employed in this work, that is, Fmoc-NA and Fmoc-FF, have basic amino groups that can accept protons readily (Figure 1). Recall that Fmoc-NA or Fmoc-FF was dissolved in a basic solvent of DMSO and then diluted in neutral water. In these regards, a water solution of DMSO with Fmoc-NA or Fmoc-FF is suggested to be a slightly basic medium, and thus 7HQ in a PNT-based hydrogel 24765

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Figure 3. Maximum-normalized absorption spectra of 7HQ in pure water, an Fmoc-NA hydrogel, and an Fmoc-FF hydrogel.

is considered to exist dominantly as A rather than N or C, already in the ground state. For this reason, we attribute the absorption band around 350
360 nm observed in both hydrogels to A. Of note is that in an aqueous solution at pH 12 the band maximum of A absorption appears at 360 nm (Figure S1 in the Supporting Information). Because of the confined environment of waterpools in PNT hydrogels, the absorption spectrum of A in PNT hydrogels is shifted to the blue slightly compared with that in a bulk basic solution. The absorption band of A in an Fmoc-NA hydrogel is observed to be red-shifted by 5 nm compared with that in an FmocFF hydrogel (Figure 3), suggesting that 7HQ molecules are H-bonded to water molecules more strongly in the Fmoc-NA hydrogel than in the Fmoc-FF hydrogel. We attribute this to the smaller cavity, where 7HQ and water molecules are contained, of the Fmoc-NA hydrogel than the cavity of the Fmoc-FF hydrogel (see above). The average size of water nanopools confined in PNT cavities of Fmoc-FF is larger than that confined in FmocNA hydrogel cavities, although the content of water is the same in both PNT hydrogels. In other words, water nanopools of the Fmoc-NA hydrogel are denser than those of the Fmoc-FF hydrogel.33 Therefore, it can be inferred that A forms a H bond with a water molecule across a shorter distance in the Fmoc-NA hydrogel than in the Fmoc-FF hydrogel. The absorption spectrum of 7HQ in both Fmoc-NA and Fmoc-FF hydrogels does not show Z absorption, presumably around 400
410 nm as observed in neat water. The excitation spectra of 7HQ also support that only A exists in the ground state in both PNT hydrogels (Figure S2 in the Supporting Information). This indicates that proton transfer from a water molecule to the imino group of A to produce Z hardly takes place in the ground state. However, upon excitation, the imino group of A* becomes much more basic than that of A, so that ESPT from a water molecule to the imino group of A* to produce Z* occurs readily in both Fmoc-FF and Fmoc-NA hydrogels, resulting in dominant Z* fluorescence with weak A* fluorescence. (See below.) 3.3. Steady-State Fluorescence Spectra. With excitation at 355 nm of A, the emission spectrum of 7HQ in an aqueous solution at pH 12 shows dominant Z* fluorescence at 508 nm with very weak A* fluorescence around 420 nm (Figure S1 in the Supporting Information). The emission spectrum of 7HQ in water shows a Z* fluorescence band only at 508 nm (Figure 4), indicating that all of the 7HQ molecules undergo ESPT rapidly along H-bonded water molecules to form Z*. In a PNT hydrogel, however, the excitation of A at 355 nm gives rise to A* fluorescence at 420 nm as well as Z* fluorescence, suggesting that not

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Figure 4. Maximum-normalized emission spectra, with excitation at 355 nm, of 7HQ in pure water, an Fmoc-NA hydrogel, and an Fmoc-FF hydrogel.

