Disassembly of Dipeptide Single Crystals Can Transform the Lipid

Disassembly of Dipeptide Single Crystals Can Transform the Lipid Membrane into a Network Meifang Fu,†,§ Qi Li,†,§ Bingbing Sun,†,§ Yang Yang,‡ Luru Dai,‡ Tommy Nylander,∥ and Junbai Li*,†,§ †

Beijing National Laboratory for Molecular Sciences (BNLMS), CAS Key Lab of Colloid, Interface and Thermodynamics, Institute of Chemistry, Chinese Academy of Sciences, 100190 Beijing, China ‡ CAS Key Laboratory for Biomedical Effects of Nanomaterials and Nanosafety, National Center for Nanoscience and Technology, 100190 Beijing, China § University of Chinese Academy of Sciences, 100049 Beijing, China ∥ Division of Physical Chemistry, Department of Chemistry, Lund University, P.O. Box 124, SE-22100 Lund, Sweden S Supporting Information *

ABSTRACT: Coupling between cytoskeleton and membranes is critical to cell movement as well as organelle formation. Here, we demonstrate that self-assembled single crystals of a dipeptide, diphenylalanine (FF), can interact with liposomes to form cytoskeleton-like structures. Under a physiological condition, disassembly of FF crystals deforms and translocates supported lipid membrane. The system exhibits similar dynamic characteristics to the endoplasmic reticulum (ER) network in cells. This bottom-up system thus indicates that external matter can participate in the deformation of liposomes, and disassembly of the nanostructures enables a system with distinct dynamic behaviors. KEYWORDS: phospholipids, self-assembly, peptides, membranes, biomimetic

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translocated accordingly. Membrane network construction in this system shares similar dynamic characteristics to ER in living cells.

oupling between cytoskeleton and membranes is critical for cellular responses to the external environment as well as intracellular organelle structures, such as Golgi apparatus and endoplasmic reticulum (ER).1−6 Cytoskeleton is composed mainly of three classes of protein fibers: actin, microtubules (MTs), and intermediate filaments. An important function of the cytoskeleton in the cytoskeleton− membrane coupling is to provide the force required for membrane movement.2 Both actin and MTs can generate force by their polymerization and depolymerization dynamics.2 Development of molecular self-assembly makes many materials with defined structures and distinct properties available.7−9 Mimicking critical cell functions with molecular materials may give us information about these functions and provide strategies to construct functional systems.10−14 Diphenylalanine (FF) dipeptide is the core recognition motif of Alzheimer’s β-amyloid polypeptide15 and can readily form various structures such as nanotubes, microtubes, and nanowires.16,17 FF assembly has been shown to display a relatively high mechanical strength;18 however, they dissolve easily in several solvents.19,20 Here, we take advantage of this property and combine FF assemblies with liposomes. Fusion of liposomes on FF assembly induces supported lipid membranes (FF−lipid complexes). When FF assembly dissolves in different buffer conditions, the supported membranes are deformed and © 2017 American Chemical Society

RESULTS AND DISCUSSION FF assembly is fabricated with single-crystalline structures (Figure 1A).17 Its XRD pattern is similar to that of the hexagonal FF single crystal structures (Figure 1B).21 Polarized optical microscope images demonstrate the birefringence property of the FF single crystals (Figure S1A). Dissolution of FF crystals in cell culture medium (Dulbecco’s modified Eagle medium (DMEM) with calf serum (CS)) is observed to ensure they dissolve completely with time (Movie S1). Supported phospholipid membranes are formed via the fusion of liposomes on FF crystals.22,23 When an aliquot of liposome dispersion is added onto a coverslip with FF crystals on it, the liposomes spread on the FF crystals immediately with some of the crystals disintegrating (Figure 1C). SEM and TEM images of FF−lipid complexes indicate the existence of lipid membranes (Figure 1D). There is no change in the XRD pattern of the FF−lipid complexes (Figure S1B), indicating that Received: May 18, 2017 Accepted: June 28, 2017 Published: June 28, 2017 7349

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Figure 1. Fabrication of FF crystal supported phospholipid membrane. (A) SEM image of FF assembly. (B) XRD pattern of FF assembly. The inset image is molecular structure of FF. (C) The adhesion of liposomes on FF crystals. The images are overlapped images of confocal fluorescence images of liposomes (red) and corresponding bright field images of FF crystals. (D) SEM image of FF−lipid complexes. The inset image is the TEM image of FF−lipid complexes. The red arrow indicates the lipid membrane on FF crystals.

