Membrane Proteins on Artificial Lipid Membranes

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This entry is adapted from the peer-reviewed paper 10.3390/ijms24087231

Membrane proteins play an important role in key cellular functions, such as signal transduction, apoptosis, and metabolism. Therefore, structural and functional studies of these proteins are essential in fields such as fundamental biology, medical science, pharmacology, biotechnology, and bioengineering.

membrane proteins ion channels nanopores giant lipid vesicles liposome proteoliposome planar bilayer lipid membrane artificial cell model

1. Introduction

Membrane proteins are a vital component of the cell membrane, which is composed of a phospholipid bilayer. They play a critical role in various cellular functions, such as molecular transportation, conversion, production, and transduction, including signal transduction, apoptosis, and metabolism ^[1]^[2]^[3]^[4]^[5]^[6]^[7]^[8]^[9]^[10]^[11]. Different types of membrane proteins on the cell membrane interact to activate these functions. G protein-coupled receptors (GPCR) and ion channels, which are types of membrane proteins, are important targets for drugs used to treat diseases such as cancer, neurodegenerative disease, metabolic disease and cardiovascular disease ^[12]^[13]^[14]^[15]^[16]^[17]. However, investigating the precise elemental reaction and structure of a membrane protein in complex cell reactions activated by several types of membrane proteins can be challenging. To overcome this issue, methodologies for investigating the function of membrane proteins purified from biological cells have been developed. In the early 1970s, proteoliposomes were first developed by reconstituting the Ca²⁺-dependent ATPase of the sarcoplasmic reticulum into nano-sized liposomes ^[18]^[19]. In the late 1980s, the proteoliposomes with K⁺-channels of the sarcoplasmic reticulum were generated using the cell-sized liposomes with diameters of approximately 10 μm, due to the measurement of single ion channel current ^[20]. Recently, cell-sized proteoliposomes with two or three types of membrane proteins were formed to connect each membrane protein reaction ^[21]^[22]^[23]. Purified membrane proteins have been crystallized to reveal their structures, and they have been reconstituted into artificial cell membranes, such as liposomes, planar lipid bilayers, and bicelles, to observe their functions ^[24]^[25]^[26]. These artificial cell membranes consist of phospholipid bilayers, and the structure and function of the membrane proteins reconstituted into them have been identified using various techniques, such as NMR, fluorescence activated cell sorter, cryo-electron microscopy (cryo-EM), fluorescence spectrophotometry, patch clamp method (for nanopores or ion channels), and optical microscopy, including fluorescence microscopy, and confocal microscopy ^[25]^[27]^[28]^[29]^[30]^[31]^[32]^[33]^[34]. These investigations have not only revealed the functions of membrane proteins but also led to the development of biological sensors using membrane proteins because of their highly specific interactions with ligands and membrane proteins ^[35]^[36]^[37]. Thus, studies of membrane proteins can have a wide range of applications (Figure 1).

Figure 1. Schematic illustration of functional studies and applications of membrane proteins on artificial lipid membranes.

2. Formation of Cell-Sized Lipid Vesicles

Membrane proteins are typically reconstituted into artificial cell membranes such as lipid vesicles, liposomes, and bicelles to maintain their structure and function. In this section, the researchers introduce methods for forming lipid vesicles and liposomes, from conventional techniques to recent microfluidic approaches. Liposomes and lipid vesicles, discovered by Bangham in the 1960s, consist of a phospholipid bilayer ^[38]. Conventional methods for liposome formation include gentle hydration and electroformation ^[39]^[40]^[41]^[42]^[43]. With the gentle hydration method, phospholipids dissolved in chloroform are added to a glass microtube, and the films are formed by evaporating the chloroform under flowing argon or nitrogen gas. The lipid films are then hydrated with an aqueous solution (pure water or buffer solution), forming giant liposomes with diameters of 1–100 μm. Nano-sized liposomes are obtained by vortexing or sonication of the lipid films ^[1]. With the electroformation method, a lipid film is formed on the surface of an indium tin oxide (ITO)-coated glass. An alternating current (AC) electric field is applied to the lipid films containing a hydrated solution (pure water or a buffer solution with low salt concentration) to enhance lipid hydration. Although the giant liposomes formed by the electroformation method have more unilamellar membranes than those formed by the gentle hydration method, the former has lower liposome production than the latter. Methods have been developed to improve the drawbacks of both the gentle hydration and electroformation methods. These conventional liposome formation methods have been used for studying membrane dynamics, functional properties of membrane proteins, and encapsulation of biological reactions.