only the forward reaction, that is, the protonation of the imino group in A* to form Z*, but also the reverse reaction, that is, the deprotonation of the iminium group in Z* to form A* again, takes place (see below). We attribute this to the spatial constraint of PNT cavities in which 7HQ and water molecules are contained.38,39b,42 In other words, the confined and viscous environment of water nanopools inside PNT hydrogel cavities induces geminate recombination between the deprotonated water molecule and the proton of the iminium group in Z*, resulting in the reformation of A*. However, the intensity of Z* fluorescence is much larger than that of A* fluorescence in both hydrogels, indicating that in a PNT hydrogel the forward reaction is dominant compared with the reverse reaction. Correspondingly, the protonation rate of A* to form Z* is found to be substantially larger than the deprotonation rate of Z* to reform A* (see below). The band maximum of Z* fluorescence is at 503 nm in an Fmoc-NA hydrogel and at 508 nm in an Fmoc-FF hydrogel, and the intensity of A* fluorescence is much larger in the Fmoc-NA hydrogel than that in the Fmoc-FF hydrogel (Figure 4). As mentioned above, 7HQ molecules in the Fmoc-NA hydrogel interact with water molecules across a shorter distance than those in the Fmoc-FF hydrogel do because of denser waterpools in the Fmoc-NA hydrogel.33 Although 7HQ molecules in both hydrogels are confined in the cavities of PNTs containing water molecules, 7HQ molecules in the Fmoc-NA hydrogel, in particular, are hemmed more tightly by water molecules than those in the Fmoc-FF hydrogel because of the narrower cavities of the Fmoc-NA hydrogel. Therefore, we consider that geminate recombination between a deprotonated water molecule, that is, a hydroxide ion, and the proton of the iminium group in Z* occurs more actively in the Fmoc-NA hydrogel than in the Fmoc-FF hydrogel. As a result, the population of A* within the lifetime of Z* is much higher in the Fmoc-NA hydrogel than in the Fmoc-FF hydrogel (Figure 4). 3.4. Time-Resolved Fluorescence Kinetic Profiles. Upon excitation at 355 nm, the fluorescence kinetic profiles of 7HQ in Fmoc-NA and Fmoc-FF hydrogels with variation of protic hydrogen isotopes have been monitored at 420 nm for A* fluorescence and at 600 nm for Z* fluorescence (Figure 5 and Figure S3 in the Supporting Information). The kinetic profile of each prototropic species has been monitored without being interfered with by the fluorescence of the other prototropic species. Nonetheless, for each sample, both kinetic profiles monitored at 420 and 600 nm have two components of the same time scales regardless of monitored wavelengths, whereas each kinetic profile has an additional long 24766

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Table 1. Fluorescence Time Constants of Excited-State 7HQ in PNT Hydrogels peptide Fmoc-NA

isotope 1

H

2

H

λema/nm

rise time/ps

420

b

600

15

decay time/ps 15 (64%)c + 200 (27%) + 2300 (9%) 200 (35%) + 3000 (65%)

420

30 (52%) + 270(38%) + 3400 (10%)

600 Fmoc-FF

1

H

30

270 (46%) + 4000 (54%)

420

20 (56%) + 180 (32%) + 2100 (12%)

600 2

H

20

420 600

40 (65%) + 240 (27%) + 3200 (8%) 40

Wavelength of monitored fluorescence. percentage. a

Figure 5. Fluorescence kinetic profiles of 7HQ in Fmoc-NA (a) and Fmoc-FF (b) hydrogels. Samples were excited at 355 nm and monitored at 420 nm for A* fluorescence (open) and at 600 nm for Z* fluorescence (closed). The protic hydrogen isotopes of 1H and 2H are indicated inside, and solid lines are the best-fitted curves to extract the time constants.

decay component (Table 1); for example, 71HQ in the Fmoc-NA hydrogel has two components of 15 and 200 ps, whereas 72HQ in the Fmoc-NA hydrogel has two components of 30 and 270 ps. The initial fractional amplitudes of those two components vary with monitored wavelengths. The longest decay times observed at 420 and 600 nm are attributed to the relaxation time constants of A* and Z*, respectively, and they are observed to be similar to each other in each sample (Table 1). These results suggest that not only proton transfer from a water molecule to the imino group of A* to form Z* but also its back reaction, that is, proton transfer from the iminium group of Z* to the deprotonated water molecule to form A* again, takes place within the lifetime of excited-state 7HQ in PNT hydrogels.38,39 This is attributed to the spatial constraint and the viscous environment of water nanopools confined in PNT cavities. Accordingly, geminate recombination between the proton of the iminium group in Z* and the deprotonated water molecule should also be considered to fit fluorescence kinetic profiles. Of note is that waterpools confined in reverse micelles have been reported to be much more viscous than bulk water.42,44 Moreover, in confined systems such

180 (17%) + 2500 (83%)

