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1Department of Physiology and Biophysics, University of Washington
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Reconstituting functional membrane proteins into giant liposomes of defined composition is a powerful approach when combined with patch-clamp electrophysiology. However, conventional giant liposome production may be incompatible with protein stability. We describe protocols for producing giant liposomes from pure lipids or small liposomes containing ion channels.
Keywords: Physiology, Issue 76, Biophysics, Molecular Biology, Biochemistry, Genetics, Cellular Biology, Proteins, Membranes, Artificial, Lipid Bilayers, Liposomes, Phospholipids, biochemistry, Lipids, Giant Unilamellar Vesicles, liposome, electrophysiology, electroformation, reconstitution, patch clamp
Collins, M. D., Gordon, S. E. Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology. J. Vis. Exp. (76), e50227, doi:10.3791/50227 (2013).
The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes >10 μm in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.
Giant liposomes (often called giant unilamellar vesicles, or GUVs) have primarily been used to study the physics and physical chemistry of lipid bilayers, including studies of bilayer deformation, lateral phase coexistence ("rafts"), membrane fusion, etc1-4. They have a grossly cell-like structure: spherical shell of membrane surrounding an aqueous interior which can easily be made different than the surrounding aqueous buffer. They are, by definition, ≈1-100 μm in diameter, so they can be imaged using a variety of light microscopy approaches. They can be made taut using osmotic gradients or mechanically applied tension, so that while generally soft, their properties can be manipulated for easy handling. In particular, controlling the "stiffness" of the liposome makes it straightforward to form "liposome-attached" or excised patches for electrophysiology. In the past, ion channel reconstitution was largely performed in planar lipid bilayers. Now, the ability to form patches from giant liposomes and use the considerable quiver of tools developed for conventional electrophysiology (fluorescence microscopy, micropipette aspiration, rapid perfusion and temperature control, etc.) makes giant liposomes increasingly attractive for reconstitution studies5,6.
Giant liposomes have been made by many strategies. In fact, giant liposomes form spontaneously by a swelling process when a dried lipid film is rehydrated4,7,8. The desire to more rapidly prepare larger, more uniform liposomes led researchers to other approaches, chief among them electroformation1,9. Electroformation also relies on hydration of a dried lipid film, but speeds the process through the application of an oscillating electric field across the lipid film. The field is applied through two electrodes, either platinum wires or Indium-Tin-Oxide (ITO) coated glass slides, separated by water or buffer and onto which the lipids are deposited. By speeding the swelling of liposomes, one achieves a higher yield of larger liposomes. Thus, electroformation has become the default method to produce giant liposomes4.
The mechanism of electroformation is not fully understood, and most of the protocols are developed empirically (e.g. 10,11). Nonetheless, we can learn a little about what to expect by considering the theory and some empirical results. It is widely believed that electroformation occurs by driving electro-osmotic flow of buffer between individual lipid bilayers stacked in the deposited lipid film10,11. Electrostatic coupling to thermal fluctuations of the lipid bilayers is probably also involved12. These hypotheses qualitatively predict upper limits for the electric field frequency and strength that can be used10,12. In particular, it is predicted that high conductivity solutions (i.e. physiological salt solutions) reduce the electrohydrodynamic forces that may initiate the liposome electroformation12. Electroosmotic flow rates generally decrease with increasing salt concentration and are frequently peaked at some electric field oscillation frequency (e.g. albeit in a different geometry, Green et al.13). Thus, higher field strengths and higher frequencies are reasonable for high conductivity solutions, within limits10.
However, membrane proteins are likely to be incompatible with the usual method of depositing lipids onto electrodes for the electroswelling procedure, namely in organic solvents which are then evaporated off to leave a thin lipid film. There are two principal paths around this difficulty: to incorporate proteins after giant liposome formation, or to adapt how the lipids are deposited. Our approach builds on others5,11 to deposit the lipids and reconstituted membrane protein together from a suspension of small or large "proteoliposomes". We describe the lengthy and more challenging process of producing proteoliposomes from purified protein and lipids elsewhere (Collins and Gordon, in review). Here we describe the protocol in the absence of any protein, but it is the same when protein is incorporated; we include results showing that proteoliposomes containing the ion channel TRPV1 can be transformed into GUVs and used for patch-clamp electrophysiology. In any electroformation approach, visual inspection of the lipid sample during the lipid deposition process is critical to success.