The gentle hydration and electroformation methods are limited in their ability to produce liposomes with uniform sizes, in their high encapsulation efficiencies, and in their asymmetric lipid distributions. Microfluidic technologies have been integrated into liposome formation to overcome these limitations and achieve a uniform size and high encapsulation efficiency. The droplet emulsion transfer method, developed by Pautot et al. ^[44]^[45], is the basis for many of the liposome formation techniques using microfluidic technologies. The liposome formation procedure with the droplet emulsion transfer method is as follows: First, water-in-oil (w/o) emulsions are generated by vortexing or sonication of buffer solution and phospholipid solution dissolved in an organic solvent such as mineral oil or n-decane. The w/o emulsions are then added to the oil phase in a microtube. When the w/o emulsions are transferred to the lipid monolayer between the oil phase and the aqueous phase, liposomes are generated. Asymmetric lipid membranes between the outer leaflet and the inner leaflet can be generated by changing the lipid compositions of the w/o emulsions and the lipid monolayer. By applying the droplet emulsion transfer method, many high-throughput methods for forming liposomes using microfluidic devices have been developed ^[46]^[47]^[48]^[49]^[50]^[51]^[52]^[53]. These methods integrate the system of w/o emulsion generation and emulsion transfer from the oil phase to the aqueous phase. Like other methods of liposome formation using microfluidic devices, a new approach has been developed that creates single or multicompartment liposomes by utilizing the de-wetting phenomena of a water-in-oil-in-water double emulsion using a coaxial microcapillary fluidic device ^[54]^[55]^[56]^[57]^[58]^[59]^[60]^[61]^[62].

Another method for forming asymmetric lipid vesicles without an organic solvent (n-decane) between the lipid bilayer is by applying a pulsed jet flow against a planar lipid bilayer in a double-well device (called the pulsed-jet flow method) ^[63]^[64]. The lipid bilayer of liposomes formed by the microfluidic technology method is normally contaminated with an organic solvent, which can cause instability of the liposomes. In contrast, liposomes formed by the pulsed-jet flow method have long-term stability (up to one week). The flip-flop phenomenon and membrane protein reconstitution have been investigated on the asymmetric lipid vesicles formed by this method ^[64]^[65]. Modification of the double-well device allows for the formation of nano-sized asymmetric lipid vesicles ^[66], automatic formation of liposomes ^[67]^[68], and the formation of vesicles-in-a-vesicle structures that mimic the asymmetric lipid composition of plasma membranes and intracellular vesicle membranes ^[69]. Multiple enzyme reactions can be created within the liposomes formed by microfluidic devices ^[70]. A complex of cell-penetrating peptides and water-soluble proteins can interact with asymmetric lipid vesicles containing negatively charged lipids on the inner leaflet, and the water-soluble proteins can be transferred into the asymmetric lipid vesicles ^[71].

3. Methodology of Reconstitution of Membrane Proteins into Cell-Sized Artificial Membrane

Giant unilamellar vesicles (GUVs) are widely used as models of artificial cells ^[72]^[73]. To understand complex biological phenomena in cells, membrane protein function has been studied by reconstituting membrane proteins into GUVs. Various methods have been developed for this, including rehydration, membrane fusion, detergent removal, and direct insertion (Table 1) ^[74]^[75]^[76]^[77]^[78]^[79]^[80]^[81]^[82]^[83]^[84]^[85]^[86]^[87]^[88]^[89]^[90].

Table 1. Selected examples of membrane proteins reconstitution in GUVs.

Protein	Organism	Type	TM Region	Complex	TM Number	Method	Membrane Composition	Ref.
BmrC/BmrD	Bacillus subtilis	ABC transporter	α−helix	BmrC/BmrD	12	detergent mediate reconstitution	DPhPC + DOPC/DOPE or DOPC/Sph/chol	^[74]
BR	Halophilic archaea	proton pump	α−helix	−	7	fusion	EPC:EPA (9:1 [mol])	^[74]
CXCR4	Homo sapiens	GPCR signaling protein	α−helix	homodimer	14 (7 × 2)	fusion	DOPC	^[75]
EmrE	Escherichia coli	multidrug transporter	α−helix	homodimer	8 (4 × 2)	direct reconstitution	POPC	^[83]
ETB	Homo sapiens	GPCR signaling protein	α−helix	−	7	fusion	DOPC	^[75]
F₁F₀-ATP synthase	Escherichia coli	ATP production	α−helix	F₁–F₀	28	rehydration	E. coli total lipid extract	^[84]
FhuA	Escherichia coli	ferrichrome-iron receptor	β−strand	−	22	fusion	EPC:Tx-DHPE (99.5:0.5)	^[74]
GL1	Homo sapiens	γ-aminobutyric acid receptor	α−helix	−	anchored	rehydration	DOPE:DOPC:DOPE-Atto647 (30:69.5:0.5)	^[85]
GLUT1	Homo sapiens	glucose transporter	α−helix	−	12	fusion	DOPC:DMPE-RhB:Biotylated PE (99.7:0.2:0.1)	^[86]
IFITM3	Homo sapiens	enveloped virus inhibitor	α−helix	−	1	detergent mediate reconstitution	POPC:cholesterol:Liss-Rho-PE (99:0.5:0.5 [mol])	^[87]
KcsA	Streptomyces lividans	potassium channels	α−helix	homotetramer	8 (2 × 4)	direct reconstitution	DOPG or DOPE	^[88]
KvAP	Aeropyrum pernix	voltage-gated potassium channels	α−helix	heteromer or homotetramer	8 (2 × 4)	rehydration	DPhPC or EPC:EPA (9:1)	^[89]
OmpF	Escherichia coli	porin	β−strand	−	16	direct reconstitution	PDMS26-b-PMOXA9	^[90]
OmpG	Escherichia coli	porin	β−strand	−	14	direct reconstitution	Outer membrane: DOPC Inner membrane: oleosin	^[76]
OmpLA	Escherichia coli	phospholipase	β−strand	homodimer	24 (12 × 2)	direct reconstitution	DOPC:DOPG (1:3)	^[77]
PR	SAR86 group of marine γ-proteobacteria	proton transport	α−helix	−	7	direct reconstitution	POPC	^[78]
RC	Rhodobacter sphaeroides	electron transport	α−helix	−	10	detergent mediate reconstitution	POPC:POPG (9:1 [mol])	^[79]
SLO	Streptococcus pyogenes	toxin	α−helix	homo 36~40 mer	36~40	rehydration	POPC, DOPC, SOPC, POPG	^[80]
TmrAB	thermus thermophilus	ABC transporter	α−helix	−	6 × 2	rehydration	POPC:POPG:POPE: biotinylated-DOPE (40:30:29:1 [mol])	^[81]
TolC	Escherichia coli	expulsion of diverse molecules from the cell	β−strand	homotrimer	12 (4 × 3)	rehydration	DOPC:DOPS:Atto647-DOPE (91.2:8:0.8)	^[85]
t-SNARE	Homo sapiens	endocytosis/exocytosis	α−helix	Syntaxin -SNAP25	1	rehydration	DOPC:DOPS:Atto647-DOPE (91.2:8:0.8)	^[85]
VDAC1	Homo sapiens	voltage dependent anion channel	β−strand	−	19	fusion	soybean polar extract: cholesterol (9:1)	^[82]