240 (8%) + 3700 (92%) b

Instant. c Initial intensity

as nanocavities of mesoporous zeolite materials or cyclodextrins, geminate recombination as well as proton transfer has been reported to take place in the excited state because of the spatial constraint.38,39a,47c Because of the back reaction of the ESPT of 7HQ, that is, geminate recombination, kinetic constants for the ESPT of 7HQ in PNT hydrogels become more complicated than those for the ESPT of 7HQ in neat water.33 We have deduced rate constants for both the protonation of the imino group in A* and the deprotonation of iminium group in Z* in PNT hydrogels through simple calculations (see below). Figure 5 shows that in both Fmoc-NA and Fmoc-FF hydrogels the ESPT of 72HQ is significantly slowed down in comparison with that of 71HQ. In confined and viscous media of water nanopools in PNT cavities, a 2 H atom is hard to transfer or diffuse away compared with a 1H atom under the same conditions because of its heavier mass. Furthermore, the activation energy for the ESPT of a 2H atom is usually larger than that for the ESPT of a 1H atom because of the lower zero-point energy of 72HQ. 3.5. Kinetic Analysis. In confined or rigid systems, following intermolecular proton transfer from a protic solvent molecule to a photobasic molecule, geminate recombination between the transferred proton and the deprotonated solvent molecule often takes place.33,38,39 Recall that two short-time constants extracted from the fluorescence kinetic profiles of 7HQ are invariant regardless of prototropic species in each sample, although their initial fractional amplitudes vary with prototropic species (Table 1). In this regard, we suggest that both the protonation of the imino group in A* to form Z* and the subsequent deprotonation of the iminium group in Z* to reform A* take place, as shown in Scheme 2. In other words, because of the confined and viscous environment of water nanopools in semirigid PNT hydrogels, proton transfer from a water molecule to the imino group of A* is considered to compete with its back reaction that is geminate recombination between the proton of the iminium group in Z* and the deprotonated water molecule, that is, a hydroxide ion. In this case, rate constants for the protonation of A* (kp) and the deprotonation of Z* (kd) (Scheme 2) should be deduced by calculations using observed kinetic constants given in Table 1 (see below). Of note is that the relaxation constants of A* (kA*) and Z* (kZ*) are taken from the longest times observed at 420 and 550 nm, respectively. 24767

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Scheme 2. Summary Scheme for the Prototropic Equilibrium and the Relaxation of Excited-State 7HQ Embedded in PNT Hydrogels

Table 2. Rate Constants for the Proton Transfer and the Relaxation of Excited-State 7HQ in PNT Hydrogels peptide Fmoc-NA

kp
1/ps

kd
1/ps

kA*
1/ps

kZ*
1/ps

1

21

43

2300

3000

2

49

63

3400

4000

1

30 54

47 100

2100 3200

2500 3700

isotope H H

Fmon-FF

H 2 H

Assuming that both the proton transfer and the geminate recombination of excited-state 7HQ in a PNT hydrogel follow a first-order reaction, we have analyzed observed kinetic constants according to following equations. The fluorescence intensity profiles of A* (IA*) and Z* (IZ*) with time (t) obey eqs 1 and 2, respectively. IA ðtÞ ¼ a1 e
λ1 t þ a2 e
λ2 t þ a3 e
λ3 t

ð1Þ

IZ ðtÞ ¼ að
e
λ1 t þ e
λ2 t þ e
λ4 t Þ

ð2Þ

where λ1, λ2, and λ3 are the respective reciprocals of three time constants having the fractional initial amplitudes of a1, a2, and a3, respectively, observed at 420 nm, whereas λ4 is the reciprocal of the longest decay time observed at 550 nm. Following the global analysis,59,60 we have obtained kp and kd (Table 2) using eqs 3
7. As mentioned above, λ3 and λ4 are used as kA* and kZ*, respectively. qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi ð3Þ λ1, 2 ¼ fX þ Y ( ðX
Y Þ2 þ 4kp kd g=2 X ¼ kp þ kA 

