Self-assembly of lipids forms various architectures such as vesicles, tubes and micelles with spherical, rod and disk shapes [1–4]. Among them, a vesicle is a ubiquitous structure seen in biological systems such as membranes surrounding cells and organelles [5–7]. Not only the unique two-dimensional bilayer structure but also the dynamic properties of the membranes arouse much interest of scientists. In extant cells, vesicle budding plays crucial roles in vital activities such as endocytosis and vesicle trafficking. Furthermore, membrane-enveloped organelles such as Goldi apparatus and endoplasmic reticulum are known to perform vesicle budding to deliver membrane proteins or soluble substances to the other organelles, plasma membrane, and extracellular space . Inspired by the dynamic behaviors of the biomembranes, transformation of synthetic membranes has been actively explored, where fusion, division and budding, triggered by heating, addition of membrane-interacting molecules, osmotic pressure or photochemical reactions have been demonstrated [9–24]. Recently, we reported light-triggered μm-size vesicle formation from particles  composed of phospholipids and multi-block amphiphiles bearing 1,4-bis(4-phenylethynyl)benzene units [26, 27]. Here for deeper understanding of this phenomenon, we investigated the effect of amphiphilicity of the chromophore as well as the effect of solute in the hydration media on the vesicle formation. In addition, we demonstrate light-triggered vesicle formation in a confined microenvironment and encapsulation of a substance in the generated vesicles as possible applications.
1,2-Dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (Rhod-PE) and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-2-1,3-benzoxadiazol-4-yl) (NBD-PE) were purchased from Avanti Polar Lipids (Alabaster, AL, USA). NaCl, glucose and sucrose were purchased from Nacalai Tesque (Kyoto, Japan). Anhydrous CHCl3, MeOH and 1,4-bis(phenylethynyl)benzene (BPEB, Fig. 1) were purchased from Wako Pure Chemicals (Osaka, Japan). These commercial reagents were used without purification. Deionized water (filtered through a 0.22 μm membrane filter, >18.2 MΩ cm) was purified in a Milli-Q system of Millipore Corp. (Billerica, MA, USA). Trimethyl (14Z,64Z,3E)-11H,61H-9,12,15,18,26,29,32,35,38,40,43,46,49,52,60,63,66,69-octadecaoxa-3,4-diaza-1,6(4,1)-ditriazola-2,5(1,3),19,22,25,53,56,59(1,4)-octabenzenacyclomonoheptacontaphan-3-ene-20,23,54,57-tetrayne-26,54,395-tricarboxylate (cycAzo-T4B2, Fig. 1) and 4,4′-(1,4-phenylenebis(ethyne-2,1-diyl))diphenol (BPEBOH, Fig. 1) was synthesized following the reported procedure [25, 28].
Bright-field, fluorescent and phase-contrast microscopy was performed with an Olympus (Tokyo, Japan) IX-71 microscope, where a U-MWU2 mirror unit (excitation filter: 330–385 nm, emission filter: 420 nm, dichroic mirror: 400 nm) was used for the fluorescence observation, and the excitation light through the filter (330–385 nm) was used for the photo-irradiation. Confocal laser scanning microscopy was performed with an Olympus IX-81 microscope. Sonication and vortex for the preparation of lipid particles were performed with a vortex genie 2 (Scientific Industries, Bohemia, NY, USA) and Bioruptor UCD-250 cell homogenizer (250 W, COSMO BIO, Tokyo, Japan), respectively.
Preparation and photo-irradiation of particles for study on amphiphilicity effect
DOPC (2.0 mM), BPEB or BPEBOH (800 μM), and glucose (16 mM) were dissolved in a mixture of anhydrous MeOH and CHCl3 (1.0/1.0 v/v) followed by evaporation of the solvents under Ar flow at 20 °C to leave a uniform thin film at the bottom of a glass test tube. The obtained film was placed under vacuum at 20 °C for longer than 1.5 h. The resulting film was then hydrated with an aqueous solution of glucose ([glucose] = 200 mM, [DOPC] = 200 μM, [BPEB or BPEBOH] = 80 μM) or NaCl ([NaCl] = 100 mM, [DOPC] = 200 μM, [BPEB or BPEBOH] = 80 μM), which was immediately subjected to sonication for 15 min at 0–5 °C. We also found that 10-s vortex mixing of the hydrated film can afford the photoresponsive particles. Photo-irradiation was performed with the light source of the microscope during the microscopic observations.
