Trace Solvent as a Predominant Factor To Tune Dipeptide Self-Assembly Juan Wang,†,‡ Kai Liu,†,‡,§ Linyin Yan,†,‡ Anhe Wang,† Shuo Bai,† and Xuehai Yan*,†,‡ †
National Key Laboratory of Biochemical Engineering, Institute of Process Engineering, and ‡Center for Mesoscience, Institute of Process Engineering, Chinese Academy of Sciences, Beijing 100190, People’s Repubic of China § University of Chinese Academy of Sciences, Beijing 100049, People’s Republic of China S Supporting Information *
ABSTRACT: Solvent molecules such as water are of key importance for tuning self-assembly in biological systems. However, it remains a great challenge to detect the role of different types of noncovalent interactions between trace solvents and biomolecules such as peptides. In this work, we discover a dominant role of trace amounts of solvents for mediation of dipeptide self-assembly, in which solvent-bridged hydrogen bonding is demonstrated as a crucial force in directing fiber formation. Hydrogen-bond-forming solvents (including ethanol, N,N-dimethylformamide, and acetone) can affect the hydrogen bonding of CO and N−H in diphenylalanine (FF) molecules with themselves, but this does not induce π−π stacking between FF molecules. The directional hydrogen bonding promotes a long-range-ordered arrangement of FF molecules, preferentially along one dimension to form nanofibers or nanobelts. Furthermore, we demonstrate that water with strong hydrogen-bond-forming capability can notably speed up structure formation with long-range order, revealing the importance of water as a trace solvent for regulation of persistent and robust fiber formation. KEYWORDS: dipeptide, self-assembly, trace solvent, fibers, hydrogen bonding
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peptide building blocks, has been found to be capable of selfassembly into a variety of complex structures, such as vertically aligned nanoforests or nanowires, films, and fiber/tube bundles.5−24 It may be the most suitable candidate for study of structural formation and regulation due to simple molecular structure and distinct intermolecular interactions. For example, diverse structures including nanotubes, nanowires, and nanofibers can be readily achieved by altering solvent properties: In an aqueous environment, FF tends to form nanotubes,6 whereas FF undergoes self-assembly for formation of long nanofibrils in nonpolar toluene and flower-like microcrystals in polar ethanol.25 This is because the solvent properties, such as polarity and hydrogen bonding ability, will directly influence local interactions of peptide self-assembly on the molecular level.9 These are closely related to solubility parameters of solvents such as Hansen solubility parameters (HSP), which include three important parameters, that is, dispersion (δd), dipole−dipole (δp), and hydrogen-bonding (δh) interactions.26 However, how the introduction of trace solvents with different solubility parameters influences local interactions of selfassembling building blocks remains to be further explored.
elf-assembling systems such as phospholipid membranes, DNA double helix, and protein folding structures play a significant role in the physiological function. Structure formation and transformation of these self-assembling architectures in terms of weak intermolecular interactions trigger life’s functions or induce various diseases. For example, protein fibril formation is thought to be a major cause of Alzheimer’s disease.1,2 In this process, water-assisted hydrogen bonding is believed to be a critical factor for self-assembly. Actually, molecular self-assembly is a spontaneous process of formation of ordered structures under the synergistic effect of intermolecular noncovalent interactions, including hydrogen bonding, π−π, electrostatic, hydrophobic, and van der Waals interactions.3 Even tiny amounts of solvents (e.g., water) will disturb the synergistic effect of these interactions and further change the assembly of macromolecules as proteins. Therefore, processes for tuning self-assembling structures of biologically relevant molecules (such as peptides) by trace solvents and insight into the noncovalent interactions between solvents and biomolecules are of crucial importance for comprehending the organization mechanism of complex systems. Herein, we set out to better understand the significance of trace solvents in tuning self-assembly of an aromatic dipeptide consisting of two covalently linked phenylalanine units (diphenylalanine, FF), which is a key structural motif in Alzheimer’s β-amyloid polypeptide.4 FF, as one of the simplest © XXXX American Chemical Society