4. Single-Channel Recording Using Planar Bilayer Lipid Membrane Systems

The function of membrane proteins on cell membranes, such as ion channels and transporters, has been studied using the patch-clamp method. Neher and Sakmann ^[91] developed the patch-clamp method to measure the function of ion channels and transporters at the single molecule level and in real time. While various patch-clamp methods have been developed for measuring ion currents of ion channels in biological target membranes, artificial planar bilayer lipid membrane (BLM) systems have recently been developed for an engineering-oriented approach ^[92]. In BLM systems, the target ion channels purified from living cells are reconstituted in the BLM, and the ionic current of the ion channels is measured using a patch clamp amplifier. BLMs allow us to perform functional studies of ion channels in a well-defined environment, such as buffer composition, lipid composition, inhibitor or activator concentration, and membrane potential.

The BLMs (Figure 2a–c) can also be formed by the painting method ^[93], the Langmuir–Blodgett method ^[94], or the droplet contact method ^[63]. In the painting method, a lipid solution (phospholipids dissolved in an organic solvent) is applied across a small aperture that separates two aqueous compartments, forming lipid monolayers. Then, a BLM is formed by the contact of each monolayer after the removal of the organic solvent ^[93]. This method of BLM formation has improved the Langmuir–Blodgett method by containing a smaller amount of an organic solvent because the presence of the organic solvent in the BLM causes denaturation of the membrane proteins. In the Langmuir–Blodgett method, BLMs are prepared by the apposition of two lipid monolayers spread at the air/water interface. The two monolayers are combined in a small aperture of a Teflon film with a thin layer of n-hexadecane or squalene ^[94]. In the droplet contact method, the lipid bilayer is formed by contacting two water droplets surrounding the lipid monolayer in the double well chip ^[63]. The painting method or the Langmuir–Blodgett method suffers from fragility and low reproducibility. Therefore, the use of high-throughput systems for pharmaceutical screenings of the BLM formed by these methods is difficult. Conversely, BLMs formed by the droplet contact method can be easily formed by simply dropping each of the solutions.

Figure 2. Illustration of BLM formation and the reconstituted ion channel. (a) Painting method. (b) Langmuir-Blodgett method. (c) Droplet contact method. (d) Vesicle Fusion method. (e) Nystatin/Ergosterol method.

5. Lipid Vesicles Containing Some Types of Membrane Proteins for Creating Complex Artificial Cell Models

To create artificial cell models that emulate the multiple functions of living cells, various membrane protein functions, such as molecular transportation, molecular conversion, and molecular production, must be reconstituted into lipid vesicles. Additionally, fluorescent probes or biomolecules such as DNA, water-soluble proteins, in vitro transcription-translation systems, and ions have been encapsulated into lipid vesicles. These complex artificial cell models, based on a bottom-up approach, have been constructed worldwide. In this section, the researchers introduce lipid vesicles containing two or more types of membrane proteins.

Ishmukhametov et al. ^[21] developed lipid vesicles reconstituted with ATP synthase and bo3-oxidase through fusion between cationic and anionic lipid vesicles containing either ATP-synthase or bo3-oxidase. This created a molecular bio-synthetic network linking the primary (proton motive force, PMF) and secondary ATP forms of biological free energy. Detergent removal using a Sephadex G-50 column reconstituted ATP synthase and bo3-oxidase into the lipid vesicles. When reduced dithiothreitol and Coenzyme Q₁ were added to the outer solution of the lipid vesicles, PMF was generated toward the inner solution because protons were also transferred to the inner solution. With PMF and KPi added to the inner solution, ATP was synthesized from ADP and pyrophosphate by ATP synthase in the outer solution. The ATP production of this molecular biosynthetic network was detected by luminescence using the luciferin-luciferase system.