Our approach may be relevant beyond the specialized application to ion channel reconstitution. In the time since we first developed this protocol and now, it has also been shown how the way in which lipids are deposited on electrodes for electroformation affects the compositional heterogeneity of the resulting GUVs. Baykal-Caglar et al.14 showed that GUVs formed from carefully dehydrated liposomes had a 2.5 times smaller variation in the miscibility transition temperature of GUVs formed from mixtures of various phospholipids and cholesterol. Their work indicates that lipids, and especially cholesterol, may precipitate from the lipid mixture when deposited from organic solvents, resulting in large spatial variation in composition of the deposited lipid film. This is especially important for studies of lipid membrane phase behavior, but may also be critical to quantitative experiments on ion channel function. Baykal-Caglar et al.'s protocol is similar but not identical to our own, and readers are encouraged to study it as well.
This protocol (see overview, Figure 1) is one of many that could be used. In principle electroformation success depends on the lipid mixture, hydration, temperature, other solutes (especially ions), and of course the voltage and frequency used in formation. As electroformation becomes better understood, we expect to refine our protocol more.
Finally, there is often a steep learning curve in electroforming giant liposomes. We suggest mastering the conventional protocol (Sections 1 and 4, and, if necessary, Section 5) before learning to deposit lipids from liposomal suspensions (Sections 2-5).
1. Deposition of Lipids from Organic Solvents: Classical Protocol
2. Small Liposome Preparation for Controlled Dehydration
3. Deposition of Lipids by Controlled Dehydration of Small Liposomes
4. Electroformation of Giant Liposomes
5. Imaging and Troubleshooting
In our examples, we prepare liposomes from a mixture of approximately 55 mol% POPC (1-palmitoyl-2-oleoyl-sn-glycero-phosphocholine), 15 mol% POPS (1-palmitoyl-2-oleoyl-sn-glycero-phosphoserine, 30 mol% cholesterol, and 0.1 mol% Texas Red-labeled 1,2-dipalmitoyl-sn-phosphoethanolamine (TxR-DPPE). This composition was chosen as approximately representative of dorsal root ganglion lipids18. We note that 15 mol% charged lipid (here POPS) is near the limit of what can be used in electroformation (e.g. Walde et al.4).
Lipid breakdown by hydrolysis and peroxidation is an especially important issue for many lipid mixtures and many sensitive experiments (see Discussion). For clarity alone, we do not use inert gas atmospheres in the video presentation. As is evident from the figures below, this does not noticeably impact the quality of the liposomes, but for sensitive studies or when using highly unsaturated lipids it is important to prevent lipid breakdown.
Since the major hurdle in electroformation is to form a good lipid film on the Indium-Tin Oxide (ITO) substrate, knowing what the lipid film should look like when dried is crucial. However, when electroformation fails, it often does so completely, resulting in no appreciable giant liposome production. Frequently, substantial detritus will be observed; this typically indicates that too much lipid has been used. Our figures show what one expects to see upon success.
Figure 2 shows two different patches of liposomes electroformed from lipids deposited out of chloroform. In the left panel, closely examine the patches indicated by arrows 2 and 3. While arrow 2 indicates a patch with poorly distinguishable liposomes, no liposomes can be brought into focus in the area indicated by arrow 3. Together, these findings indicate a generally poor preparation, probably due to there being too much lipid deposited in this area. In the right panel, such "fuzzy" regions are less common, indicating a generally better preparation. We find that it is rare to completely eliminate such imperfections.
Figure 3 shows liposomes successfully electroformed from lipids deposited by dehydration of small liposomes, at two magnifications, imaged using Texas Red epifluorescence. Liposomes are sparse, but there are several good specimens. Because size varies, several are out of focus. The arrows indicate three good quality liposomes ranging in size from ~5-20 μm, two of which appear in the 40x magnification image. Note the clear "ring" at the edge: this indicates a unilamellar or nearly unilamellar liposome. While some spots of lipid appear to have collapsed or never formed lipid, this is a good quality result.
Finally, our purpose in developing and publishing this protocol is to prepare GUVs with intact, functional mammalian ion channels that are suitable for conventional patch-clamp electrophysiology. In Figure 4 we demonstrate capsaicin-activated TRPV1 ionic currents recorded in lipid membrane patches excised from GUVs formed using this protocol. The current can be blocked by Ruthenium Red, and is not activated by the capsaicin vehicle (0.1% ethanol, 138 mM NaCl, 3 mM HEPES pH 7.4) alone. The preparation of the proteoliposomes for electrophysiology experiments is far beyond the scope of the present protocol, but is the subject of a submitted article.
|Salt||Molality @ 20 °C and saturation||%RH @ 10 °C||%RH @ 20 °C||%RH @ 30 °C|
|Magnesium Chloride Hexahydrate||5.8||33||33||32|
|Potassium Carbonate Dihydrate||8.0||47||44||42|
Table 1. Relative humidity (%RH) of several common salts. Relative humidity data from Greenspan19 and Rockland20; saturation data from the International Critical Tables21.