Received: October 19, 2015 Accepted: January 12, 2016
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DOI: 10.1021/acsnano.5b06567 ACS Nano XXXX, XXX, XXX−XXX
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ACS Nano In this work, we for the first time discover that a trace amount of solvent may be a dominant factor for directing and mediating the dipeptide self-assembly. The hydrogen-bonding (δh) interaction of trace solvents is found to play a key role in tuning the formation of fibers with long-range order. Hydrogen-bond-forming solvents such as ethanol, N,N-dimethylformamide (DMF), and acetone, although a small amount is introduced for tuning self-assembly, interact with dipeptide molecules presumably through solvent-bridged hydrogen bonding. The solvent-bridged hydrogen bonding promotes a long-range-ordered arrangement of FF molecules, preferentially along one dimension to form nanofibers or nanobelts (Scheme 1). The finding of trace-solvent-tuning self-assembly may Scheme 1. Schematic Depiction of a Phase Transition Induced by Trace Amounts of Solventa
a
Hydrogen-bond-forming solvents, such as ethanol, DMF, and acetone, can induce FF to form fiber structures. Figure 1. SEM images of (a) FF/CH2Cl2, (b) FF/CH2Cl2/ethanol, (c) FF/CH2Cl2/DMF, and (d) FF/CH2Cl2/acetone systems. The amount of ethanol (DMF or acetone) in CH2Cl2 is 5% (v/v). (e) AFM height image of the FF/CH2Cl2/ethanol system, showing the presence of well-defined fibers. (f) Corresponding height profiles along the green line, showing a fiber height of approximately 15 nm.
provide references for understanding structure formation and transformation of biomolecules in biological systems. This also opens up an alternative way for mediating assembly and functional integration of a range of biology-derived molecules.
RESULTS AND DISCUSSION We chose FF/dichloromethane as a model. In the FF/ dichloromethane solution, there is no hydrogen bond or π−π interaction between FF and dichloromethane (CH2Cl2) molecules. Three types of trace solvents (Table S1) are used. Type I is a solvent that has hydrogen-bonding (δh) interactions with FF, including ethanol, DMF, and acetone. Type II is a solvent with π−π interactions with FF, including toluene. Type III is a solvent that only has van der Waals interactions with FF, including n-hexane. The optical microscopy images of samples in solution showed that FF underwent crystallization in pure CH2Cl2, while gelling in CH2Cl2 with a trace amount of hydrogen-bond-forming solvents (Figure S1). Scanning electron microscopy (SEM) images of the samples dried by solvent evaporation show that the FF crystal has a flower-like morphology and the “flower” is composed of short rods (Figure 1a), while the addition of hydrogen-bond-forming solvents successfully induces FF to form fiber or ribbon structures (Figure 1b−d). These observations suggest that hydrogen-bonding (δh) interactions play a unique and key role in fiber formation. Moreover, to avoid the influence of different vapor pressures between CH2Cl2 and other solvents on the morphologies developed during the solvent evaporation drying process, the samples were freeze-dried for atomic force microscopy (AFM) observation with assistance of liquid nitrogen. The AFM image of the freeze-dried gel shows that it consists of well-defined fibers (Figure 1e), and the height of the fiber is about 15 nm (Figure 1f), consistent with the SEM observation for the solvent evaporation dried samples.
Next, we will characterize the effect of hydrogen bonding in detail. Take the ethanol system as a typical example. We observed that the precipitates formed in FF/CH2Cl2/ethanol systems were mainly microcrystals if the content of ethanol was lower than 3% (Figure S2). The turbidity of these systems was increased with increasing content of ethanol. When the ethanol content was increased to 3%, the systems formed an organogel with little opacity (Figure S2). SEM images show that, after the addition of ethanol, the short rods (Figure 1a) appear flaky (Figure 2a,b). When the ethanol content is increased to 3%, an organogel consisting of long fibers is formed (Figure 2c,d). These fibers intertwine with each other firmly to form a network. Transmission electron microscopy (TEM) images also agree with the SEM observations (Figure S2). The results of SEM and TEM images indicate that even trace amounts of