Berhanu et al. ^[22] developed nano-sized lipid vesicles containing ATP synthase and BR for synthesizing proteins of an in vitro transcription–translation system using ATP produced from the nano-sized lipid vesicles by light. Detergent removal using bio-beads reconstituted ATP synthase and BR into the nano-sized lipid vesicles, which were then encapsulated into cell-sized lipid vesicles along with the in vitro transcription–translation system. When light was irradiated to the cell-sized lipid vesicles, BR on the nano-sized lipid vesicle membranes transported protons into the nano-sized lipid vesicles. By increasing the proton concentration into the inner solution of the nano-sized lipid vesicles, ATP-synthase converted ADP to ATP in the outer solution of the nano-sized lipid vesicles (in other words, into the inner solution of the cell-sized lipid vesicles). The produced ATP was consumed as substrates for messenger RNA, energy for phosphorylation of guanosine diphosphate, or energy for aminoacylation of transfer RNA.

Lee et al. ^[23] developed nano-sized lipid vesicles containing ATP synthase, PR, and plant-derived photosystem II (PSII) for controlling ATP production by changing the proton concentration and switching PR or PSII activity using red or green light. ATP synthase, PR, and PSII were reconstituted in the nano-sized lipid vesicles using a detergent-mediated method. The activation mechanism of the nano-sized lipid vesicles containing ATP-synthase, PR, and PSII was as follows: red light irradiated to the nano-sized lipid vesicles generated protons into the inner solution of the nano-sized lipid vesicles using PSII, which subsequently increased the proton concentration and caused ATP synthase to convert ADP to ATP in the outer solution. Conversely, green light caused protons to move from the inner to the outer solution of the nano-sized lipid vesicles, decreasing the proton concentration and halting ATP synthesis. Cell-sized lipid vesicles containing three types of membrane protein-reconstituted nano-sized lipid vesicles, ionophore, and actin experienced actin polymerizations and morphological changes depending on the ATP concentration generated from the nano-sized lipid vesicles.