Preparation and photo-irradiation of particles for applications
Typical procedure: DOPC (2.0 mM), Rhod-PE (60 μM) and cycAzo-T4B2 (400 μM) were dissolved in a mixture of anhydrous MeOH and CHCl3 (1.0/1.0, v/v) followed by evaporation of the solvents under Ar flow at 20 °C to leave a uniform thin film at the bottom of a glass test tube. The obtained film was placed under vacuum at 20 °C for longer than 1.5 h. The resulting film was then hydrated with an aqueous solution of sucrose and glucose mixture ([sucrose] = [glucose] = 100 mM; [DOPC] = 200 μM, [Rhod-PE] = 6.0 μM, [cycAzo-T4B2] = 40 μM), which was immediately subjected to sonication for 15 min at 0–5 °C. We also found that 10-s vortex mixing of the hydrated film can afford the photoresponsive particles. Photo-irradiation was performed with the light source of the microscope during the microscopic observations.
Effect of BPEB unit on light-triggered vesicle formation
In our previous paper , we reported that the BPEB units in cycAzo-T4B2 are likely the essential components enabling the light-triggered vesicle formation, since hydrogenation of the ethynylene bonds in the BPEB units resulted in no vesicle formation under irradiation. In order to get a more direct evidence for the role of the BPEB unit for the vesicle formation, we prepared particles including BPEB and BPEBOH.
Upon light irradiation to the particles prepared from a mixture of DOPC and BPEBOH at 25 °C (λ = 330–385 nm), μm-size vesicle budding from the particles were promptly observed (Fig. 2a, b). However, interestingly, the particles prepared from a mixture of DOPC and BPEB did not show the vesicle formation under the light illumination (Fig. 2c, d). Thus, while it was confirmed that the BPEB unit enables photo-generation of the vesicles from the particle, it is also suggested that there is an another factor responsible for this phenomenon.
For fluorometric visualization of the phospholipid component, particles composed of DOPC, NBD-PE, and BPEBOH or BPEB were prepared. The resulting particles including BPEBOH also showed the light-triggered vesicle budding. Under confocal laser scanning (CLS) microscopic observation, a particle including BPEBOH visualized the fluorescence of BPEBOH over the particle upon excitation with 405-nm light (Fig. 3a). Green fluorescence from the NBD group upon excitation with 473-nm light mostly overlapped with the BPEBOH fluorescence (Fig. 3b, c). These observations indicate that BPEBOH has high miscibility with the phospholipids and does not form domains in the particles. In sharp contrast, a particle prepared from DOPC, NBD-PE, and BPEB showed the green fluorescence of the NBD group only at limited areas in the particle (Fig. 3f), although most parts of the particle displayed the fluorescence of BPEB (Fig. 3e). These contrasting results between BPEBOH and BPEB suggest that the amphiphilicity of the chromophore, which allows blending with DOPC, is also an important factor for the light-triggered vesicle formation.
Effect of ionic solute on light-triggered vesicle formation
It is known that the zwitterionic structure of a phosphatidylcholine, such as DOPC, strengthens the multilayer membranes due to the formation of strong electrostatic interactions between the bilayer membranes. As reported previously, the particles are consisting of multilayer membranes . This suggests a possible effect of ionic solutes to facilitate detachment of the membranes from the particles to encourage the formation of vesicles . When the particles, prepared from a mixture of DOPC and BPEBOH in the aqueous solution of glucose (200 mM), was irradiated, they left a portion of dormant “husks” after the vesicle formation. In contrast, it was found that the particles prepared from a mixture of DOPC and BPEBOH in 100 mM NaCl aq. (Fig. 4a), in place of glucose aq., generated vesicles by irradiation, where most portions of the particle change into membranes. Once a membrane emerged from a particle (Fig. 4b), it grew continuously (Fig. 4c), and finally, membrane structures over 10-μm diameter were formed with almost no husk remaining (Fig. 4d). Obviously, NaCl encourages the photo-induced vesicle formation.
Vesicle formation in microenvironment
The notable advantage of photo-generation of vesicles from the particles is that this methodology is potentially applicable for preparation of vesicles in a confined space like a cell, just like the vesicle budding. As a demonstration for proving this concept, we conducted a light-driven vesicle budding from the particle within a microchamber. For this demonstration, the particles were prepared from a mixture of DOPC, Rhod-PE and cycAzo-T4B2 ([DOPC] = 200 μM, [Rhod-PE] = 6.0 μM, [cycAzo-T4B2] = 40 μM), where the composition was optimized for a few-minutes real-time observation of the budding and migration of the vesicles by fluorescence microscopy. The particles dispersed in an aqueous media were successfully trapped in a microchamber (90 μm width, 30 μm depth) by putting the aqueous suspension of the particles onto a microchamber array chip composed of an epoxy resin, which was then placed under reduced pressure for 10 min at 20 °C. Bright-field microscopy displayed a particle in a microchamber (Fig. 5a, c). The particle showed bright fluorescence upon irradiation of 405-nm and 532-nm laser beams, indicating the existence of cycAzo-T4B2 and Rhod-PE, respectively. The laser beam (405 nm, 5–20 mW) focused into a 10-μm spot was irradiated to the particle at 20 °C for the vesicle formation, meanwhile whole area of the chamber was irradiated with 532-nm laser light to excite Rhod-PE, allowing for observation by a fluorescence microscope (Fig. 5e–h). Actually, a dozen of fluorescent spheres, rapidly flitting around the chamber, were observed after photo-irradiation for 15 s, where further irradiation induced the continuous formation of fluorescent spherical objects in the microchamber. Diameter of the observed fluorescent spheres ranged from 1.5 to 3.6 μm. Although the photo-generated vesicles initially flitted in the microchamber, they stopped the flitting immediately, probably due to adsorption onto the bottom surface of the microchamber. After photo-irradiation for 75 s, the particle image observable by bright-field microscopy diminished (Fig. 5b, d), suggesting that the lipids contained in the particle turned into the fluorescent spheres, namely, the photo-generated μm-size vesicles.