ethanol can successfully induce FF to form various morphologies. The solution-based Fourier transform infrared spectra (FTIR) of FF in pure CH2Cl2 (Figure 3a) show a strong amide I absorption band (vibration of CO) in the vicinity of 1675 cm−1 and an amide II absorption band (in-plane vibration of N−H) near 1602 cm−1, indicating a hydrogen-bonded βsheet secondary structure.27,28 When a trace amount of ethanol was added, the peaks at 1602 cm−1 red-shifted to about 1587 cm−1, and the peaks at 1675 cm−1 shifted to about 1686 cm−1 in gel systems (Figure 3a). This is because the addition of ethanol affects the hydrogen bonding of CO and N−H in FF molecules. The detailed hydrogen bonding pattern can be B
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are obviously changed and the signals of the hexagonal structure disappear. This reveals that long-range order rather than a hexagonal structure is formed. Furthermore, FL (Figure 3d) of the microcrystals and fibers obtained with ethanol content of 2 and 3%, respectively, shows the same emission peak at about 293 nm. This value, close to that obtained in 100% ethanol (285 nm),25,31 reveals that no π−π stacking structures form in these systems. In this system, there is no hydrogen-bonding or π−π interaction between FF and CH2Cl2 molecules. Therefore, in pure CH2Cl2, crystallization is favored with growth of FF into each dimension at a more or less comparable rate. When hydrogen-bond-forming solvents are added to CH2Cl2, directional hydrogen bonding will promote the growth (assembly) of FF molecules in one dimension and result in the formation of fibers. In order to prove the effect of hydrogen bonding, water, which has stronger hydrogen-bonding (δh) ability than ethanol, was added to the solution. Considering the immiscibility between water and CH2Cl2, water was used to partially replace ethanol. Specifically, ethanol was first mixed with water. Then, the ethanol/water mixture was mixed with CH2Cl2. The ethanol/water content in CH2Cl2 is fixed at 2%, but the ratios of water to ethanol are 1:19, 1:9, and 1:4. It is interesting to note that when the ratio of water to ethanol reached 1:9, the systems form gels with fiber as well as ribbon pattern. This phenomenon is not observed in the system with only 2% ethanol/CH2Cl2 mixed solvents (Figure S4). The replacement of water by ethanol results in a further red shift from 1587 to 1562 cm−1 in the FTIR spectra (Figure S5a). The XRD pattern in the system with the ratio of water to ethanol of 1:9 shows a peak at 2θ of 5°, corresponding to a d spacing of 17.6 Å (Figure S5b). The result indicates that the thickness of the β-sheet monolayer in the gel phase is about 1.76 nm. The FL of microcrystals and fibers reveals the same emission peak, indicating that no π−π stacking structures form in these systems (Figure S5c). Therefore, it can be seen that the hydrogen-bond interactions play a decisive role in formation of fiber structures in our systems. Similar to ethanol, as both DMF and acetone can form hydrogen bonds with FF, the addition of a trace amount of DMF (and acetone) also can induce FF to form fiber and even ribbon structures (the results of DMF and acetone systems are shown in Figures S7−S10). In addition, two counter-examples also prove the important role of hydrogen bonding in fiber formation. We observed that the addition of a trace amount of toluene and n-hexane (both of them are not hydrogen-bondforming solvents) cannot induce the formation of fiber structures (Figures S11 and S12). The above observations together suggest that hydrogen-bonding (δh) interactions in the HSP parameters play a unique and key role in fiber formation, and these interactions can be manipulated by slight variations of solvent composition. Actually, many works have been reported to regulate and control the self-assembly and fiber formation of peptides,2,8−10,25,32−42 and these studies suggest that different kinds of interactions, such as hydrogen-bond, π−π, and hydrophobic interactions, can play an important role in fiber formation in certain self-assembly systems. However, in common biological systems, especially in Alzheimer’s amyloid fibril growth, the hydrogen-bonding interaction is essential.2 Moreover, it is interesting to note that the directional hydrogen bonding can promote the further growth and longrange-ordered arrangement of molecules in the original fibers
Figure 2. SEM images of FF/CH2Cl2/ethanol systems with ethanol contents of (a) 1%, (b) 2%, (c) 3%, and (d) 5%.
