Photoinduced Fusion of Lipid Bilayer Membranes - ACS Publications

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Photo-induced fusion of lipid bilayer membranes Yui Suzuki, Ken H. Nagai, Anatoly Zinchenko, and Tsutomu Hamada Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.7b00448 • Publication Date (Web): 13 Feb 2017 Downloaded from http://pubs.acs.org on February 14, 2017

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Photo-induced fusion of lipid bilayer membranes Yui Suzuki,1 Ken H. Nagai,1 Anatoly Zinchenko,2 Tsutomu Hamada*, 1 1

School of Materials Science, Japan Advanced Institute of Science and Technology, 1-1

Asahidai, Nomi, Ishikawa 923-1292, Japan 2

Graduate School of Environmental Studies, Nagoya University, 1 Furo-cho, Chikusa-ku,

Nagoya, Aichi 464-8601, Japan

ABSTRACT: We have developed a novel system for photocontrol of the fusion of lipid vesicles through the use of a photosensitive surfactant containing an azobenzene moiety (AzoTAB). Real-time microscopic observations clarified a change in both the surface area and internal volume of vesicles during fusion. We also determined the optimal cholesterol concentrations and temperature for inducing fusion. The mechanism of fusion can be attributed to a change in membrane tension, which is caused by the solubilization of lipids through the isomerization of AzoTAB. We used a micropipette technique to estimate membrane tension, and discuss the mechanism of fusion in terms of membrane elastic energy. The obtained results regarding this novel photo-induced fusion could lead to a better understanding of the mechanism of membrane fusion in living cells and may also see wider applications, such as in drug delivery and biomimetic material design.

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Introduction Living cells are enclosed by an elastic membrane that shows structural dynamics under physical or chemical stress. This flexibility of the membrane helps cellular and extracellular substances to pass through the membrane barrier.1 Recently, this intriguing property of a fluid membrane has attracted considerable attention because membrane transport may be useful for the application of synthetic substances such as medicines. Along these lines, many biomembranemimetic studies have been performed with the use of artificial lipid vesicles (spherical membrane shells of lipid bilayers).2 The most important issue in these studies is how to control the membrane dynamics. The design of molecular systems that produce biomimetic dynamical motion is an outstanding challenge. Since membrane morphological changes are governed by the elastic free energy of the membrane, it is important to understand the physico-chemical characteristics of the membrane if we wish to construct and manipulate controllable membrane systems.3 A better understanding of the principles that underlie membrane reactions should help to extend the possibility of biomimetic technology. In the area of membrane dynamics, membrane fusion is one of the most essential processes. Fusion plays a role in many biological transport functions such as fertilization, neurotransmission and intracellular trafficking.4 With respect to technological applications, such as drug delivery, fusion could be an effective method for transporting molecules encapsulated by lipid vesicles into target cells.4 However, the physico-chemical principles of membrane fusion are not well understood and need to be elucidated to promote progress in membrane engineering. To characterize membrane fusion, studies on model membrane systems, such as artificial lipid


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vesicles, have been conducted. Fusion has been observed with various molecules or sources of stimulation, such as fusogenic peptides,5-7 synthetic lipid,8,9 poly ethylene glycol,10,11 ions,12 and an electric field pulse.13 These experimental results show that specialized proteins are not necessary for membrane fusion. In addition, studies on an elastic model of membrane fusion have been reported. Since Kozlov and Markin first attempted to explain fusion in terms of membrane bending energy,14 the morphological changes during fusion has been revealed.15-20 Most recently, coarse-grained models of lipid bilayers have also been developed.21-26 According to those theoretical studies, membrane fusion is expected to proceed through a hemifusion intermediate followed by pore expansion (see Figure S1). In recent studies of membrane fusion, it has been suggested that membrane tension due to area strain is a significant contributor to fusion events.27 Membrane tension is a restoring force, which works when the area per lipid of the membrane differs from the optimal area. In biological systems, tension is induced by osmotic pressure or membrane interaction with the cytoskeleton.28,29 The hypothesis of lateral tension-induced fusion has been supported experimentally in protein-free model membrane systems. In planar lipid membranes, fusion pores opened under a membrane tension 10-fold greater than that in a living cell.30 Membrane tension generated by osmotic stress has been shown to drive vesicle-to-planar membrane fusion.31 In addition to membrane tension, the fusion process is also promoted by local curvature stress, which is driven by fusion proteins (such as SNAREs or viral fusion proteins) in a cell.27 The hemifusion intermediate is stabilized by negative spontaneous curvature that is induced by inverted corn-type lipids, such as cholesterol, phosphatidylethanolamine, and unsaturated lipids, or a temperature increase, where the thermal motion of lipid tails effectively promotes membrane splaying.4