Encapsulation of substance within vesicles
Encapsulation of substances is one of the important applications of vesicles for delivery [30–35]. Here we tried to encapsulate sucrose within the μm-size vesicles. A particle suspension (200 μl) was prepared from a mixture of DOPC (2.0 mM) and cycAzo-T4B2 (400 μM) in 200 mM sucrose aq. by sonication (250 W, 15 min, 0–5 °C). After UV light (330–385 nm, 1 kW, Asahi Spectra LAX-1000, Tokyo, Japan) was irradiated to the particles suspension for 90 min at 20 °C, 100 μl of the suspension was taken out and mixed with 100 μl of 200 mM glucose aq. Phase-contrast microscopy visualized μm-size dark spheres in the resulting suspension (Fig. 6). It is known that, under phase-contrast microscopy, the area containing sucrose aq. appears darker than the one containing glucose aq. because of the larger refractive index of sucrose aq. than glucose aq. . Thus, this result clearly suggests that sucrose was encapsulated within the photo-generated vesicles.
In the present article, it was indicated that not only the BPEB unit but also the amphiphilicity of the chromophore is necessary for the light-triggered vesicle budding from the particle. It is expected that the capability of the small amphiphiles such as BPEBOH to form μm-size vesicles by light could expand the possible applications of vesicles. Effectiveness of ionic solutes to enhance the budding was also demonstrated. In addition to these fundamental aspects, the light-triggered vesicle formation from particles was applied to μm-size vesicle preparation in a confined microchamber and substrate encapsulation, which are difficult to be performed by conventional μm-size vesicle preparation techniques . Since light has advantages as a local and contactless stimulus, we believe that the present study lends to the regulation of vesicles within a confined space such as a cell.
This work was partially supported by the ministry of education, culture, sports, Science and Technology in Japan (MEXT), Grants-in-Aid for Young Scientists S (21675003), Scientific Research on Innovative Areas “Spying minority in biological phenomena” (No.3306), (23115003), and the Management Expenses Grants for National Universities Corporations to K.K.
T. Shimizu, M. Masuda, H. Minamikawa. Chem. Rev. 105, 1401 (2005).
E. A. Chandross, R. D. Miller. Chem. Rev. 99, 1641 (1999).
T. Kunitake. Angew. Chem., Int. Ed. Engl. 31, 709 (1992).
G. M. Whitesides, J. P. Mathias. Science 254, 1312 (1991).
G. M. Whitesides, J. P. Mathias.)| false Science 254, 1312 (1991). 10.1126/science.1962191
P. L. Luisi. “Why giant vesicles?” in Giant Vesicles, Perspectives in Supramolecular Chemistry, Vol. 6, P. L. Luisi, P. Walde (Eds.), Wiley-Interscience, Weinheim (2000).
P. L. Luisi. “Why giant vesicles?” in)| false Giant Vesicles, Perspectives in Supramolecular Chemistry, Vol. 6, P. L. Luisi, P. Walde (Eds.), Wiley-Interscience, Weinheim (2000). 10.1002/9780470511534
E. Sackmann. “Biological membranes architecture and function” in Structure and Dynamics of Membranes, 1st ed., R. Lipowsky, E. Sackmann (Eds.), Elsevier, Amsterdam (1995).
E. Sackmann. “Biological membranes architecture and function” in)| false Structure and Dynamics of Membranes, 1st ed., R. Lipowsky, E. Sackmann (Eds.), Elsevier, Amsterdam (1995). 10.1016/S1383-8121(06)80018-7
D. D. Lasic. “General introduction to liposomes” in Liposomes: From Physics to Applications, D. D. Lasic (Ed.), Elsevier, Amsterdam (1993).
Intracellular Vesicular Traffic. Molecular Biology of the Cell, 5th ed., B. Alberts, A. Johnson, J. Lewis, M. Raff, K. Roberts, P. Walter (Eds.), Garland Science, New York (2007).