Figure 3. (a) FTIR spectra of FF/CH2Cl2/ethanol systems. (b) Optimized hydrogen-bonding structure between FF (line model) and ethanol (stick model). (c) XRD patterns and (d) FL spectra (λexcitation = 259 nm) of FF/CH2Cl2/ethanol systems with different ethanol contents.
clearly observed from the optimized structure of FF (line model) and ethanol (stick model) obtained by density functional theory (DFT) simulation in Figure 3b (the calculation method is listed in the Supporting Information). One can see that in ethanol abundant criss-crossed intermolecular hydrogen bonds (dashed cyan line) are remarkably increased by the hydroxyl groups. To determine whether the intermolecular interactions change during the solvent evaporation drying process, the IR spectra of dried samples were also measured. The results show that there are no discernible differences between the solution-based and the dried samples, except that the IR signals (around 1600 cm−1) of solution-based samples are weaker than those of dried samples (Figure S3). Therefore, it is conclusive to carry out X-ray diffraction (XRD) and fluorescence spectroscopy (FL) measurements in the dried state for further identifying intermolecular interactions. The XRD patterns (Figure 3c) of microcrystals formed in pure CH2Cl2 as well as in 1 and 2% ethanol/CH2Cl2 are similar to those of the hexagonal structure determined in a study of FF single crystals and nanotubes.29,30 If the content of ethanol is increased to 3%, the XRD patterns C
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ethanol, water, DMF, and acetone) can affect the bonding of CO and N−H in FF molecules. The directional hydrogen bonding can promote a long-range-ordered arrangement of FF molecules in one dimension, resulting in the formation of persistent and robust nanofibers as well as nanobelts. Our findings demonstrate the crucial role of solvent-bridged hydrogen bonding in terms of a trace amount of solvents in regulating biomolecular self-assembly, providing a reference for understanding the essential role of a small amount of solvents such as water for protein fibrillation and gene mutation, relative to various diseases, in complex biological systems.
to transform into nanobelts after being aged at room temperature for several days (Figure 4a−d). The FTIR
METHODS Materials. The dipeptide (L-Phe-L-Phe, FF) and 1,1,1,3,3,3hexafluoro-2-propanol (HFP) were purchased from Sigma. Dichloromethane (CH2Cl2) was obtained from Beijing InnoChem Science & Technology Co., Ltd. The water content in CH2Cl2 is less than 50 ppm. Ethanol, DMF, acetone, toluene, and n-hexane were purchased from Beijing Chemical Works (Beijing, China) and dried with molecular sieves before being used. Preparation of the Samples. The CH2 Cl2/CH3CH2OH mixtures with different ethanol contents were prepared by adding ethanol into CH2Cl2. Then, a freshly prepared FF/HFP solution (0.4 M, 20 μL) was diluted to a final concentration of 16 mM in CH2Cl2/ CH3CH2OH mixtures. The mixtures with other different trace solvents (DMF, acetone, toluene, or n-hexane) were prepared the same way. Microscopy Studies. Optical microscopy (Olympus IX73) was utilized to observe the samples in the solution. The SEM images were taken with a S-4800 scanning electron microscope, and the samples were allowed to dry by solvent evaporation, followed by sputtering a thin layer of gold. TEM images were recorded by using a JEM-2100 transmission electron microscope with samples placed onto the carbon-coated copper grids. AFM images were collected by FASTSCANBIO (Bruker) in a tapping mode. The samples were freeze-dried with the assistance of liquid nitrogen. Spectroscopy. FTIR spectra of the samples dried by solvent evaporation were measured by using a TENSOR 27 FTIR spectrometer (Bruker). The dried microcrystals were pressed into KBr pellets, and the gels were dried on a CaF2 plate. The solutionbased FTIR spectra were measured by using a Nicolet 380 FTIR spectrometer (Thermo, America), and the KBr windows were not pressed by KBr powder but by commercial and transparent disks with 4 mm thickness. The samples were put between two KBr windows to avoid solvent evaporation. The fluorescence was measured by a model FL-4500 spectrofluorometer (Hitachi, Tokyo, Japan), and the samples were excited at 259 nm. X-ray Diffraction. Data were collected at room temperature on a Empyrean (Panalytical, Nederland) instrument equipped with a Cu filter under the following conditions: scan step size, 0.026° 2θ; Cu Kα1 radiation (λ = 1.5406 Å). All samples were dried by solvent evaporation on a silica substrate. Density Functional Theory Simulation. See Supporting Information for details.