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Here, we developed a new experimental system for the fusion of cell-sized giant unilamellar vesicles (GUVs) under light irradiation. Our system uses a photosensitive surfactant (azobenzene trimethylammonium bromide; AzoTAB) to induce membrane tension under light. AzoTAB is an amphiphilic molecule that has a cationic head and a hydrophobic tail containing an azobenzene component, which photoisomerizes between its trans and cis forms under visible light and UV irradiation, respectively (Scheme 1). Recently, Baigl and co-workers reported that AzoTAB does not deform GUVs under visible light, whereas it bursts individual vesicles under UV irradiation.32 They also clarified that there was a significant difference in affinity between the cis and trans isomers of AzoTAB for lipid vesicles using spectroscopy.32 Since cis isomers are less hydrophobic than trans isomers, trans-AzoTAB is incorporated into GUVs more efficiently than cis-AzoTAB. Generally, surfactants with a large hydrophilic part extract lipids from a bilayer vesicle to make small micelles (solubilization), which leads to a decrease in the surface area of the vesicle, i.e., area strain of the membrane.33 Thus, under UV irradiation, cis-isomerized AzoTAB tends to move to an outer solution while extracting some lipids from GUVs. Motivated by these results, we used AzoTAB to regulate membrane tension and demonstrated membrane fusion under the application of light. Since light is a superior external stimulus than other stimuli, in that its intensity and duration are both easy to control, real-time observation is both very clear and smooth.

Scheme 1. Photoisomerization reaction of AzoTAB


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Experimental section Materials. We used a surfactant that contains an azobenzene moiety (azobenzene trimethylammonium bromide; AzoTAB) to photocontrol the membrane morphology. The details of the synthesis of AzoTAB have been described previously.34 Lipid vesicles were made of 1,2diolyoyl-sn-glycerol-3-phosphocholine (DOPC) and various amounts of cholesterol. For fluorescent observations, we used N-(rhodamine red-X)-1,2-dihexadecanoyl-sn-glycero-3phosphoethanolamine triethylammonium salt (rhodamine-PE). DOPC and cholesterol were purchased from Avanti Polar Lipids, and rhodamine-PE was obtained from Invitrogen. Deionized water obtained from a Millipore Milli Q purification system was used to prepare reagents. Preparation of GUVs. To prepare lipid vesicles, we used an electroformation method.35 First, 40 µL of a 10 mM solution containing a mixture of DOPC and cholesterol in chloroform was


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spread on an indium tin oxide (ITO) electrode. After the lipid film was dried, 400 µL of sucrose solution (300 mM) was introduced between the two electrodes separated by a 1mm silicone spacer. Electroformation was performed using a sinusoidal AC field (1V, 10Hz) for 2h. Preparation of the sample chamber. The vesicle suspension was extracted and mixed with the same volume of a solution containing glucose (300 mM) and AzoTAB (1 mM). The final concentrations of lipids and AzoTAB were both 0.5 mM. This solution was dropped on a glass coverslip, covered with another smaller coverslip at a spacing of 1 mm, and left to stand for 2h in the dark to sediment vesicles in the bottom of the chamber by gravity. Microscopic observation. We observed lipid vesicles with a phase contrast microscope (IX 71, Olympus). For UV irradiation, we used a standard filter set (WU, Olympus; λex = 330-385 nm, dichroic mirror 400 nm, λem = 420 nm), with an extra-high-pressure mercury lamp. The power of the irradiated light on the microscopy stage was measured using an optical power measurement system (X-Cite XP750 & XR2100, EXFO). For the microscope stage, we used a temperature controller (TOKAI HIT) so that we could investigate the membrane dynamics under a wide range of temperatures (the microscopic setup is shown in Figure 1).


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Figure 1. Microscopic setup for the photoirradiation of GUVs.