Figure 2. Deformation of supported membranes into vesicles in cell culture medium. (A) Confocal images indicate vesicles form accompanying FF crystal disassembly. (B) Dual color confocal fluorescence images show that the lipid membrane (Rhod-PE, red) covers the FF crystal (FITC, cyan), and vesicles form with FF crystal disassembly. White arrows indicate the bulges on the edge of the final vesicle.

two vesicles on a FF crystal, the vesicles can adjust their orientations, and the movement induces their fusion (Figure S3C). In an effort to decrease the complexity of the system, CS is removed from cell culture medium. FF−lipid complexes in DMEM tend to deposit on the coverslip, and a membrane network is generated beneath the vesicles (Figure 3A). This may be induced by the coverslip that provides support for tubule extension. We confirm this by using phosphate buffered solution (PBS), another buffer used in cell culture, and CS (5%)−PBS. The results are consistent with those observed in systems containing DMEM. Supported membranes are deformed into vesicles in CS-PBS, while they tend to form tubular structures in PBS (Figure S4). After decreasing the lipid amount, tubular structures and networks become more clear (Figure 3B). We then analyze the formation mechanisms of tubular structures and networks separately. Dissolution of isolated FF crystals in DMEM generates membrane tubular structures (Movie S3). Isolated FF−lipid complexes and their corresponding bright field image (shown

the lipid membranes do not change FF crystal structure. Fourier transform infrared spectroscopy (FTIR) images of FF crystal and FF−lipid complexes both show the characteristic peaks at 1606 and 1687 cm−1, which correspond to the antiparallel β-sheet structure of the FF crystal (Figure S2).17 FF−lipid complexes are then immersed in cell culture medium. It is indicated that the supported membranes are deformed into vesicles accompanying FF crystal dissolution (Figure 2A, Movie S2). Dual color imaging is conducted where FF is coassembled with FITC (cyan) and phospholipid is coassembled with Rhod PE (red) (Figure 2B). The lipid membrane is gradually deformed into a vesicle with the dissolution of FF crystal and the fluorescence signal from the cyan channel disappears gradually. FITC does not associate with FF molecules through covalent bonds, so the fluorescence signals from FITC only indirectly reflect FF crystal behavior. There are two bulges at the edge of the final vesicle, which may be induced by the interference from dissolved FF molecules (see also Figure S3A). During this process, disassembly of FF crystals induces vesicles movement (Figure S3A,B). If there are 7350