References

Gilman, A.G. G proteins: Transducers of receptor-generated signals. Annu. Rev. Biochem. 1987, 56, 615–649.
Springer, T.A. Adhesion Receptors of the Immune System. Nature 1990, 346, 425–434.
Poitelon, Y.; Kopec, A.M.; Belin, S. Myelin Fat Facts: An Overview of Lipids and Fatty Acid Metabolism. Cells 2020, 9, 812.
Schenk, P.W.; Snaar-Jagalska, B.E. Signal Perception and Transduction: The Role of Protein Kinases. Biochim. Biophys. Acta Mol. Cell Res. 1999, 1449, 1–24.
Martin, T.F.J. Phosphoinositide Lipids as Signaling Molecules: Common Themes for Signal Transduction, Cytoskeletal Regulation, and Membrane Trafficking. Annu. Rev. Cell Dev. Biol. 1998, 14, 231–264.
Qin, J.; Guo, Y.; Xue, B.; Shi, P.; Chen, Y.; Su, Q.P.; Hao, H.; Zhao, S.; Wu, C.; Yu, L.; et al. ER-Mitochondria Contacts Promote MtDNA Nucleoids Active Transportation via Mitochondrial Dynamic Tubulation. Nat. Commun. 2020, 11, 4471.
Li, L.; Conradson, D.M.; Bharat, V.; Kim, M.J.; Hsieh, C.H.; Minhas, P.S.; Papakyrikos, A.M.; Durairaj, A.S.; Ludlam, A.; Andreasson, K.I.; et al. A Mitochondrial Membrane-Bridging Machinery Mediates Signal Transduction of Intramitochondrial Oxidation. Nat. Metab. 2021, 3, 1242–1258.
Raff, M.C. Social Controls on Cell Survival and Cell Death. Nature 1992, 356, 397–400.
Zhang, Y.; Chen, X.; Gueydan, C.; Han, J. Plasma Membrane Changes during Programmed Cell Deaths. Cell Res. 2018, 28, 9–21.
Han, Y.; Liu, D.; Li, L. PD-1/PD-L1 Pathway: Current Researches in Cancer. Am. J. Cancer Res. 2020, 10, 727–742.
Mizushima, N.; Klionsky, D.J. Protein Turnover via Autophagy: Implications for Metabolism. Annu. Rev. Nutr. 2007, 27, 19–40.
Congreve, M.; de Graaf, C.; Swain, N.A.; Tate, C.G. Impact of GPCR Structures on Drug Discovery. Cell 2020, 181, 81–91.
Davenport, A.P.; Scully, C.C.G.; de Graaf, C.; Brown, A.J.H.; Maguire, J.J. Advances in Therapeutic Peptides Targeting G Protein-Coupled Receptors. Nat. Rev. Drug Discov. 2020, 19, 389–413.
Lee, Y.; Warne, T.; Nehmé, R.; Pandey, S.; Dwivedi-Agnihotri, H.; Chaturvedi, M.; Edwards, P.C.; García-Nafría, J.; Leslie, A.G.W.; Shukla, A.K.; et al. Molecular Basis of β-Arrestin Coupling to Formoterol-Bound Β1-Adrenoceptor. Nature 2020, 583, 862–866.
Koivisto, A.P.; Belvisi, M.G.; Gaudet, R.; Szallasi, A. Advances in TRP Channel Drug Discovery: From Target Validation to Clinical Studies. Nat. Rev. Drug Discov. 2022, 21, 41–59.
Rosenbaum, M.I.; Clemmensen, L.S.; Bredt, D.S.; Bettler, B.; Strømgaard, K. Targeting Receptor Complexes: A New Dimension in Drug Discovery. Nat. Rev. Drug Discov. 2020, 19, 884–901.
He, S.; Moutaoufik, M.T.; Islam, S.; Persad, A.; Wu, A.; Aly, K.A.; Fonge, H.; Babu, M.; Cayabyab, F.S. HERG Channel and Cancer: A Mechanistic Review of Carcinogenic Processes and Therapeutic Potential. Biochim. Biophys. Acta Rev. Cancer 2020, 1873, 188355.
Pullman, M.E.; Penefsky, H.S.; Datta, A.; Racker, E. Partial Resolution of the Enzymes Catalyzing Oxidative Phosphorylation. J. Biol. Chem. 1960, 235, 3322–3329.
Racker, E. Reconstitution of a Calcium Pump with Phospholipids and a Purified Ca++—Adenosine Triphosphatase from Sacroplasmic Reticulum. J. Biol. Chem. 1972, 247, 8198–8200.
Keller, B.U.; Hedrich, R.; Vaz, W.L.C.; Criado, M. Single Channel Recordings of Reconstituted Ion Channel Proteins: An Improved Technique. Pflügers Arch. Eur. J. Physiol. 1988, 411, 94–100.
Ishmukhametov, R.R.; Russell, A.N.; Berry, R.M. A Modular Platform for One-Step Assembly of Multi-Component Membrane Systems by Fusion of Charged Proteoliposomes. Nat. Commun. 2016, 7, 13025.
Berhanu, S.; Ueda, T.; Kuruma, Y. Artificial Photosynthetic Cell Producing Energy for Protein Synthesis. Nat. Commun. 2019, 10, 1325.
Lee, K.Y.; Park, S.J.; Lee, K.A.; Kim, S.H.; Kim, H.; Meroz, Y.; Mahadevan, L.; Jung, K.H.; Ahn, T.K.; Parker, K.K.; et al. Photosynthetic Artificial Organelles Sustain and Control ATP-Dependent Reactions in a Protocellular System. Nat. Biotechnol. 2018, 36, 530–535.
Seddon, A.M.; Curnow, P.; Booth, P.J. Membrane Proteins, Lipids and Detergents: Not Just a Soap Opera. Biochim. Biophys. Acta Biomembr. 2004, 1666, 105–117.
Denisov, I.G.; Sligar, S.G. Nanodiscs in Membrane Biochemistry and Biophysics. Chem. Rev. 2017, 117, 4669–4713.
