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).

1. Deposition of Lipids from Organic Solvents: Classical Protocol
<ol>
	<li>Remove lipids from storage at -20 &deg;C or -80 &deg;C; warm to RT. Caution: lipids are extremely hygroscopic, and many are sensitive to oxygen. Cover lipids in dry Argon or Nitrogen gas and in all steps minimize exposure to air.</li>
	<li>If necessary, suspend the lipids in chloroform or cyclohexane at 1-10 mg/ml; note that manufacturer's stated concentrations are typically nominal only. CAUTION: Wear appropriate gloves and other personal protective equipment when using organic solvents. Remove the PPE quickly if solvent is spilled on them, as most materials are still permeable to these solvents and they only provide temporary protection.</li>
	<li>Clean both sides of two 25 x 37.5 mm ITO coated glass slides with ethanol</li>
	<li>Mix the lipids to achieve the desired molar ratios. For each 1 cm2 of area of ITO slides to be coated with lipid, mix ~15-20 &mu;g of lipids. For example, if both 25 x 37.5 mm slides were to be fully coated, we would typically use 30 &mu;l of 10 mg/ml lipid mixture. Note: add 0.1 mol% Texas Red-DPPE for fluorescent imaging, and see below for cautions about fluorescent dyes.</li>
	<li>Verify that the ITO coated side of the glass slides is facing up by measuring the surface resistance with an ohmmeter or multimeter.</li>
	<li>Aspirate the lipids into a solvent-resistant syringe. Caution: No glues or plastics, other than PTFE, should contact the organic solvent.</li>
	<li>With the syringe needle not quite touching the ITO surface, slowly apply the lipid moving the needle back and forth across the slide. Cover the surface as evenly, looking for a "rainbow sheen" on the glass surface.</li>
	<li>Place the slides quickly under &lt; 1 Torr (1 mmHg) vacuum for 0.5-1 hr to remove any trace solvent. Release vacuum with inert gas.</li>
	<li>Apply a silicone gasket, with a thin layer of silicone vacuum grease on both sides. Leave at least 5 mm of uncovered slide exposed at one end of the slide.</li>
	<li>Proceed immediately to Section 4.</li>
</ol>
2. Small Liposome Preparation for Controlled Dehydration
<ol>
	<li>Prepare the lipid mixture as in Steps 1.1 to 1.4.</li>
	<li>Dry the lipids using a stream of Argon or Nitrogen gas in a 10-15 mm diameter culture tube. For small quantities of lipids, &le;0.5 mg, the resulting film can be hydrated directly after placing it under vacuum for 0.5-1 hr to remove residual solvent; proceed to step 2.5. Larger amounts of lipid tend to trap organic solvents in a thick gel, even under vacuum, and are better prepared by lyophilization; see steps 2.3-4.</li>
	<li>Suspend the dried lipid film in cyclohexane, cover with Argon and seal with a stopper, place the tube in a cold block and freeze at -80 &deg;C for 1 hr minimum.</li>
	<li>Place the cold block and lipid sample in a vacuum system capable of reaching 100 mTorr. Apply vacuum; it will "stall" at about 1 Torr as solvent sublimes off. Once solvent is completely removed (~1 hr) vacuum will drop below this level. Release vacuum with inert gas and seal the tube or hydrate immediately.</li>
	<li>While the lipids are under vacuum, degas hydration buffer using vacuum and sparging15-17.</li>
	<li>Hydrate the lipids using degassed buffer at final concentration 1-10 mg/ml. Allow the lipids to hydrate slowly. After 30 min-1 hr, vortex to break up remaining clumps. Wait an additional 0.5-1 hr to allow for complete hydration. Use an osmoticant, such as sorbitol or sucrose up to 200 mM, to prevent complete dehydration in steps below.</li>
	<li>Prepare 100-200 nm diameter liposomes by extrusion. See manufacturer's extrusion protocol for details. Note that the buffer must have less than ~25 mM ionic strength, or special electroformation voltage protocols will be needed in Section 4.</li>
</ol>
3. Deposition of Lipids by Controlled Dehydration of Small Liposomes
<ol>
	<li>Caution: Our protocol requires lipids to be placed in vapor equilibrium with saturated salt solutions for many hours. We strongly recommend performing the protocol in an inert gas atmosphere, using a glove box or similar enclosure.</li>
	<li>Caution: If preparing samples containing proteins, avoid complete dehydration. Hydrate the atmosphere in any enclosure with a beaker of warm water, or a sonicator based humidifier. Use a hygrometer to ensure there is at least 30% relative humidity. Use sorbitol or sucrose in the small liposome buffer as an additional precaution to prevent total dehydration.</li>