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restrained state (Figure 4C, double-headed arrow, left). The restrained membrane appears to expand (Figure 4D, doubleheaded arrow) accompanying the forward motion of the moving membrane. After the moving membrane reaches the farthest location (Figure 4D), it retreats to the location shown in Figure 4E (the full line between Figure 4D,E). At the same time, the restrained membrane swells further (Figure 4E, double-headed arrows). The moving membrane continues to retreat by rolling over, pouring the phospholipid constituents into the left chamber rapidly (Figure 4F). The tubular structure undergoes several morphological changes (Figure 4G, H) before it reaches equilibrium (Figure 4I). We show two other typical structures generated by this system in Figure S5 that share similar formation process. The fluorescence intensities are heterogeneous whether in the intermediate or in the final structures (see also Figure S6A). The heterogeneous fluorescence as well as the separation of the membranes (Figure 4B) may reflect heterogeneous organization of the membrane. FF crystals can be seen as closepacking of hydrophobic tubes with hydrophobic side chains emanating from the hydrophilic core.21 The hydrophobic surface may associate with phospholipid tails, inducing membrane monolayer on it.24,25 After depositing FF−lipid complexes onto a coverslip, we find that excess membranes spill out (Figure S6B). A similar phenomenon has been observed when supported bilayers were prepared in high salt solution and then washed with water.26 This system has been found to possess a large fluid reservoir of a single bilayer.27 In the current study, the exceeded membranes exhibit different fluorescence intensities (Figure S6B). When phospholipid/FF molar ratio is increased from 0.004 to 0.03, the exceeded membranes form two layers with different fluorescence intensities (Figure S6C,D). This finding may demonstrate the presence of multibilayers on the surface of the FF crystal. Some vesicles formed in cell culture medium demonstrate multilayers (Figure S7), which is consistent with the exceeded membranes. Based on the observation in Figure 4, we speculate that the fluid bilayer reservoir will be compressed as the FF crystal dissolves in DMEM,28 while the monolayer will associate with the remaining FF assembly and adopt a constrained form. When FF−lipid complexes cross-link one another, dissolution of FF crystals deforms supported membranes into networks in DMEM (Movie S4). The FF−lipid complex disintegrates above other membrane tubes, linking the tubes and creating tube junctions (Figure 5K,L, marked with the cyan dotted line). Every element in the interacted FF−lipid complexes still follows the pattern described above. However, the interactions among the complexes add other dynamic feature. One feature is the redistribution of lipid constituents in the range of the whole network (Figure 5K,L, indicated with cyan arrows). Another feature is the translocation of membrane tubes. Translocation of membrane tubules is an important factor in the construction of ER tubule networks in cells. Ring closure that makes the ER less densely reticulated is based on the sliding of one tubule along another at the point where they are joined.29 This behavior has also been observed in an in vitro system where the force generated by kinesin upon the MTs produced the polygonal networks. The author attributed the translocation to the elastic responses of lipid membrane to tension and its fluid property.30 After a close check of the network construction process, we conclude that there are three main translocation methods for

Figure 3. Membrane network formation in DMEM. (A) FF−lipid complexes (phospholipid/FF molar ratio is 0.02) dissolve into lipid vesicles. A magnified image shows that a membrane network is beneath the vesicles. (B) After lipid amount is decreased (phospholipid/FF molar ratio is 0.004), membrane tubular structures and network are more clear.

in the right column) are shown in Figure 4A. As they dissemble, the FF crystals leave part of immobile restrained membrane

Figure 4. Formation of membrane tubular structure through dissolution of isolated FF crystals in DMEM.

(Figure 4B, dotted lines). The left side FF crystal leaves a knot in addition (Figure 4B, arrow), where the membranes begin to move separately. While one part of the membrane continues move forward (the full line between Figure 4B−D), another part of the membrane is left on the pathway and adopts a 7351

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Figure 5. Construction of membrane network through dissolution of cross-linked FF crystals in DMEM. The scale bars in inset images represent 2 μm.

tubular structures with multichambers. This asymmetry structure forms through controlled shape shifts of the lipid membrane, which is reminiscent of the concept of supramolecular differentiation. In the pioneering example, tube-like protuberances spontaneously formed on the surface of spherical particles.37 Once the experimental conditions are decided, the asymmetry structures will form spontaneously and depend on the inherent molecular interactions. In the system presented here, the construction process is solely controlled by the dissolution of FF crystals without the involvement of other external forces. Supramolecular systems that have the differentiation characteristics will not only benefit the material science but also may enable us to better understand biosystems.