Dürr, U.H.N.; Gildenberg, M.; Ramamoorthy, A. The Magic of Bicelles Lights up Membrane Protein Structure. Chem. Rev. 2012, 112, 6054–6074.
Kim, H.J.; Howell, S.C.; Van Horn, W.D.; Jeon, Y.H.; Sanders, C.R. Recent Advances in the Application of Solution NMR Spectroscopy to Multi-Span Integral Membrane Proteins. Prog. Nucl. Magn. Reson. Spectrosc. 2009, 55, 335–360.
Hong, M.; Zhang, Y.; Hu, F. Membrane Protein Structure and Dynamics from NMR Spectroscopy. Annu. Rev. Phys. Chem. 2012, 63, 1–24.
Binder, D.; Drepper, T.; Jaeger, K.E.; Delvigne, F.; Wiechert, W.; Kohlheyer, D.; Grünberger, A. Homogenizing Bacterial Cell Factories: Analysis and Engineering of Phenotypic Heterogeneity. Metab. Eng. 2017, 42, 145–156.
Frank, J. Single-Particle Imaging of Macromolecules by Cryo-Electron Microscopy. Annu. Rev. Biophys. Biomol. Struct. 2002, 31, 303–319.
Fernandez-Leiro, R.; Scheres, S.H.W. Unravelling Biological Macromolecules with Cryo-Electron Microscopy. Nature 2016, 537, 339–346.
Hilgemann, D.W. Cytoplasmic ATP-Dependent Regulation of Ion Transporters and Channels: Mechanisms and Messengers. Annu. Rev. Physiol. 1997, 59, 193–220.
Darszon, A.; Labarca, P.; Nishigaki, T.; Espinosa, F. Ion Channels in Sperm Physiology. Physiol. Rev. 1999, 79, 481–510.
Wan, Y.; Zong, C.; Li, X.; Wang, A.; Li, Y.; Yang, T.; Bao, Q.; Dubow, M.; Yang, M.; Rodrigo, L.A.; et al. New Insights for Biosensing: Lessons from Microbial Defense Systems. Chem. Rev. 2021, 122, 8126–8180.
Misawa, N.; Osaki, T.; Takeuchi, S. Membrane Protein-Based Biosensors. J. R. Soc. Interface 2018, 15, 20170952.
Misawa, N.; Fujii, S.; Kamiya, K.; Osaki, T.; Takaku, T.; Takahashi, Y.; Takeuchi, S. Construction of a Biohybrid Odorant Sensor Using Biological Olfactory Receptors Embedded into Bilayer Lipid Membrane on a Chip. ACS Sens. 2019, 4, 711–716.
Yamada, T.; Sugiura, H.; Mimura, H.; Kamiya, K.; Osaki, T.; Takeuchi, S. Highly Sensitive VOC Detectors Using Insect Olfactory Receptors Reconstituted into Lipid Bilayers. Sci. Adv. 2021, 7, 2013–2026.
Bangham, A.D.; Horne, R.W. Negative Staining of Phospholipids and Their Structural Modification by Surface-Active Agents as Observed in the Electron Microscope. J. Mol. Biol. 1964, 8, 660–668.
Akashi, K.I.; Miyata, H.; Itoh, H.; Kinosita, K. Preparation of Giant Liposomes in Physiological Conditions and Their Characterization under an Optical Microscope. Biophys. J. 1996, 71, 3242–3250.
Horger, K.S.; Estes, D.J.; Capone, R.; Mayer, M. Films of Agarose Enable Rapid Formation of Giant Liposomes in Solutions of Physiologic Ionic Strength. J. Am. Chem. Soc. 2009, 131, 1810–1819.
Lira, R.B.; Dimova, R.; Riske, K.A. Giant Unilamellar Vesicles Formed by Hybrid Films of Agarose and Lipids Display Altered Mechanical Properties. Biophys. J. 2014, 107, 1609–1619.
Angelova, M.I.; Dimitrov, D.S. Liposome Electroformation. Faraday Discuss. Chem. Soc. 1986, 81, 303–311.
Dimitrov, D.S.; Angelova, M.I. Lipid Swelling and Liposome Formation Mediated by Electric Fields. Bioelectrochem. Bioenerg. 1988, 19, 323–336.
Pautot, S.; Frisken, B.J.; Weitz, D.A. Production of Unilamellar Vesicles Using an Inverted Emulsion. Langmuir 2003, 19, 2870–2879.
Pautot, S.; Frisken, B.J.; Weitz, D.A. Engineering Asymmetric Vesicles. Proc. Natl. Acad. Sci. USA 2003, 100, 10718–10721.
Tan, Y.C.; Hettiarachchi, K.; Siu, M.; Pan, Y.R.; Lee, A.P. Controlled Microfluidic Encapsulation of Cells, Proteins, and Microbeads in Lipid Vesicles. J. Am. Chem. Soc. 2006, 128, 5656–5658.
Hu, P.C.; Li, S.; Malmstadt, N. Microfluidic Fabrication of Asymmetric Giant Lipid Vesicles. ACS Appl. Mater. Interfaces 2011, 3, 1434–1440.
Matosevic, S.; Paegel, B.M. Stepwise Synthesis of Giant Unilamellar Vesicles on a Microfluidic Assembly Line. J. Am. Chem. Soc. 2011, 133, 2798–2800.
Abkarian, M.; Loiseau, E.; Massiera, G. Continuous Droplet Interface Crossing Encapsulation (CDICE) for High Throughput Monodisperse Vesicle Design. Soft Matter 2011, 7, 4610–4614.
Karamdad, K.; Law, R.V.; Seddon, J.M.; Brooks, N.J.; Ces, O. Preparation and Mechanical Characterisation of Giant Unilamellar Vesicles by a Microfluidic Method. Lab Chip 2015, 15, 557–562.
Karamdad, K.; Law, R.V.; Seddon, J.M.; Brooks, N.J.; Ces, O. Studying the Effects of Asymmetry on the Bending Rigidity of Lipid Membranes Formed by Microfluidics. Chem. Commun. 2016, 52, 5277–5280.