ð4Þ

Y ¼ kd þ kZ

ð5Þ

a1 =a2 ¼ ðX
λ2 Þ=ðλ1
XÞ

ð6Þ

Y ¼ λ1 þ λ2
X

ð7Þ

On one hand, the value of kp is found to be much larger than the value of kd in both Fmoc-NA and Fmoc-FF hydrogels regardless of protic hydrogen isotopes because Z* is energetically lower than A* in aqueous media. Correspondingly, the emission spectra of 7HQ in both hydrogels show dominant Z* fluorescence with weak A* fluorescence (Figure 4). The kinetic isotope effect of k(1H)/k(2H) is deduced to be 2.3 for kp and 1.5 for kd in

the Fmoc-NA hydrogel and 1.8 for kp and 2.1 for kd in the FmocFF hydrogel. A 2H atom is heavier than a 1H atom, and the zeropoint energy of 72HQ is lower than that of 71HQ, requiring a larger activation energy for the ESPT of 72HQ. Accordingly, in extremely viscous water nanopools confined in PNT cavities, a 2 H atom is hard to transfer or diffuse away in comparison with a 1 H atom. The fact that kinetic isotope effect is as large as ∼2 for both kp and kd supports that proton diffusion is not important in PNT hydrogels. On the other hand, the values of both kp and kd are significantly larger in the Fmoc-NA hydrogel than in the Fmoc-FF hydrogel. We attribute this to the fact that the cavities of Fmoc-NA PNTs are smaller than those of Fmoc-FF PNTs (see above). Whereas the content of water is the same in both Fmoc-NA and Fmoc-FF hydrogels, the Fmoc-FF PNT cavity is larger than the Fmoc-NA PNT cavity, leading to denser waterpools in the Fmoc-NA hydrogel.33 Therefore, we consider that narrower spaces inside Fmoc-NA hydrogel cavities make 7HQ molecules have more chances to form strong H bonds with water molecules. Consequently, the ESPT of 7HQ and its back reaction occur more readily in the Fmoc-NA hydrogel than in the FmocFF hydrogel. In biological systems, water is often contained in a small pocket such as a cell membrane, and such confined water is known to be much more viscous than bulk water.47
49 Accordingly, highly viscous water nanopools confined inside cavities of semirigid PNT hydrogels can be a good biomimetic system. Therefore, our results would provide clues in understanding confinement effects on biological processes as well as proton-transport dynamics in viscous biological environment.

4. CONCLUSIONS We have explored the ESPT of 7HQ embedded in hydrogels based on PNTs that were prepared by the self-assembly of Fmocβ-(2-naphthyl)-L-alanine (Fmoc-NA) and Fmoc-diphenylalanine (Fmoc-FF). 7HQ molecules are confined in water nanopools inside the PNT cavities of the hydrogels, which are mildly basic, and thus 7HQ exists as an anionic species (A) dominantly, rather than a normal species or a cationic species, already in the ground state. Because of the confined and extremely viscous media of water nanopools in PNT hydrogel cavities, proton transfer from a water molecule to the imino group of A* to form the zwitterionic species (Z*) of 7HQ competes with its back reaction that is geminate recombination between the deprotonated water molecule and the proton of the iminium group of Z* to form A* again. Consequently, the spatial constraint of viscous waterpools in semirigid PNT hydrogels induces a prototropic equilibrium between A* and Z*. The rate constant of A* protonation (kp) is found to be much larger than the rate constant of Z* deprotonation (kd) in both Fmoc-NA and Fmoc-FF hydrogels because Z* is energetically lower than A*. Water nanopools are denser in an Fmoc-NA hydrogel than in an Fmoc-FF hydrogel so that H bonds between 7HQ and water molecules form across a shorter distance in the Fmoc-NA hydrogel. Accordingly, both kp and kd are significantly larger in the Fmoc-NA hydrogel than in the Fmoc-FF hydrogel. The kinetic isotope effect of k(1H)/k(2H) has been found to be ∼2. In confined and viscous environment of waterpools in PNT hydrogel cavities, a 2H atom is hard to transfer or diffuse away compared with a 1H atom because of its heavier mass. Our results can shed light on fundamental mechanistic elucidation for the molecular dynamics of proton relay in biologically confined systems such as protein membranes. 24768

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’ ASSOCIATED CONTENT

bS

Supporting Information. Absorption and emission spectra of anionic species, excitation spectra, and fluorescence kinetic profiles in a time window of 2.5 ns. This material is available free of charge via the Internet at http://pubs.acs.org.

’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. Tel: +82-(2)-880-4368. Fax: +82-(2)875-6624.