the membrane tubules. As shown in Figure 5A,B, the complex indicated by the cyan ellipse moves upward along another complex and the membrane tube indicated by the cyan line is pulled up. At the same time, the lateral tension in the membrane tube builds up to the maximum, after which it relaxes and drags the complex down (Figure 5B,C). Other cases of this method can be found in Figure 5G−I where the membrane tubes marked with dotted lines move with the remaining complex. Comparing the inset images in Figure 5A,F and D,E, we find that the membrane tubes are translocated via fusion of phospholipid vesicles or tubes. Translocation via fusion is fast and occurs within 5 s in the current imaging condition. It has been shown that minimization of surface free energy optimizes the layout of nanotube−vesicle networks.31 The inset images in Figure 5J,K show that the membrane tube (the dotted line) translocates and forms a more stable symmetrical three-way junction (Figure 5L, white arrow) instead of maintaining the original cross-intersection (Figure 5J, inset image). Amphiphilic molecules, especially lipids, can self-assemble into a wide range of morphologies, and their shapes can be changed through molecular modification or environmental stimulus.32,33 For example, it has been shown that lipid vesicles can exhibit the extrusion and contraction of pseudopod-like structures under a pH gradient.33 The lipid morphology change here is controlled by the dissolution of dipeptide assembly. Compared with relatively simple and uniform tubes formed from lipid only,34−36 our system constructs the asymmetry

CONCLUSIONS In conclusion, we demonstrate that disassembly of supramolecular dipeptide single crystals at a physiological condition induces the deformation and translocation of liposomes. This character is similar to the cytoskeleton−membrane coupling in cells, where disassembly of the cytoskeleton can provide the force required for membrane movement. Liposomes here are deformed into vesicles, tubular structures, or networks that depend on buffer conditions. During membrane network construction, membrane tubules can be translocated through different means. This dynamic behavior is reminiscent of ER in living cells, in which tubule translocation is an important factor to construct the ER morphology. This study thus indicates that the external matter can participate in the deformation of 7352

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ACS Nano liposomes, and disassembly of the nanostructures enables a system with distinct dynamic behaviors.

vesicles and vesicle fusion induced by dissolution of FF crystals, images of FF−lipid complexes in PBS-CS and PBS, membrane tubular structures induced by disassembly of FF single crystals in DMEM, fluorescent images and profiles of FF−lipid complexes at different molar ratios, and multilayer properties of the vesicles (PDF) Dissolution of FF crystals in cell culture medium (AVI) Lipid membranes are deformed into vesicles in cell culture medium (AVI) Lipid membranes are deformed into tubular structures in DMEM (AVI) Lipid membranes are deformed into networks in DMEM (AVI)