Calculation of the surface area and volume of vesicles throughout fusion. The 2D outline of the vesicle shape was extracted from the image data. All of the Y coordinates of this outline were calculated for every 2 pixels (0.34 µm) of X coordinate. We approximated the 3D vesicle shape by the sum of the thin cylinders, and calculated the surface area and volume. Calculation of the surface area by micromanipulation. We used a micromanipulator (TransferMan 4, Eppendorf) with an ultra-fine glass capillary (Piezo drill tip, Eppendorf; OD = 8 µm, ID = 6 µm) to observe membrane shrinkage under UV. We aspirated a vesicle with a capillary, and measured the difference in the shape of the deformed vesicle between trans- and


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cis-isomers under a constant suction pressure. We then obtained the rate of change of the surface area as previously reported.36 Measurement was conducted with the use of low-intensity UV (1530 µW). Under high-intensity UV (100 µW), vesicles ruptured. The photo-responsive motion of vesicles, such as area expansion and fusion, proceeded within a few second. We performed the measurements just after the end of these events. Results and Discussion We observed two adjacent vesicles in the presence of AzoTAB, and examined the response of the morphology to photo-irradiation. We changed the fraction of cholesterol of vesicles and temperature of the microscopic stage. Figure 2A shows typical images of UV-induced fusion of GUVs of DOPC/cholesterol (1:1) at 70 °C (Movie S1). In addition to vesicle fusion, we observed the adhesion of vesicles in the absence of fusion (DOPC GUV at 22 °C, Figure 2B). Figure 2C shows the conditions under which fusion and adhesion were observed. We found that fusion was induced by cis-isomerization of AzoTAB when we used vesicles containing 40-60% cholesterol at temperatures of 40-70°C. Notably, we confirmed that trans-isomerization of AzoTAB does not induce fusion behaviors (Figure S4). In our observations, the intensity of UV irradiation was 6 µW. High power UV (>50 mW) does not tend to induce fusion (Figure S5).


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Figure 2. (A) Time sequence images of the fusion of two GUVs made of DOPC/cholesterol (4:6) in the presence of AzoTAB (0.5 mM) at 70°C. The numbers in the upper left of each image denote the elapsed time (in seconds). The scale bar represents 10 µm. (B) Time sequence images of the adhesion of two GUVs made of only DOPC in the presence of AzoTAB (0.5 mM) at 22°C. The scale bar represents 10 µm. (C) Effects of cholesterol mole fraction and temperature on fusion. The open circles denote when the adhesion of vesicles was observed, and filled circles denote when both adhesion and fusion (more than 5%) were observed (see also Figure S2).

The fusion event occurred within an experimental time-scale of approximately 1 second (Figure 2A). From the video images, we calculated the changes in surface area and the volume of vesicles throughout the fusion process (Figure 3A). The volume of the fused vesicle was essentially equal to the sum of the initial two vesicles. In contrast, the total surface area decreased during fusion. This probably results from membrane fragments generated from contacting area of two vesicles as shown in Figure 2A. Figure 3B shows the rates of change for the volume and surface area during fusion for several events. The surface area of fused vesicles decreased by approximately 20%. Conservation of the internal volume indicates that there was no loss of vesicle content during this photo-induced fusion. A decrease in area with a conservation of volume during fusion was also reported in a previous study with electrofusion.13


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Figure 3. (A) Time-dependent change in the surface area and volume during fusion.

(B)

Distribution of the rates of change for the surface area and volume of fusing vesicles. The number of observations is n=10.


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To examine the properties of fusion intermediate, we prepared two different adjacent vesicles; a fluorescent-labeled vesicle (DOPC/rhodamine-PE (98:2)) and an unlabeled vesicle (only DOPC), and checked dye movement during UV-induced behaviors. At room temperature (22°C), the fluorescent dye did not move into another vesicle, whereas the dye did migrate after adhesion at 70°C (Figure 4). This result implies that the adhering DOPC vesicles under a tension achieve hemifusion at high temperature. The UV-induced hemifusion doesn’t require cholesterol.


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Figure 4. Dye migration caused by UV-induced adhesion, and plots of the gray value of fluorescent images along the white dashed line of (A) and (B). The vesicles are composed of only DOPC and the temperature is 70°C. The scale bar represents 10 µm.