’ ACKNOWLEDGMENT This work was financially supported by research grants through the National Research Foundation (NRF) of Korea by the Ministry of Education, Science, and Technology (2011-0001216, 20110003074, and 2011-0028981). J.Y.L. is thankful for NRF grants (2011-0001211 and 2011-0015767), whereas S.-Y.P. acknowledges the Seoul fellowship and the BK21 scholarship. ’ REFERENCES (1) Iijima, S. Nature 1991, 354, 56–58. (2) Gattuso, G.; Menzer, S.; Nepogodiev, S. A.; Stoddart, J. F.; Williams, D. J. Angew. Chem., Int. Ed. Engl. 1997, 36, 1451–1454. (3) Harada, A.; Li, J.; Kamachi, M. Nature 1993, 364, 516–518. (4) Nelson, J. C; Saven, J. G.; Moore, J. S.; Wolynes, P. G. Science 1997, 277, 1793–1796. (5) Konig, B. Angew. Chem., Int. Ed. Engl. 1997, 36, 1833–1835. (6) Schnur, J. M. Science 1993, 262, 1669–1676. (7) Perutz, M. F.; Finch, J. T.; Berriman, J.; Lesk, A. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 5591–5595. (8) Graveland-Bikker, J. F.; Ipsen, R.; Otte, J.; Kruif, C. G. Langmuir 2004, 20, 6841–6846. (9) Abrahams, B. F.; Hoskins, B. F.; Michail, D. M.; Robson, R. Nature 1994, 369, 727–729. (10) Mitchell, J. C.; Harris, J. R.; Malo, J.; Bath, J.; Turberfield, A. J. J. Am. Chem. Soc. 2004, 126, 16342–16343. (11) Whrtesides, G. M.; Mathias, J. P.; Seto, C. T. Science 1991, 254, 1312–1319. (12) Mao, C.; Solis, D. J.; Reiss, B. D.; Kottmann, S. T.; Sweeney, R. Y.; Hayhurst, A.; Georgiou, G.; Iverson, B.; Belcher, A. M. Science 2004, 303, 213–217. (13) Nam, K. T.; Kim, D.-W.; Yoo, P. J.; Chiang, C. Y.; Meethong, N.; Hammond, P. T.; Chiang, Y.-M.; Belcher1, A. M. Science 2006, 312, 885–888. (14) Lehn, J.-M. Supramolecular Chemistry: Concepts and Perspectives; VCH Weinheim: New York, 1995; pp 1
30. (15) Ghadiri, M. R. Adv. Mater. 1995, 7, 675–677. (16) Buriak, J. M.; Ghadiri, M. R. Mater. Sci. Eng., C 1997, 4, 207–212. (17) Gorbitz, C. H. Chem.—Eur. J. 2001, 7, 5153–5159. (18) Reches, M.; Gazit, E. Curr. Nanosci. 2006, 2, 105–111. (19) Ghadiri, M. R.; Granja, J. R.; Milligan, R. A.; McRee, D. E.; Khazanovich, N. Nature 1993, 366, 324–327. (20) Zhang, S. Nat. Biotechnol. 2003, 21, 1171–1178. (21) Patolsky, F.; Weizmann, Y.; Willner, I. Nat. Mater. 2004, 3, 692–695. (22) Smith, A. M.; Williams, R. J.; Tang, C.; Coppo, P.; Collins, R. F.; Turner, M. L.; Saiani, A.; Ulijn, R. V. Adv. Mater. 2008, 20, 37–41. (23) Reches, M; Gazit, E. Nat. Nanotechnol. 2006, 1, 195–200. (24) Vegners, R.; Shestakova, I.; Kalvinsh, I.; Ezzell, R. M.; Janmey, P. A. J. Pep. Sci. 1995, 1, 371–378. (25) Orbach, R.; Adler-Abramovich, L.; Zigerson, S.; Mironi-Harpaz, I.; Seliktar, D.; Gazit, E. Biomacromolecules 2009, 10, 2646–2651.