A. Diguet, M. Yanagisawa, Y.-J. Liu, E. Brun, S. Abadie, S. Rudiuk, D. Baigl. J. Am. Chem. Soc. 134, 4898 (2012).
T. Hamada, R. Sugimoto, M. C. Vestergaard, T. Nagasaki, M. Takagi. J. Am. Chem. Soc. 132, 10528 (2010).
Y. Yu, J. A. Vroman, S. C. Bae, S. Granick. J. Am. Chem. Soc. 132, 195 (2010).
Z. Wang, K. Yasuhara, H. Ito, M. Mukai, J. Kikuchi. Chem. Lett. 39, 54 (2010).
G. P. Robbins, M. Jimbo, J. Swift, M. J. Therien, D. A. Hammer, I. J. Dmochowski. J. Am. Chem. Soc. 131, 3872 (2009).
K. Ishii, T. Hamada, M. Hatakeyama, T. Nagasaki, M. Takagi. ChemBioChem 10, 251 (2009).
K. Ishii, T. Hamada, M. Hatakeyama, T. Nagasaki, M. Takagi.)| false ChemBioChem 10, 251 (2009). 19132694
M. S. Long, A.-S. Cans, C. D. Keating. J. Am. Chem. Soc. 130, 756 (2008).
G. T. Charras. J. Microsc. 231, 466 (2008).
G. T. Charras.)| false J. Microsc. 231, 466 (2008). 10.1111/j.1365-2818.2008.02059.x
M. Yanagisawa, M. Imai, T. Taniguchi. Phys. Rev. Lett. 100, 148102 (2008).
E. Brückner, P. Sonntag, H. Rehage. Langmuir 17, 2308 (2001).
E. Brückner, P. Sonntag, H. Rehage.)| false Langmuir 17, 2308 (2001). 10.1021/la000256i
G. Cevc, H. Richardsen. Adv. Drug Deliver. Rev. 38, 207 (1999).
P. G. Petrov, J. B. Lee, H.-G. Döbereiner. Europhys. Lett. 48, 435 (1999).
J. Käs, E. Sackmann. Biophys. J. 60, 825 (1991).
J. Käs, E. Sackmann.)| false Biophys. J. 60, 825 (1991). 1742455
T. Kunitake, N. Nakashima, M. Shimomura, Y. Okahata, K. Kano, T. Ogawa. J. Am. Chem. Soc. 102, 6642 (1980).
T. Shima, T. Muraoka, T. Hamada, M. Morita, M. Takagi, H. Fukuoka, Y. Inoue, T. Sagawa, A. Ishijima, Y. Omata, T. Yamashita, K. Kinbara. Langmuir30, 7289 (2014).
T. Muraoka, T. Shima, T. Hamada, M. Morita, M. Takagi, K. V. Tabata, H. Noji, K. Kinbara. J. Am. Chem. Soc. 134, 19788 (2012).
T. Muraoka, T. Shima, T. Hamada, M. Morita, M. Takagi, K. Kinbara. Chem. Commun. 47, 194 (2011).
D. P. Flaherty, T. Kiyota, Y. Dong, T. Ikezu, J. L. Vennerstrom. J. Med. Chem. 53, 7992 (2010).
P. L. Luisi, M. Allegretti, T. P. de Souza, F. Steiniger, A. Fahr, P. Stano. ChemBioChem 11, 1989 (2010).
P. L. Luisi, M. Allegretti, T. P. de Souza, F. Steiniger, A. Fahr, P. Stano.)| false ChemBioChem 11, 1989 (2010). 10.1002/cbic.201000381
N. Ben-Haim, P. Broz, S. Marsch, W. Meier, P. Hunziker. Nano Lett. 8, 1368 (2008).
V. Noireaux, A. Libchaber. Proc. Natl. Acad. Sci. USA 101, 17669 (2004).
V. Noireaux, A. Libchaber.)| false Proc. Natl. Acad. Sci. USA 101, 17669 (2004). 10.1073/pnas.0408236101
P.-A. Monnard. J. Membrane Biol. 191, 87 (2003).
P.-A. Monnard.)| false J. Membrane Biol. 191, 87 (2003). 10.1007/s00232-002-1046-0
K. Tsumoto, S. M. Nomura, Y. Nakatani, K. Yoshikawa. Langmuir 17, 7225 (2001).
K. Tsumoto, S. M. Nomura, Y. Nakatani, K. Yoshikawa.)| false Langmuir 17, 7225 (2001). 10.1021/la010887s
M. Mally, J. Majhenc, S. Svetina, B. Žekš. Biophys. J. 83, 944 (2002).
M. Mally, J. Majhenc, S. Svetina, B. Žekš.)| false Biophys. J. 83, 944 (2002). 12124276