Figure 4. SEM images of FF/CH2Cl2/ethanol systems after (a) 1 day, (b) 3 days, (c) 5 days, and (d) 15 days. (e) FTIR spectra and (f) XRD patterns of FF/CH2Cl2/ethanol systems with different aging time. The amount of ethanol in CH2Cl2 is 3% (v/v).
absorption bands of the gel, aging after 5 days, in Figure 4e show that the peaks at 1587 cm−1 obviously blue shift back to about 1602 cm−1, revealing that the nanobelts have a crystalline structure. The results of XRD (Figure 4f) also agree with the FTIR observations, as one can see that the XRD patterns change and the signals of hexagonal structures appear. Taken together, these results indicate that, at short-range, higher ethanol content can induce formation of a fibrous gel with homogeneity but recrystallizes into hierarchically organized anisotropic nanobelts at long times. On the mesoscale, the morphological transition from metastable gels to crystallized nanobelts is governed by the growth kinetics. Nevertheless, the gels consisting of nanobelts remain stable at least for half of a month. Therefore, apparently, a trace amount of solvents, specifically interacting with peptide molecules, plays a key role in tuning the molecular organization and the finally formed structures.
ASSOCIATED CONTENT S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsnano.5b06567. Figures S1−S12, Table S1, and the method of DFT simulation (PDF)
CONCLUSIONS In conclusion, we have discovered that a trace amount of solvents can be a predominant factor to tune dipeptide selfassembly. It has been demonstrated that bridged hydrogen bonding in the process of dipeptide self-assembly plays a crucial role in mediating long-range-ordered fiber formation, whereas the other noncovalent interactions, including π−π and van der Waals interactions, are not key factors for long-range growth. The addition of hydrogen-bond-forming solvents (including
AUTHOR INFORMATION Corresponding Author
*E-mail:
[email protected]. Notes
The authors declare no competing financial interest. D
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(18) Ma, H. C.; Fei, J. B.; Li, Q.; Li, J. B. Photo-Induced Reversible Structural Transition of Cationic Diphenylalanine Peptide SelfAssembly. Small 2015, 11, 1787−1791. (19) Liu, X. C.; Zhu, P. L.; Fei, J. B.; Zhao, J.; Yan, X. H.; Li, J. B. Synthesis of Peptide-Based Hybrid Nanobelts with Enhanced Color Emission by Heat Treatment or Water Induction. Chem. - Eur. J. 2015, 21, 9461−9467. (20) Yan, X.; Li, J.; Möhwald, H. Self-Assembly of Hexagonal Peptide Microtubes and Their Optical Waveguiding. Adv. Mater. 2011, 23, 2796−2801. (21) Yan, X.; Su, Y.; Li, J.; Frueh, J.; Möhwald, H. Uniaxially Oriented Peptide Crystals for Active Optical Waveguiding. Angew. Chem., Int. Ed. 2011, 50, 11186−11191. (22) Yan, X.; Zhu, P.; Li, J. Self-Assembly and Application of Diphenylalanine-Based Nanostructures. Chem. Soc. Rev. 2010, 39, 1877−1890. (23) Adler-Abramovich, L.; Gazit, E. The Physical Properties of Supramolecular Peptide Assemblies: From Building Block Association to Technological Applications. Chem. Soc. Rev. 2014, 43, 6881−6893. (24) Burgess, N. C.; Sharp, T. H.; Thomas, F.; Wood, C. W.; Thomson, A. R.; Zaccai, N. R.; Brady, R. L.; Serpell, L. C.; Woolfson, D. N. Modular Design of Self-Assembling Peptide-Based Nanotubes. J. Am. Chem. Soc. 2015, 137, 10554−10562. (25) Zhu, P.; Yan, X.; Su, Y.; Yang, Y.; Li, J. Solvent-Induced Structural Transition