EXPERIMENTAL SECTION Construction of FF Crystals. The FF crystals were constructed with a method described previously.17 FF (3 mg. Sigma-Aldrich, purity by thin layer chromatography, ≥98%, CAS 2577-40-4) was dissolved in 45 μL of 1,1,3,3,6,6-hexafluoro-2-propanol (HFP), and then 105 μL of water was poured into the solution, followed by ultrasonic treatment at room temperature for 5 or 10 min. For the FITC labeled FF crystals, FITC was dissolved in HFP (1 mg/mL), and the synthesis procedure was unchanged. The FF crystal solution was allowed to stand for 12 h. Then the HFP was removed via vacuum drying. Small Unilamellar Vesicle Preparation. 1,2-Dioleoyl-sn-glycero3-phosphoethanolamine (DOPE, 20 mol %, Avanti, >99%, CAS: 400405-1) in 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC, Avanti, >99%, CAS: 4235-95-4) with 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (Rhod PE, Avanti, >99%, CAS: 384833-00-5) (typically 0.5 mol %) were dissolved in chloroform. Chloroform was evaporated under nitrogen flow, followed by vacuum for 6 h. The dry lipid film was rehydrated in PBS (Solarbio, Cat. no. P1020-500) (to a final concentration of 1 mM), vortexed, and left to stand for 30 min. Lipid dispersions were sonicated to clarity (9.9 s on, 9.9 s off for 20 min) using an Ultrasonic Cell Disruptor (SCIENTZ-IID, Ningbo Scientz Biotechnology CO.LTD, Newton, CT). Fabrication of FF Crystal Supported Membrane. FF crystal supported phospholipid membranes were formed by fusion of phospholipid vesicles.22 The dispersions of the lipid vesicles with different volumes and the corresponding equal volume of water were mixed with dried FF nanorods (NRs), and the complexes were vortexed for 30 min. Varied phospholipid/FF molar ratios (0.004− 0.03) were used. The excess vesicles were removed via centrifugation (10000 rpm, 1 min), and phosphate buffered solution (PBS) was exchanged with water 3 times. Dissolution of FF Crystals. FF crystals and the centrifuged FF− lipid complexes were mixed with either CS (5%, Gibco, Cat. no. 16010-159)−DMEM/high glucose (HyClone, Cat. no. SH30022.01) or DMEM/high glucose (400 μL) via brief vortex. Then the solutions were transferred into tissue culture dishes with glass bottoms. The volumes of PBS and CS (5%)-PBS were 1800 μL. XRD and FTIR Characterization. X-ray diffraction (XRD) patterns were measured at 25 °C using Rigaku. Data were collected on a Rigaku D/max 2500 instrument equipped with a Cu filter under the following conditions: scan speed, 2°/min; Cu Kα radiation, λ = 1.5405 Å. Fourier transform infrared spectroscopy (FTIR) was carried out with a TENSOR-27 in the range of 4000−400 cm−1. Fluorescence and Electron Microscope Observation. The FF NRs and the centrifuged FF−lipid complexes were characterized by using scanning electronic microscope (SEM, S-4800, Hitachi, Tokyo, Japan), transmission