Blosser, M.C.; Horst, B.G.; Keller, S.L. CDICE Method Produces Giant Lipid Vesicles under Physiological Conditions of Charged Lipids and Ionic Solutions. Soft Matter 2016, 12, 7364–7371.
Romanov, V.; McCullough, J.; Gale, B.K.; Frost, A. A Tunable Microfluidic Device Enables Cargo Encapsulation by Cell- or Organelle-Sized Lipid Vesicles Comprising Asymmetric Lipid Bilayers. Adv. Biosyst. 2019, 3, 1900010.
Shum, H.C.; Lee, D.; Yoon, I.; Kodger, T.; Weitz, D.A. Double Emulsion Templated Monodisperse Phospholipid Vesicles. Langmuir 2008, 24, 7651–7653.
Arriaga, L.R.; Datta, S.S.; Kim, S.H.; Amstad, E.; Kodger, T.E.; Monroy, F.; Weitz, D.A. Ultrathin Shell Double Emulsion Templated Giant Unilamellar Lipid Vesicles with Controlled Microdomain Formation. Small 2014, 10, 950–956.
Kim, S.H.; Kim, J.W.; Cho, J.C.; Weitz, D.A. Double-Emulsion Drops with Ultra-Thin Shells for Capsule Templates. Lab Chip 2011, 11, 3162–3166.
Caschera, F.; Lee, J.W.; Ho, K.K.Y.; Liu, A.P.; Jewett, M.C. Cell-Free Compartmentalized Protein Synthesis inside Double Emulsion Templated Liposomes with In Vitro Synthesized and Assembled Ribosomes. Chem. Commun. 2016, 52, 5467–5469.
Deshpande, S.; Caspi, Y.; Meijering, A.E.C.; Dekker, C. Octanol-Assisted Liposome Assembly on Chip. Nat. Commun. 2016, 7, 10447.
Deng, N.-N.; Yelleswarapu, M.; Huck, W.T.S. Monodisperse Uni- and Multicompartment Liposomes. J. Am. Chem. Soc. 2016, 138, 7584–7591.
Deng, N.; Huck, W.T.S. Microfluidic Formation of Monodisperse Coacervate Organelles in Liposomes. Angew. Chem. Int. Ed. Engl. 2017, 56, 9736–9740.
Michelon, M.; Huang, Y.; de la Torre, L.G.; Weitz, D.A.; Cunha, R.L. Single-Step Microfluidic Production of W/O/W Double Emulsions as Templates for β-Carotene-Loaded Giant Liposomes Formation. Chem. Eng. J. 2019, 366, 27–32.
Ushiyama, R.; Koiwai, K.; Suzuki, H. Plug-and-Play Microfluidic Production of Monodisperse Giant Unilamellar Vesicles Using Droplet Transfer across Water–Oil Interface. Sens. Actuators B Chem. 2022, 355, 131281.
Funakoshi, K.; Suzuki, H.; Takeuchi, S. Lipid Bilayer Formation by Contacting Monolayers in a Microfluidic Device for Membrane Protein Analysis. Anal. Chem. 2006, 78, 8169–8174.
Kamiya, K.; Kawano, R.; Osaki, T.; Akiyoshi, K.; Takeuchi, S. Cell-Sized Asymmetric Lipid Vesicles Facilitate the Investigation of Asymmetric Membranes. Nat. Chem. 2016, 8, 881–889.
Kamiya, K.; Arisaka, C.; Suzuki, M. Investigation of Fusion between Nanosized Lipid Vesicles and a Lipid Monolayer toward Formation of Giant Lipid Vesicles with Various Kinds of Biomolecules. Micromachines 2021, 12, 133.
Kamiya, K.; Osaki, T.; Takeuchi, S. Formation of Nano-Sized Lipid Vesicles with Asymmetric Lipid Components Using a Pulsed-Jet Flow Method. Sens. Actuators B Chem. 2021, 327, 128917.
Gotanda, M.; Kamiya, K.; Osaki, T.; Fujii, S.; Misawa, N.; Miki, N.; Takeuchi, S. Sequential Generation of Asymmetric Lipid Vesicles Using a Pulsed-Jetting Method in Rotational Wells. Sens. Actuators B Chem. 2018, 261, 392–397.
Gotanda, M.; Kamiya, K.; Osaki, T.; Miki, N.; Takeuchi, S. Automatic Generation System of Cell-Sized Liposomes. Sens. Actuators B Chem. 2019, 292, 57–63.
Kamiya, K.; Osaki, T.; Takeuchi, S. Formation of Vesicles-in-a-Vesicle with Asymmetric Lipid Components Using a Pulsed-Jet Flow Method. RSC Adv. 2019, 9, 30071–30075.
Elani, Y.; Law, R.V.; Ces, O. Vesicle-Based Artificial Cells as Chemical Microreactors with Spatially Segregated Reaction Pathways. Nat. Commun. 2014, 5, 5305.
Miwa, A.; Kamiya, K. Control of Enzyme Reaction Initiation inside Giant Unilamellar Vesicles by the Cell-Penetrating Peptide-Mediated Translocation of Cargo Proteins. ACS Synth. Biol. 2022, 11, 3836–3846.
Kamiya, K. Development of Artificial Cell Models Using Microfluidic Technology and Synthetic Biology. Micromachines 2020, 11, 559.
Kamiya, K.; Takeuchi, S. Giant Liposome Formation toward the Synthesis of Well-Defined Artificial Cells. J. Mater. Chem. B 2017, 5, 5911–5923.
Dezi, M.; Di Cicco, A.; Bassereau, P.; Lévy, D. Detergent-Mediated Incorporation of Transmembrane Proteins in Giant Unilamellar Vesicles with Controlled Physiological Contents. Proc. Natl. Acad. Sci. USA 2013, 110, 7276–7281.