ARTICLE

(26) Mahler, A.; Reches, M.; Rechter, M.; Cohen, S.; Gazit, E. Adv. Mater. 2006, 18, 1365–1370. (27) Langer, R.; Vacanti, J. P. Science 1993, 260, 920–926. (28) Almany, L.; Seliktar, D. Biomaterials 2005, 26, 2467–2477. (29) Hartgerink, J. D.; Granja, J. R.; Milligan, R. A.; Ghadiri, M. R. J. Am. Chem. Soc. 1996, 118, 43–50. (30) Sanchez-Quesada, J.; Isler, M. P.; Ghadiri, M. R. J. Am. Chem. Soc. 2002, 124, 10004–10005. (31) Sanchez-Quesada, J.; Kim, H. S.; Ghadiri, M. R. Angew. Chem., Int. Ed. 2001, 40, 2503–2506. (32) Motesharei, K.; Ghadiri, M. R. J. Am. Chem. Soc. 1997, 119, 11306–11312. (33) Amdursky, N.; Orbach, R.; Gazit, E.; Huppert, D. J. Phys. Chem. C 2009, 113, 19500–19505. (34) (a) Cohen, B.; Segal, J.; Huppert, D. J. Phys. Chem. A 2002, 106, 7462–7467. (b) Leiderman, P.; Genosar, L.; Huppert, D. J. Phys. Chem. A 2005, 109, 5965–5977. (c) Gepshtein, R.; Leiderman, P.; Genosar, L.; Huppert, D. J. Phys. Chem. A 2005, 109, 9674–9684. (35) Mathias, G.; Marx, D. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 6980–6985. (36) (a) Faxen, K.; Gilderson, G.; Adelroth, P.; Brzezinski, P. Nature 2005, 437, 286–289. (b) Kohen, A.; Cannio, R.; Bartolucci, S.; Klinman, J. P. Nature 1999, 399, 496–499. (37) (a) Park, S.-Y.; Jang, D.-J. J. Am. Chem. Soc. 2010, 132, 297–302. (b) Park, S.-Y.; Lee, Y.-S.; Jang, D.-J. Phys. Chem. Chem. Phys. 2011, 13, 3730–3736. (38) Lee, Y.-S.; Kwon, O.-H.; Park, H. J.; Franz, J.; Jang, D.-J. J. Photochem. Photobiol., A 2008, 194, 105–109. (39) (a) Park, S.-Y.; Yu, H.; Park, J.; Jang, D.-J. Chem.—Eur. J. 2010, 16, 12609–12615. (b) Park, S.-Y.; Lee, Y.-S.; Jang, D.-J. Phys. Chem. Chem. Phys. 2008, 10, 6703–6707. (40) (a) Bardez, E.; Chatelain, A.; Larrey, B.; Valeur, B. J. Phys. Chem. 1994, 98, 2357–2366. (b) Bardez, E.; Fedorov, A.; Berberan-Santos, M. N.; Martinho, J. M. G. J. Phys. Chem. A 1999, 103, 4131–4136. (41) (a) Tokumura, K.; Itoh, M. J. Phys. Chem. 1984, 88, 3921–3923. (b) Lavin, A.; Collins, S. Chem. Phys. Lett. 1993, 204, 96–100. (c) Bohra, A.; Lavin, A.; Collins, S. J. Phys. Chem. 1994, 98, 11424–11427. (42) (a) Park, S.-Y.; Kwon, O.-H.; Kim, T. G.; Jang, D.-J. J. Phys. Chem. C 2009, 113, 16110–16115. (b) Kwon, O.-H.; Kim, T. G.; Lee, Y.-S.; Jang, D.-J. J. Phys. Chem. B 2006, 110, 11997–12004. (c) Park, H. J.; Kwon, O.-H.; Ah, C. S.; Jang, D.-J. J. Phys. Chem. B 2005, 109, 3938–3943. (43) (a) Park, S.-Y.; Lee, Y.-S.; Kwon, O.-H.; Jang, D.-J. Chem. Commun. 2009, 926–928. (b) Park, S.-Y.; Kim, B.; Lee, Y.-S.; Kwon, O.-H.; Jang, D.-J. Photochem. Photobiol. Sci. 2009, 8, 1611–1617. (44) (a) Garcia-Ochoa, I.; Bisht, P. B.; Sanchez, F.; Martinez-Ataz, E.; Santos, L.; Tripathi, H. B.; Douhal, A. J. Phys. Chem. A 1998, 102, 8871–8880. (b) Angulo, G.; Organero, J. A.; Carranza, M. A.; Douhal, A. J. Phys. Chem. B 2006, 110, 24231–24237. (c) Douhal, A.; Angulo, G.; Gil, M.; Organero, J. A.; Sanz, M.; Tormo, L. J. Phys. Chem. B 2007, 111, 5487–5493. (45) Park, S.-Y.; Jeong, H.; Jang, D. J. J. Phys. Chem. B 2011, 115, 6023–6031. (46) (a) Chudoba, C.; Nibbering, E. T. J.; Elsaesser, T. J. Phys. Chem. A 1999, 103, 5625–5628. (b) Rini, M.; Dreyer, J.; Nibbering, E. T. J.; Elsaesser, T. Chem. Phys. Lett. 2003, 374, 13–19. (47) (a) Douhal, A.; Lahmani, F.; Zewail, A. H. Chem. Phys. 1996, 207, 477–4983. (b) Douhal, A.; Fiebig, T.; Chachisvilis, M.; Zewail, A. H. J. Phys. Chem. A 1998, 102, 1658–1660. (c) Chachisvilis, M.; GarciaOchoa, I.; Douhal, A.; Zewail, A. H. Chem. Phys. Lett. 1998, 293, 153–159. (d) Pal, S. K.; Zewail, A. H. Chem. Rev. 2004, 104, 2099–2123. (48) (a) Moilanen, D. E.; Fenn, E. E.; Wong, D.; Fayer, M. D. J. Phys. Chem. B 2009, 113, 8560–8568. (b) Spry, D. B.; Fayer, M. D. J. Phys. Chem. B 2009, 113, 10210–10221. (49) (a) Pal, S.; Balasubramanian, S.; Bagchi, B. J. Chem. Phys. 2004, 120, 1912–1920. (b) Pal, S.; Maiti, P. K.; Bagchi, B.; Hynes, J. T. J. Phys. Chem. B 2006, 110, 26396–26402. (c) Rey, R.; Moller, K. B.; Hynes, J. T. J. Phys. Chem. A 2002, 106, 11993–11996. (d) Laage, D.; Hynes, J. T. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 11167–11172. 24769