of Self-Assembled Dipeptide: From Organogels to Microcrystals. Chem. - Eur. J. 2010, 16, 3176−3183. (26) Lan, Y.; Corradini, M.; Weiss, R.; Raghavan, S.; Rogers, M. To Gel or Not To Gel: Correlating Molecular Gelation with Solvent Parameters. Chem. Soc. Rev. 2015, 44, 6035−6058. (27) Lamm, M. S.; Rajagopal, K.; Schneider, J. P.; Pochan, D. J. Laminated Morphology of Nontwisting Beta-Sheet Fibrils Constructed via Peptide Self-Assembly. J. Am. Chem. Soc. 2005, 127, 16692−16700. (28) Elfrink, K.; Ollesch, J.; Stoehr, J.; Willbold, D.; Riesner, D.; Gerwert, K. Structural Changes of Membrane-Anchored Native PrPC. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 10815−10819. (29) Gö rbitz, C. H. Nanotube Formation by Hydrophobic Dipeptides. Chem. - Eur. J. 2001, 7, 5153−5159. (30) Görbitz, C. H. The Structure of Nanotubes Formed by Diphenylalanine, the Core Recognition Motif of Alzheimer’s βAmyloid Polypeptide. Chem. Commun. 2006, 2332−2334. (31) Gadde, S.; Batchelor, E. K.; Weiss, J. P.; Ling, Y.; Kaifer, A. E. Control of H- and J-Aggregate Formation via Host-Guest Complexation using Cucurbituril Hosts. J. Am. Chem. Soc. 2008, 130, 17114− 17119. (32) Huang, R.; Qi, W.; Su, R.; Zhao, J.; He, Z. Solvent and Surface Controlled Self-Assembly of Diphenylalanine Peptide: From Microtubes to Nanofibers. Soft Matter 2011, 7, 6418−6421. (33) Wang, Y.; Qi, W.; Huang, R.; Yang, X.; Wang, M.; Su, R.; He, Z. Rational Design of Chiral Nanostructures from Self-Assembly of a Ferrocene-Modified Dipeptide. J. Am. Chem. Soc. 2015, 137, 7869− 7880. (34) Cui, H.; Muraoka, T.; Cheetham, A. G.; Stupp, S. I. SelfAssembly of Giant Peptide Nanobelts. Nano Lett. 2009, 9, 945−951. (35) Vidyasagar, A.; Sureshan, K. M. Stoichiometric Sensing to Opt between Gelation and Crystallization. Angew. Chem., Int. Ed. 2015, 54, 12078−12082. (36) Ma, M.; Kuang, Y.; Gao, Y.; Zhang, Y.; Gao, P.; Xu, B. AromaticAromatic Interactions Induce the Self-Assembly of Pentapeptidic Derivatives in Water To Form Nanofibers and Supramolecular Hydrogels. J. Am. Chem. Soc. 2010, 132, 2719−2728. (37) Tsonchev, S.; Niece, K. L.; Schatz, G. C.; Ratner, M. A.; Stupp, S. I. Phase Diagram for Assembly of Biologically-Active Peptide Amphiphiles. J. Phys. Chem. B 2008, 112, 441−447. (38) Fu, I. W.; Markegard, C. B.; Nguyen, H. D. Solvent Effects on Kinetic Mechanisms of Self-Assembly by Peptide Amphiphiles via Molecular Dynamics Simulations. Langmuir 2015, 31, 315−324. (39) Draper, E. R.; Eden, E. G.; McDonald, T. O.; Adams, D. J. Spatially Resolved Multicomponent Gels. Nat. Chem. 2015, 7, 848− 852.
ACKNOWLEDGMENTS We acknowledge financial support from the National Natural Science Foundation of China (Project Nos. 21522307, 21473208, and 91434103), the Talent Fund of the Recruitment Program of Global Youth Experts, the CAS visiting professorships for senior international scientists (Project No. 2015VTA033), and the Chinese Academy of Sciences (CAS). Prof. Yanqiang Zhang is thanked for his assistance in solutionbased IR measurement. The first two authors contributed equally to this work. REFERENCES (1) Krone, M. G.; Hua, L.; Soto, P.; Zhou, R.; Berne, B. J.; Shea, J.