Fenz, S.F.; Sachse, R.; Schmidt, T.; Kubick, S. Cell-Free Synthesis of Membrane Proteins: Tailored Cell Models out of Microsomes. Biochim. Biophys. Acta Biomembr. 2014, 1838, 1382–1388.
Suzuki, M.; Kamiya, K. Cell-Sized Asymmetric Phospholipid-Amphiphilic Protein Vesicles with Growth, Fission, and Molecule Transportation. iScience 2023, 26, 106086.
Ohnishi, S.; Kamiya, K. Formation of Giant Lipid Vesicle Containing Dual Functions Facilitates Outer Membrane Phospholipase. ACS Synth. Biol. 2021, 10, 1837–1846.
Eaglesfield, R.; Madsen, M.A.; Sanyal, S.; Reboud, J.; Amtmann, A. Cotranslational Recruitment of Ribosomes in Protocells Recreates a Translocon-Independent Mechanism of Proteorhodopsin Biogenesis. iScience 2021, 24, 102429.
Altamura, E.; Milano, F.; Tangorra, R.R.; Trotta, M.; Omar, O.H.; Stano, P.; Mavelli, F. Highly Oriented Photosynthetic Reaction Centers Generate a Proton Gradient in Synthetic Protocells. Proc. Natl. Acad. Sci. USA 2017, 114, 3837–3842.
Kajii, K.; Shimomura, A.; Higashide, M.T.; Oki, M.; Tsuji, G. Effects of Sugars on Giant Unilamellar Vesicle Preparation, Fusion, PCR in Liposomes, and Pore Formation. Langmuir 2022, 38, 8871–8880.
Diederichs, T.; Tampé, R. Single Cell-like Systems Reveal Active Unidirectional and Light-Controlled Transport by Nanomachineries. ACS Nano 2021, 15, 6747–6755.
Varnier, A.; Kermarrec, F.; Blesneac, I.; Moreau, C.; Liguori, L.; Lenormand, J.L.; Picollet-D’Hahan, N. A Simple Method for the Reconstitution of Membrane Proteins into Giant Unilamellar Vesicles. J. Membr. Biol. 2010, 233, 85–92.
Soga, H.; Fujii, S.; Yomo, T.; Kato, Y.; Watanabe, H.; Matsuura, T. In Vitro Membrane Protein Synthesis inside Cell-Sized Vesicles Reveals the Dependence of Membrane Protein Integration on Vesicle Volume. ACS Synth. Biol. 2014, 3, 372–379.
Almendro-Vedia, V.G.; Natale, P.; Mell, M.; Bonneau, S.; Monroy, F.; Joubert, F.; López-Montero, I. Nonequilibrium Fluctuations of Lipid Membranes by the Rotating Motor Protein F1F0-ATP Synthase. Proc. Natl. Acad. Sci. USA 2017, 114, 11291–11296.
Motta, I.; Gohlke, A.; Adrien, V.; Li, F.; Gardavot, H.; Rothman, J.E.; Pincet, F. Formation of Giant Unilamellar Proteo-Liposomes by Osmotic Shock. Langmuir 2015, 31, 7091–7099.
Jang, H.S.; Cho, Y.K.; Granick, S. Biologically-Active Unilamellar Vesicles from Red Blood Cells. Biomater. Sci. 2019, 7, 1393–1398.
Guo, X.; Steinkühler, J.; Marin, M.; Li, X.; Lu, W.; Dimova, R.; Melikyan, G.B. Interferon-Induced Transmembrane Protein 3 Blocks Fusion of Diverse Enveloped Viruses by Altering Mechanical Properties of Cell Membranes. ACS Nano 2021, 15, 8155–8170.
Yanagisawa, M.; Iwamoto, M.; Kato, A.; Yoshikawa, K.; Oiki, S. Oriented Reconstitution of a Membrane Protein in a Giant Unilamellar Vesicle: Experimental Verification with the Potassium Channel KcsA. J. Am. Chem. Soc. 2011, 133, 11774–11779.
Aimon, S.; Manzi, J.; Schmidt, D.; Larrosa, J.A.P.; Bassereau, P.; Toombes, G.E.S. Functional Reconstitution of a Voltage-Gated Potassium Channel in Giant Unilamellar Vesicles. PLoS ONE 2011, 6, 25529.
dos Santos, E.C.; Belluati, A.; Necula, D.; Scherrer, D.; Meyer, C.E.; Wehr, R.P.; Lörtscher, E.; Palivan, C.G.; Meier, W. Combinatorial Strategy for Studying Biochemical Pathways in Double Emulsion Templated Cell-Sized Compartments. Adv. Mater. 2020, 32, 2004804.
Neher, E.; Sakmann, B. Single-Channel Currents Recorded from Membrane of Denervated Frog Muscle Fibres. Nature 1976, 260, 799–802.
Hirano-Iwata, A.; Ishinari, Y.; Yamamoto, H.; Niwano, M. Micro- and Nano-Technologies for Lipid Bilayer-Based Ion-Channel Functional Assays. Chem. Asian J. 2015, 10, 1266–1274.
Mueller, P.; Rudin, D.O.; Ti Tien, H.; Wescott, W.C. Reconstitution of Cell Membrane Structure In Vitro and Its Transformation into an Excitable System. Nature 1962, 194, 979–980.
Montal, M.; Mueller, P. Formation of Bimolecular Membranes from Lipid Monolayers and a Study of Their Electrical Properties. Proc. Natl. Acad. Sci. USA 1972, 69, 3561–3566.

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Subjects: Biophysics; Biotechnology & Applied Microbiology; Biochemistry & Molecular Biology

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Toshiyuki Tosaka

Koki Kamiya

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Update Date: 04 May 2023

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