dx.doi.org/10.1021/jp207245q |J. Phys. Chem. C 2011, 115, 24763–24770

The Journal of Physical Chemistry C

ARTICLE

(50) (a) Bhattacharyya, K. Chem. Commun. 2008, 2848–2857. (b) Nandi, N.; Bhattacharyya, K.; Bagchi, B. Chem. Rev. 2000, 100, 2013–2045. (51) (a) Siwick, B. J.; Cox, M. J.; Bakker, H. J. J. Phys. Chem. B 2008, 112, 378–389. (b) Siwick, B. J.; Bakker, H. J. J. Am. Chem. Soc. 2007, 129, 13412–13420. (c) Cox, M. J.; Timmer, R. L. A.; Bakker, H. J.; Park, S.; Agmon, N. J. Phys. Chem. A 2009, 113, 6599–6606. (d) Tielrooij, K. J.; Cox, M. J.; Bakker, H. J. ChemPhysChem 2009, 10, 245–251. (52) (a) Mohammed, O. F.; Pines, D.; Nibbering, E. T. J.; Pines, E. Angew. Chem., Int. Ed. 2007, 46, 1458–1461. (b) Rini, M.; Magnes, B.-A.; Pines, E.; Nibbering, E. T. J. Science 2003, 301, 349–352. (c) Mohammed, O. F.; Pines, D.; Dreyer, J.; Pines, E.; Nibbering, E. T. J. Science 2005, 310, 83–86. (53) Chen, H.; Voth, G. A.; Agmon, N. J. Phys. Chem. B 2010, 114, 333–339. (54) Marx, D.; Tuckerman, M. E.; Hutter, J.; Parrinello, M. Nature 1999, 397, 601–604. (55) Eigen, M. Angew. Chem., Int. Ed. Engl. 1964, 3, 1–19. (56) Zundel, G.; Metzger, H. Z. Phys. Chem. 1968, 58, 225–233. (57) Marx, D. ChemPhysChem 2006, 7, 1848–1870. (58) Chan, W. C.; White, P. D. Fmoc Solid Phase Peptide Synthesis: A Practical Approach; Oxford University Press: New York, 2000; pp 41
76. (59) Giestas, L.; Yihwa, C.; Lima, J. C.; Vautier-Giongo, C.; Lopes, A.; Macanita, A. L.; Quina, F. H. J. Phys. Chem. A 2003, 107, 3263–3269. (60) Lima, J. C.; Abreu, I.; Brouillard, R.; Macanita, A. L. Chem. Phys. Lett. 1998, 298, 189–195.

24770

dx.doi.org/10.1021/jp207245q |J. Phys. Chem. C 2011, 115, 24763–24770