We then quantified the membrane tension of GUV in the presence of AzoTAB. As the UVinduced solubilization of AzoTAB proceeds, the actual surface area of the vesicle differs from the equilibrium area, which is the optimal surface area per lipid molecule on the vesicle surface, provided that the vesicle volume remains constant. Consequently, solubilization leads to area strain in the membrane (Figure 5A). The membrane tension σ is proportional to the area strain as σ = κa(A-A0)/A0 (2) where κa is an area stretching elastic modulus, A is the surface area and A0 is the equilibrium area of a vesicle.37 In our experiments, membrane tension is induced when trans-AzoTAB isomerizes to the cis form. We consider that A0 and A correspond to the surface area of vesicles with cis- and trans-AzoTAB, respectively. We measured the surface area of a vesicle before and after UV irradiation by a micromanipulation technique (Fig. S6, Movie S2). We aspirated a vesicle with a glass micropipette, and measured the difference in the shape of the deformed vesicle between the trans- and cis-isomers under a constant suction pressure. This experiment was carried out with DOPC vesicles under room temperature. Figure 5B shows the distribution of the rate of change for the area in photosensitive vesicles. The average ratio was 2.6±1.7%. For an area stretching modulus of DOPC membranes under room temperature of 265 mN/m,38 the membrane tension is estimated to be 7 mN/m. In addition, the area stretching modulus has been reported to increase with an increase in the cholesterol fraction (κa of DOPC/cholesterol (1:1)


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vesicles was around three-fold larger than κa of DOPC vesicles), and is essentially independent of temperature (around 5 % shift between 15 and 45 °C).39,40 Assuming that the photo-induced change in surface area of vesicles does not depend on cholesterol mixing fraction and temperature in our experimental condition, we estimate the membrane tension of DOPC/cholesterol (1:1) vesicles to be 21 mN/m. Our experimental results show that hemifusion (DOPC vesicles, 70 °C) and complete fusion (DOPC/cholesterol (1:1) vesicles, 70 °C) were achieved under tensions of 7 mN/m and 21 mN/m, respectively. A previous numerical study reported that the surface area of lipid vesicles needs to change by 7-25% to achieve fusion.25 This change corresponds to a membrane tension of about 16-58 mN/m. It was also reported that hemifusion was induced under smaller tensions than those to achieve fusion.24 Our experimental results are comparable to the value from the simulation. It is also noted that DOPC vesicles achieved hemifusion not at 22 °C but at 70 °C, although the lateral tension is essentially independent of temperature. This indicates that temperature-induced negative spontaneous curvature of membrane lipids facilitates the formation of hemifusion states.


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Figure 5. (A) Proposed mechanism of the UV-induced increase in tension. (B) Distribution of the rate of change for the membrane area upon UV irradiation. The total number of observations was n=30.


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Next, we discuss the process of pore expansion after hemifusion with a phenomenological model. The free energy of pore expansion as a function of the pore radius r can be described in terms of membrane tension σ and the line tension of the pore edge γ: F(r) = 2πrγ − πr2σ. The activation energy is given by F* = πγ2/σ. The pore line tension can be expressed, approximately, in terms of the membrane bending modulus, κb, and the distance between the mid-planes of the fusing membranes, H (Figure S7A), as γ = π (κb/H) where we assumed, for simplicity, that the line tension is derived from the bending energy at the pore rim.27 The membrane bending rigidity is κb = 20 kBT,40 and a recent study showed that bending rigidity of DOPC membranes is almost unchanged by both the cholesterol fraction and temperature.40,41 The addition of 50% cholesterol increases σ to reduce the activation energy by two-thirds (Figure S7B). Thus, DOPC/Chol vesicles tend to show fusion. When the activation energies for membranes consisting of DOPC/Chol (1:1) is a few kBT, we obtained H = 20~30 nm (we use κb = 20 kBT and σ = 21 mN/m). The confirmation of H value is beyond the scope of this paper, because a resolution is 166 nm/pix in our microscopic observation. Recently, FrançoisMartin et al. showed that the activation energy is ~20 kBT for spontaneous fusion observed within 5 min,42 which corresponds to H ≈ 10 nm. It was reported that the distance of two


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membranes with fusion proteins is H ≈ 13 nm.18 It should be also mentioned that, although we here discussed with a constant H, H can be a dynamic variable during the process of fusion.43,44 Kawamoto et al also reported that the activation energy of phenomenological models is overestimated by comparing with coarse-grained molecular dynamics.26 Lipid conformational change, which is not considered in the phenomenological models, can reduce the free energy barrier. High degree of lipid conformational change at high temperature may reduce the activation energy, which possibly explains that fusion requires high temperature in our experiments. The overestimation also implies that H would be smaller than that in the phenomenological model.