Figure 1. Schematic of giant liposome electroformation from small liposomes. (Top left) Small liposomes are deposited in an array of small droplets, < 5 μl. (Top center) The liposomes are dehydrated under controlled relative humidity by placing them in a sealed container above a saturated salt solution. A hygrometer is optional. (Top right) Once dehydrated to a sticky film of lipid and (possibly) osmoticant such as sorbitol, the lipids are rehydrated in a hyperosmotic buffer (see protocol section 5). (Lower left) the electroformation chamber is sealed with a second ITO slide on top. (Lower right) Finally, the chamber is heated above the lipid chain melting temperature, and connected to a signal source providing an oscillating electric field across the two ITO slides.
Figure 2. Electroformed giant liposomes formed from solvent-deposited lipids. (Left) Poor preparations have a few good quality liposomes (1) many more regions containing small, barely discernible liposomes, and many "fuzzy" regions with no discernible liposome formation. (2). (Right) While good preparations will still contain some regions of poorly discernible liposomes or "fuzz", there are many large liposomes (1 and 2). Imaged using an inverted epifluorescence microscope equipped with a Chroma 41004 Texas Red filter cube and Stanford Photonics XR/MEGA-10 S30 intensified CCD camera.
Figure 3. Electroformed giant liposomes formed from dehydration-deposited lipids. (Left) At 10x magnification, several liposomes are visible. Three liposomes are labeled. (right) The same view as at left, at 40x magnification, with the same liposomes (2 and 3) labeled. The number of these liposomes is typically much smaller than when electroforming from solvent-deposited lipid, but is entirely adequate for many purposes. Imaged using an inverted epifluorescence microscope equipped with a Chroma 41004 Texas Red filter cube and Stanford Photonics XR/MEGA-10 S30 intensified CCD camera.
Figure 4. Capsaicin activated TRPV1 current recorded in a membrane patch excised from a GUV formed using this protocol. A. The leak current indicated a 500 MΩ seal resistance before capsaicin was added. B. Saturating capsaicin (>20 μM, in 0.1% ethanol) activates a large TRPV1 current; the current is blocked by Ruthenium Red, and is not activated by vehicle (0.1% ethanol in buffer) alone (data not shown). C. The current returns to near baseline as capsaicin is washed out of the patch.
Figure 5. Plans for electroformation chamber. Included is an overview, showing the complete assembly, and dimensioned schematics for the top and bottom plastic pieces. Use acrylic or another easily machined clear plastic. The thin Kapton film heater should be connected to the temperature controller (see Table of specific reagents and equipment), and the EMI gasket foam should be connected to the function generator, so that the sine wave signal is applied across the two ITO coated slides. A hole is provided for a temperature probe. Flathead 10-32 screws are placed in the countersunk holes in the bottom piece, and pass through the upper piece. 10-32 nuts are used to secure the assembly. When fully assembled, the top and bottom plastic pieces should be flush with each other on two sides (the sides parallel to the EMI gasket.) Click here to view larger figure.
Electroformation of giant liposomes has developed into a flexible technique compatible with diverse lipids, preparations, and buffers. Careful control of the lipid deposition process is most critical to success. We have presented simple tools to make controlled deposition of lipids from small liposome preparations a straightforward process. The relative humidity is critical to proper dehydration of the initial liposomes, and the optimum value will vary with the initial concentration of solutes in the liposome suspension. Lower relative humidity is required to dehydrate more concentrated samples to an adequate level.
The exact hydration protocol and electric field used for electroformation remains a point of discussion1,4,7,10,11,22. The simplest protocols should be mastered first before attempting liposome electroformation in the presence of high salt concentrations, charged lipids, or very high concentrations of high melting temperature lipids. Once those skills are mastered, more advanced protocols typically divide the electroformation process into three parts: initial swelling, growth, and detachment. Initial swelling seems to be favored by slowly increasing the applied electric field11. A second, optional stage at fixed field strength can help control the final size of the liposomes. Finally, decreasing the frequency of the oscillating field may help to detach liposomes from the ITO surface. Higher frequencies are needed to electroform liposomes in physiological salt conditions11, but liposomes can be formed over an extremely wide range of field frequencies and amplitudes10.