electron microscope (TEM, JEM-1011), polarizing microscope (POM, Leica DMLP), confocal fluorescent microscope (Olympus), and fluorescence microscope (home-built). Images generated by the confocal microscope are pseudocolored. Our custom built setup is based on an Olympus IX-81 inverted microscope body (Olympus) with a 150×, 1.45 NA oil immersion objective (PlanApoN, Olympus). It is equipped with an EMCCD (electron multiplying charge coupled device), and the frequency used was 26.89 Hz (37 ms/ frame). A 561 nm diode laser (Sapphire 561-200 CW; Coherent) was used for excitation.

AUTHOR INFORMATION Corresponding Author

*[email protected]. ORCID

Tommy Nylander: 0000-0001-9420-2217 Junbai Li: 0000-0001-9575-3125 Notes

The authors declare no competing financial interest.

ACKNOWLEDGMENTS The authors acknowledge the financial support from the National Natural Science Foundation of China (Project Nos. 21433010, 21320102004, and 21673056) and National Basic Research Program of China (973 program, 2013CB932802). REFERENCES (1) Raucher, D.; Stauffer, T.; Chen, W.; Shen, K.; Guo, S.; York, J. D.; Sheetz, M. P.; Meyer, T. Phosphatidylinositol 4,5-Bisphosphate Functions as a Second Messenger that Regulates Cytoskeleton-Plasma Membrane Adhesion. Cell 2000, 100, 221−228. (2) Gurel, P. S.; Hatch, A. L.; Higgs, H. N. Connecting the Cytoskeleton to the Endoplasmic Reticulum and Golgi. Curr. Biol. 2014, 24, R660−672. (3) Stephens, D. J. Functional Coupling of Microtubules to Membranes-Implications for Membrane Structure and Dynamics. J. Cell Sci. 2012, 125, 2795−2804. (4) Koster, D. V.; Mayor, S. Cortical Actin and the Plasma Membrane: Inextricably Intertwined. Curr. Opin. Cell Biol. 2016, 38, 81−89. (5) Sheetz, M. P.; Sable, J. E.; Dobereiner, H.-G. Continuous Membrane-Cytoskeleton Adhesion Requires Continuous Accommodation to Lipid and Cytoskeleton Dynamics. Annu. Rev. Biophys. Biomol. Struct. 2006, 35, 417−434. (6) Sheetz, M. P. Cell Control by Membrane-Cytoskeleton Adhesion. Nat. Rev. Mol. Cell Biol. 2001, 2, 392−396. (7) He, Q.; Cui, Y.; Li, J. Molecular Assembly and Application of Biomimetic Microcapsules. Chem. Soc. Rev. 2009, 38, 2292−2303. (8) Yan, X.; Zhu, P.; Li, J. Self-Assembly and Application of Diphenylalanine-Based Nanostructures. Chem. Soc. Rev. 2010, 39, 1877−1890. (9) Ariga, K.; Li, J.; Fei, J.; Ji, Q.; Hill, J. P. Nanoarchitectonics for Dynamic Functional Materials from Atomic-/Molecular-Level Manipulation to Macroscopic Action. Adv. Mater. 2016, 28, 1251−1286. (10) Tu, Y.; Peng, F.; Adawy, A.; Men, Y.; Abdelmohsen, L. K.; Wilson, D. A. Mimicking the Cell: Bio-Inspired Functions of Supramolecular Assemblies. Chem. Rev. 2016, 116, 2023−2078. (11) Liu, A. P.; Fletcher, D. A. Biology Under Construction: In Vitro Reconstitution of Cellular Function. Nat. Rev. Mol. Cell Biol. 2009, 10, 644−650.