Conclusion We demonstrated a novel example of membrane engineering that consists of vesicle fusion induced by photo-irradiation. The solubilization reaction of a photo-sensitive surfactant was used to increase the membrane tension of vesicles. We obtained a phase diagram of vesicle fusion in terms of both the cholesterol contents and temperature. The addition of cholesterol and an increase in temperature tend to induce fusion through increasing lateral tension and negative spontaneous curvature. We discussed the mechanism of photo-induced vesicle fusion in terms of the elastic free energy of the membrane, including the membrane tension of stretching and the line tension of a fusion pore. To the best of our knowledge, this is the first experimental report to determine the membrane tension required to promote vesicle-vesicle fusion. These results could be useful for the development of strategies for light-triggered bioreactors and novel drugdelivery systems.


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ASSOCIATED CONTENT Supporting Information. Figures S1-S7, movies and movie legends. This material is available free of charge via the Internet at http://pubs.acs.org. AUTHOR INFORMATION Corresponding Author *[email protected] Notes The authors declare no competing financial interest. ACKNOWLEDGMENT We thank Prof. Damien Baigl (École Normale Supérieure) and Dr. Mafumi Hishida (Tsukuba University) for their useful discussions. This work was supported by MEXT KAKENHI Grant Nos. 15H00807 and 26103516, JSPS KAKENHI Grant No. 15K12538, AMED-CREST, AMED, and the Shibuya Foundation. REFERENCES (1) Alberts, B.; Johnson, A.; Lewis, J.; Raff, M.; Roberts, K.; Walter, P. Molecular Biology of the Cell, 5th edition; Garland Science: New York, 2008; pp 749-812.


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(2) Kooijmans, S. A. A.; Vader, P.; van Dommelen, S. M.; van Solinge, W. W.; Schiffelers, R. M. Exosome Mimetics: A Novel Class of Drug Delivery Systems. Int. J. Nanomed. 2012, 7, 1525−1541. (3) Hamada, T.; Sugimoto, R.; Vestergaard, M. C.; Nagasaki, T.; Takagi, M. Membrane Disk and Sphere: Controllable Mesoscopic Structures for the Capture and Release of a Targeted Object. J. Am. Chem. Soc. 2010, 132, 10528-10532. (4) Markvoort, A. J.; Marrink, S. J. Lipid Acrobatics in the Membrane Fusion Arena. In Membrane Fusion; Chernomordik, L. V.; Kozlov, M. M., Eds.; Elsevier: New York, 2011; pp 260-261, 273-276. (5) Takiguchi, K.; Nomura, F.; Inaba, T.; Takeda, S.; Saitoh, A.; Hotani, H. Liposomes Possess Drastic Capabilities for Topological Transformation. ChemPhysChem 2002, 3, 571-573. (6) Nomura, F.; Inaba, T.; Ishikawa, S.; Nagata, M.; Takahashi, S.; Hotani, H. Microscopic Observations Reveal that Fusogenic Peptides Induce Liposome Shrinkage Prior to Membrane Fusion. Proc. Natl. Acad. Sci. 2004, 101, 3420-3425. (7) Nikolaus, J.; Stöckl, M.; Langosch, D.; Volkmer, R.; Herrmann, A. Direct Visualization of Large and Protein-Free Hemifusion Diaphragms. Biophys. J. 2010, 98, 1192-1199. (8) Menger, F. M.; Balachander, N. Chemically-Induced Aggregation, Budding, and Fusion in Giant Vesicles: Direct Observation by Light Microscopy. J. Am. Chem. Soc. 1992, 114, 58625863.