The observation that giant liposomes will form spontaneously upon rehydration of dehydrated liposomes with a hypo-osmotic buffer7 indicates several keys to success. First, the liposome swelling process begins immediately, so that for electroformation, some groups have even found it useful to apply the AC electric field before rehydrating the lipid film10. Second, it is likely that fluid flow drives the liposomal swelling process, and that electroformation occurs at least in part because of electro-osmotically driven flow. This places limits on the useful frequency and amplitude range that can be used in electroformation10.
A major point of discussion in the last several years has been the potential for lipid hydrolysis and peroxidation23,24. The first defense against lipid degradation of any kind is to remove oxygen from all materials used in the GUV preparation: since peroxidation depends on molecular oxygen23, this should slow the process considerably. A combination of vacuum and sparging with inert gas should be used to remove as much oxygen as possible15-17. Sonication and light vacuum are ineffective. We use an oxygen test kit (Chemetrics K7501) to verify that we have removed oxygen as thoroughly as possible. Hydrolysis is more pernicious, but is reduced by maintaining neutral pH25, and by reducing the voltage used in electroformation24. Some authors advocate the use of titanium electrodes in place of ITO slides23,24,26,27. We prefer to avoid the possibility of contaminating our samples with titanium (or any metal), since it has in the past been known to interfere with liposome experiments28,29, but it might help to prevent some peroxidation or hydrolysis effects. It goes without saying that prolonged exposure to high temperatures accelerates lipid breakdown30. Finally, thin-layer chromatography24,31, colorimetric assays23, and other methods31 exist to quantify lipid degradation.
A similar issue relates to the fluorescent lipid dyes used to image GUVs, especially for studies of lateral phase separation30,32. In sensitive experiments, dyes should be chosen carefully30,33, and used at minimal concentration.
We are not aware of any consensus on whether the ITO electrode slides can be reused. Many groups do reuse their ITO slides, while some do not. Recent work indicates that the slides do degrade over time, but can be annealed to regain high performance34; this degradation had only a small effect for lipid mixtures including zwitterionic or anionic lipids, but was critical for mixtures containing cationic lipids. We reuse our slides without annealing.
Although there is great variability in the protocols used to electroform giant liposomes, the theoretical understanding and practical experience with the technique is constantly improving. Already liposomes can be formed with large amounts of charged lipids, or in high salt buffers. The key remains efficient deposition of the lipids on the electrode surface, which is the core of our protocol.
The authors declare that they have no competing financial interests.
We thank Bryan Venema and Eric Martinson for constructing the electroformation apparatus. This work was funded by grants from the National Institutes of General Medical Sciences of the National Institutes of Health (R01GM100718 to SEG) and the National Eye Institute of the National Institutes of Health (R01EY017564 to SEG).
|Digital Multimeter||Agilent Technologies, www.agilent.com||U1232A or similar||Any multimeter will do, but avoid old style analog ohmmeters which apply much more current to the resistance under test.|
|Fluke||117 or 177||Any multimeter will do, but avoid old style analog ohmmeters which apply much more current to the resistance under test.|
|Function Generator||Agilent Technologies, www.agilent.com||33210A or similar||Most function generators work for simple protocols. This programmable model is useful for advanced electroformation protocols. Make sure the generator can drive 10 V peak-to-peak into a 50 Ω load|
|ITO coated glass slides||Delta Technologies, Loveland, CO www.delta-technologies.com||CB-90IN-S107 or similar||Break these in half to make two slides, 25 mm x 37 mm|
|Temperature controller||Omega Engineering Stamford, CT www.omega.com||CNi3233 or similar|
|Hygrometer||Extech, Nashua, NH, www.extech.com||445815|
|Silicone rubber sheet||McMaster-Carr Elmhurst, IL www.mcmaster.com||87315K64||Use USP Grade VI silicone for its high purity|
|EMI gasket||Laird Technologies www.lairdtech.com||4202-PA-51H-01800 or similar||Distributed by Mouser www.mouser.com|
|TxR-DHPE||Life Technologies, Carlsbad, CA www.lifetechnologies.com||T1395MP||Other fluorescently labeled lipids are available, but TxR-DHPE is one of the brightest and most photostable.|
|POPC||Avanti Polar Lipids, Alabaster, AL www.avantilipids.com||850457P or 850457C||Lipids can be ordered as powders (P) or in chloroform (C)|
|POPS||Avanti Polar Lipids||840034P/C|
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