-E. Role of Water in Mediating the Assembly of Alzheimer Amyloid-β Aβ16−22 Protofilaments. J. Am. Chem. Soc. 2008, 130, 11066−11072. (2) Knowles, T. P.; Fitzpatrick, A. W.; Meehan, S.; Mott, H. R.; Vendruscolo, M.; Dobson, C. M.; Welland, M. E. Role of Intermolecular Forces in Defining Material Properties of Protein Nanofibrils. Science 2007, 318, 1900−1903. (3) Whitesides, G. M.; Grzybowski, B. Self-Assembly at All Scales. Science 2002, 295, 2418−2421. (4) Reches, M.; Gazit, E. Casting Metal Nanowires within Discrete Self-Assembled Peptide Nanotubes. Science 2003, 300, 625−627. (5) Hamley, I. W. Peptide Nanotubes. Angew. Chem., Int. Ed. 2014, 53, 6866−6881. (6) Kim, J.; Han, T. H.; Kim, Y.-I.; Park, J. S.; Choi, J.; Churchill, D. C.; Kim, S. O.; Ihee, H. Role of Water in Directing Diphenylalanine Assembly into Nanotubes and Nanowires. Adv. Mater. 2010, 22, 583− 587. (7) Zou, Q.; Liu, K.; Abbas, M.; Yan, X. Peptide-Modulated SelfAssembly of Chromophores towards Biomimetic Light-Harvesting Nanoarchitectonics. Adv. Mater. 2015, DOI: 10.1002/ adma.201502454. (8) Korevaar, P. A.; Newcomb, C. J.; Meijer, E. W.; Stupp, S. I. Pathway Selection in Peptide Amphiphile Assembly. J. Am. Chem. Soc. 2014, 136, 8540−8543. (9) Mason, T. O.; Chirgadze, D. Y.; Levin, A.; Adler-Abramovich, L.; Gazit, E.; Knowles, T. P. J.; Buell, A. K. Expanding the Solvent Chemical Space for Self-Assembly of Dipeptide Nanostructures. ACS Nano 2014, 8, 1243−1253. (10) Tang, C.; Smith, A. M.; Collins, R. F.; Ulijn, R. V.; Saiani, A. Fmoc-Diphenylalanine Self-Assembly Mechanism Induces Apparent pK(a) Shifts. Langmuir 2009, 25, 9447−9453. (11) Yan, X.; Cui, Y.; He, Q.; Wang, K.; Li, J.; Mu, W.; Wang, B.; Ouyang, Z.-c. Reversible Transitions between Peptide Nanotubes and Vesicle-Like Structures Including Theoretical Modeling Studies. Chem. - Eur. J. 2008, 14, 5974−5980. (12) Fleming, S.; Ulijn, R. V. Design of Nanostructures Based on Aromatic Peptide Amphiphiles. Chem. Soc. Rev. 2014, 43, 8150−8177. (13) Guo, C.; Luo, Y.; Zhou, R.; Wei, G. Probing the Self-Assembly Mechanism of Diphenylalanine-Based Peptide Nanovesicles and Nanotubes. ACS Nano 2012, 6, 3907−3918. (14) Jeon, J.; Mills, C. E.; Shell, M. S. Molecular Insights into Diphenylalanine Nanotube Assembly: All-Atom Simulations of Oligomerization. J. Phys. Chem. B 2013, 117, 3935−3943. (15) Tamamis, P.; Adler-Abramovich, L.; Reches, M.; Marshall, K.; Sikorski, P.; Serpell, L.; Gazit, E.; Archontis, G. Self-Assembly of Phenylalanine Oligopeptides: Insights from Experiments and Simulations. Biophys. J. 2009, 96, 5020−5029. (16) Liu, K.; Xing, R.; Chen, C.; Shen, G.; Yan, L.; Zou, Q.; Ma, G.; Möhwald, H.; Yan, X. Peptide-Induced Hierarchical Long-Range Order and Photocatalytic Activity of Porphyrin Assemblies. Angew. Chem., Int. Ed. 2015, 54, 500−505. (17) Zhang, H.; Fei, J. B.; Yan, X. H.; Wang, A. H.; Li, J. B. EnzymeResponsive Release of Doxorubicin from Monodisperse DipeptideBased Nanocarriers for Highly Efficient Cancer Treatment In Vitro. Adv. Funct. Mater. 2015, 25, 1193−1204. E
DOI: 10.1021/acsnano.5b06567 ACS Nano XXXX, XXX, XXX−XXX
Article
ACS Nano (40) Kumar, V. A.; Shi, S.; Wang, B. K.; Li, I.-C.; Jalan, A. A.; Sarkar, B.; Wickremasinghe, N. C.; Hartgerink, J. D. Drug-Triggered and Cross-Linked Self-Assembling Nanofibrous Hydrogels. J. Am. Chem. Soc. 2015, 137, 4823−4830. (41) Löwik, D. W.; Leunissen, E.; Van den Heuvel, M.; Hansen, M.; van Hest, J. C. Stimulus Responsive Peptide Based Materials. Chem. Soc. Rev. 2010, 39, 3394−3412. (42) Ramakers, B.; van Hest, J.; Löwik, D. Molecular Tools for the Construction of Peptide-Based Materials. Chem. Soc. Rev. 2014, 43, 2743−2756.
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