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(9) Menger, F. M.; Lee, S. J. Induced Morphological Changes in Synthetic Giant Vesicles: Growth, Fusion, Undulation, Excretion, Wounding, and Healing. Langmuir 1995, 11, 36853689. (10) Lee, J.; Lentz, B. R. Secretory and Viral Fusion may Share Mechanistic Events with Fusion Between Curved Lipid Bilayers. Proc. Natl. Acad. Sci. 1998, 95, 9274-9279. (11) Haque, M. E.; McIntosh, T. J.; Lentz, B. R. Influence of Lipid Composition on Physical Properties and PEG-Mediated Fusion of Curved and Uncurved Model Membrane Vesicles: “Nature’s Own” Fusogenic Lipid Bilayer. Biochemistry 2001, 40, 4340-4348. (12) Haluska, C. K.; Riske, K. A.; Marchi-Artzner, V.; Lehn, J. M.; Lipowsky, R.; Dimova, R. Time Scales of Membrane Fusion Revealed by Direct Imaging of Vesicle Fusion with High Temporal Resolution. Proc. Natl. Acad. Sci. 2006, 103, 15841-15846. (13) Büschl, R.; Ringsdorf, H.; Zimmermann, U. Electric Field-Induced Fusion of Large and Polymerizable Liposomes from Natural Lipids. FEBS Lett. 1982, 150, 38-42. (14) Kozlov, M. M.; Markin, V. S. Possible Mechanism of Membrane Fusion. Biofizika, 1983, 28, 242-247. (15) Chernomordik, L. V.; Melikyan, G. B.; Chizmadzhev, Y. A. Biomembrane Fusion: A New Concept Derived from Model Studies using Two Interacting Planar Lipid Bilayers. Biochim. Biophys. Acta. 1987, 906, 309-352. (16) Siegel, D. P. Energetics of Intermediates in Membrane Fusion: Comparison of Stalk and Inverted Micellar Intermediate Mechanisms. Biophys. J. 1993, 65, 2124-2140.


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(17) Kuzmin, P. I.; Zimmerberg, J.; Chizmadzhev, Y. A.; Cohen, F. S. A Quantitative Model for Membrane Fusion Based on Low-Energy Intermediates. Proc. Natl. Acad. Sci. 2001, 98, 7235-7240. (18) Kozlovsky, Y.; Kozlov, M. M. Stalk Model of Membrane Fusion: Solution of Energy Crisis. Biophys. J. 2002, 82, 882-895. (19) Katsov, K.; Müller, M.; Schick, M. Field Theoretic Study of Bilayer Membrane Fusion. I. Hemifusion Mechanism. Biophys. J. 2004, 87, 3277-3290. (20) Katsov, K.; Müller, M.; Schick, M. Field Theoretic Study of Bilayer Membrane Fusion: II. Mechanism of a Stalk-Hole Complex. Biophys. J. 2006, 90, 915-926. (21) Marrink, S. J.; Mark, A. E. Molecular dynamics simulation of the formation, structure, and dynamics of small phospholipid vesicles. J. Am. Chem. Soc. 2003, 125, 15233-15242. (22) Stevens, M.; Hoh, J.; Woolf, T. Insights into the Molecular Mechanism of Membrane Fusion from Simulation: Evidence for the Association of Splayed Tails. Phys. Rev. Lett. 2003, 91, 188102. (23) Noguchi, H.; Takasu, M. Fusion Pathways of Vesicles: A Brownian Dynamics Simulation. J. Chem. Phys. 2001, 115, 9547-9551. (24) Shillcock, J.; Lipowsky, R. Tension-induced Fusion of Bilayer Membranes and Vesicles. Nature Materials 2005, 4, 225-228. (25) Gao, L.; Lipowsky, R.; Shillcock, J. Tension-Induced Vesicle Fusion: Pathways and Pore Dynamics. Soft Matter 2008, 4, 1208-1214.


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(41) Gracià, R.S.; Bezlyepkina, N.; Knorr, R. L.; Lipowsky, R.; Dimova, R. Effect of Cholesterol on the Rigidity of Saturated and Unsaturated Membranes: Fluctuation and Electrodeformation Analysis of Giant Vesicles. Soft Matter 2010, 6, 1472-1482. (42) François-Martin C.; Rothman J. E.; Pincet F. Low energy cost for optimal speed and control of membrane fusion. Proc. Natl. Acad. Sci. 2017, 114, 1238-1241. (43) Chizmadzhev, Y. A.; Cohen, F. S.; Shcherbakov, A.; Zimmerberg, J. Membrane mechanics can account for fusion pore dilation in stages. Biophys. J. 1995, 69, 2489-2500. (44) Chizmadzhev, Y. A.; Kuzmin, P. I.; Kumenko, D. A.; Zimmerberg, J.; Cohen, F. S. Dynamics of Fusion Pores Connecting Membranes of Different Tensions. Biophys. J. 2000, 78, 2241-2256.


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Photoinduced Fusion of Lipid Bilayer Membranes - ACS Publications

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