ASSOCIATED CONTENT S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsnano.7b03468. POM images of the FF assembly and XRD pattern of FFlipid complexes, FTIR images of FF crystal and FF-lipid complexes, bright field images showing movement of the 7353

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ACS Nano (12) Koga, S.; Williams, D. S.; Perriman, A. W.; Mann, S. PeptideNucleotide Microdroplets as a Step Towards a Membrane-Free Protocell Model. Nat. Chem. 2011, 3, 720−724. (13) Liu, K.; Xing, R.; Li, Y.; Zou, Q.; Mohwald, H.; Yan, X. Mimicking Primitive Photobacteria: Sustainable Hydrogen Evolution Based on Peptide-Porphyrin Co-Assemblies with a Self-Mineralized Reaction Center. Angew. Chem., Int. Ed. 2016, 55, 12503−12507. (14) Schwille, P. Bottom-Up Synthetic Biology: Engineering in a Tinkerer’s World. Science 2011, 333, 1252−1254. (15) Reches, M.; Gazit, E. Casting Metal Nanowires Within Discrete Self-Assembled Peptide Nanotubes. Science 2003, 300, 625−627. (16) 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. (17) Li, Q.; Jia, Y.; Dai, L.; Yang, Y.; Li, J. Controlled Rod Nanostructured Assembly of Diphenylalanine and Their Optical Waveguide Properties. ACS Nano 2015, 9, 2689−2695. (18) Kol, N.; Adler-Abramovich, L.; Barlam, D.; Shneck, R. Z.; Gazit, E.; Rousso, I. Self-Assembled Peptide Nanotubes are Uniquely Rigid Bioinspired Supramolecular Structures. Nano Lett. 2005, 5, 1343− 1346. (19) Andersen, K. B.; Castillo-Leon, J.; Hedstrom, M.; Svendsen, W. E. Stability of Diphenylalanine Peptide Nanotubes in Solution. Nanoscale 2011, 3, 994−998. (20) Nezammahalleh, H.; Amoabediny, G.; Kashanian, F.; Foroughi Moghaddam, M. H. An Investigation on the Chemical Stability and a Novel Strategy for Long-Term Stabilization of Diphenylalanine Nanostructures in Aqueous Solution. Results Phys. 2015, 5, 11−19. (21) Gorbitz, C. H. The Structure of Nanotubes Formed by Diphenylalanine, the Core Recognition Motif of Alzheimer’s BetaAmyloid Polypeptide. Chem. Commun. 2006, 2332−2334. (22) Dabkowska, A. P.; Niman, C. S.; Piret, G.; Persson, H.; Wacklin, H. P.; Linke, H.; Prinz, C. N.; Nylander, T. Fluid and Highly Curved Model Membranes on Vertical Nanowire Arrays. Nano Lett. 2014, 14, 4286−4292. (23) Reimhult, E.; Höök, F.; Kasemo, B. Intact Vesicle Adsorption and Supported Biomembrane Formation from Vesicles in Solution: Influence of Surface Chemistry, Vesicle Size, Temperature, and Osmotic Pressure. Langmuir 2003, 19, 1681−1691. (24) Czolkos, I.; Jesorka, A.; Orwar, O. Molecular Phospholipid Films on Solid Supports. Soft Matter 2011, 7, 4562−4576. (25) Wang, S. T.; Fukuto, M.; Yang, L. In Situ X-Ray Reflectivity Studies on the Formation of Substrate-Supported Phospholipid Bilayers and Monolayers. Phys. Rev. E 2008, 77, 031909. (26) Pucadyil, T. J.; Schmid, S. L. Real-Time Visualization of Dynamin-Catalyzed Membrane Fission and Vesicle Release. Cell 2008, 135, 1263−1275. (27) Pucadyil, T. J.; Schmid, S. L. Supported Bilayers with Excess Membrane Reservoir: a Template for Reconstituting Membrane Budding and Fission. Biophys. J. 2010, 99, 517−525. (28) Nissen, J.; Jacobs, K.; Radler, J. O. Interface Dynamics of Lipid Membrane Spreading on Solid Surfaces. Phys. Rev. Lett. 2001, 86, 1904−1907. (29) Lee, C.; Chen, L. B. Dynamic Behavior of Endoplasmic Reticulum in Living Cells. Cell 1988, 54, 37−46. (30) Vale, R. D.; Hotani, H. Formation of Membrane Networks In Vitro by Kinesin-Driven Microtubule Movement. J. Cell Biol. 1988, 107, 2233−2241. (31) Lobovkina, T.; Dommersnes, P. G.; Tiourine, S.; Joanny, J. F.; Orwar, O. Shape Optimization in Lipid Nanotube Networks. Eur. Phys. J. E: Soft Matter Biol. Phys. 2008, 26, 295−300. (32) Kim, C. J.; Kurauchi, S.; Uebayashi, T.; Fujisaki, A.; Kimura, S. Morphology Change from Nanotube to Vesicle and Monolayer/ Bilayer Alteration by Amphiphilic Block Polypeptides Having Aromatic Groups at C Terminal. Bull. Chem. Soc. Jpn. 2017, 90, 568−573. (33) Nawa, E.; Yamamoto, D.; Shioi, A. Chemotactic Amoeboid-Like Shape Change of a Vesicle under a pH Gradient. Bull. Chem. Soc. Jpn. 2015, 88, 1536−1544.

(34) Jesorka, A.; Stepanyants, N.; Zhang, H.; Ortmen, B.; Hakonen, B.; Orwar, O. Generation of Phospholipid Vesicle-Nanotube Networks and Transport of Molecules Therein. Nat. Protoc. 2011, 6, 791−805. (35) Paxton, W. F.; Bouxsein, N. F.; Henderson, I. M.; Gomez, A.; Bachand, G. D. Dynamic Assembly of Polymer Nanotube Networks via Kinesin Powered Microtubule Filaments. Nanoscale 2015, 7, 10998−11004. (36) Roux, A. The Physics of Membrane Tubes: Soft Templates for Studying Cellular Membranes. Soft Matter 2013, 9, 6726−6736. (37) Bairi, P.; Minami, K.; Hill, J. P.; Nakanishi, W.; Shrestha, L. K.; Liu, C.; Harano, K.; Nakamura, E.; Ariga, K. Supramolecular Differentiation for Construction of Anisotropic Fullerene Nanostructures by Time-Programmed Control of Interfacial Growth. ACS Nano 2016, 10, 8796−8802.

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