	<li>Prepare a saturated salt solution of appropriate relative humidity. Target humidity depends on the osmolarity of the vesicle preparation. For preparations with physiological osmolarity, use 30-45% RH, while low osmolarity vesicle preparations can be adequately dehydrated in equilibrium with 75-90% RH (See also Table 1).</li>
	<li>Put the saturated salt solution and excess salt in a tightly sealable container with an interior shelf. Commercial food containers for yogurt and granola work very well. Replace the shelf after filling, and make sure fluid is 5-10 mm below the shelf.</li>
	<li>Clean ITO coated slides as in Section 1. Use the multimeter to determine which side is conductive, and label the non-conductive side with the sample name.</li>
	<li>Lightly grease both sides of the silicone (USP Grade VI) gasket(s) with one or multiple holes.</li>
	<li>Place the slides conductive side up on the bench, and apply the silicone gasket(s) to each slide to which lipid will be applied, making sure there is at least 5 mm of exposed slide at one end to connect to the electroformation apparatus. Smooth the gasket to ensure a good seal.</li>
	<li>Dilute the vesicle preparation in a low-salt isosmotic buffer to ~1-2 mg/ml. We find that higher concentrations produce poor results.</li>
	<li>Apply lipid to the slide in 1-10 &mu;l drops. Smaller drops generally produce better results.</li>
	<li>Place the slide on the interior shelf above the saturated salt solution and seal the container tightly. Leave at RT for 3 hr to O/N. The container can be placed at 4 &deg;C, but note that for some salts, the RH depends strongly on temperature, and that cold will increase equilibration time.</li>
	<li>The resulting film may have a slight rainbow sheen to it, but in any case should appear nearly desiccated. Highly osmotic starting solutions may be impossible to dry completely to a lipid film; this will not adversely affect the results.</li>
</ol>
4. Electroformation of Giant Liposomes
<ol>
	<li>Lipid films, especially those formed from dehydrated liposomes, are easily dislodged. Carefully hydrate each well by placing a 27 G syringe needle at the edge of the gasket and slowly applying buffer. Buffers can include water, &le;200 mM sucrose, &le;1 M sorbitol and &le;5 mM HEPES. More than 10 mM salt solution may require alternate electroformation voltage protocols. Overfill each gasket well by ~10%.</li>
	<li>Note: once the lipids are hydrated, proceed quickly through the remaining steps, since lipid films begin to delaminate immediately. Yield and size are maximized if electroformation begins promptly.</li>
	<li>In one smooth motion, apply a second ITO slide, conductive face in, to the top of the gasket. Make sure to have at least 5 mm of overhang outside the gasket area and opposite the first overhang. Press gently to ensure a good seal.</li>
	<li>Clean the two overhangs using ethanol and verify that sides facing the gasket are conductive with your multimeter.</li>
	<li>The chamber can be further secured with parafilm or light tension spring clips if desired.</li>
	<li>Connect the electroformation chamber to a voltage source by securing aluminum or copper bars, "EMI gasket foam", or conductive-adhesive tape to the conductive surfaces of the two slides. We use EMI gasket foam. Plans for a jig and clamp are shown in Figure 5.</li>
	<li>Verify that there is no electrical short between the contacts, and that the contacts connect properly to the ITO surfaces, using the multimeter.</li>
	<li>Heat the chamber 10 &deg;C above the highest melting temperature of any of the lipids present; e.g. for DPPC, heat to 52 &deg;C minimum, 10 &deg;C above the chain melting temperature of 42 &deg;C.</li>
	<li>For low-salt buffers, apply a 10 Hz sine wave, ~0.7 Vrms for each millimeter gap between the two ITO coated surfaces, for 60-90 min. For high salt buffers, other voltage protocols should be used (see references).</li>
</ol>
5. Imaging and Troubleshooting
<ol>
	<li>Image the liposome using an inverted microscope equipped with a filter cube for Rhodamine or Texas Red dyes. The liposome should be spherical, predominantly unilamellar by eye, and free of defects such as "strings" hanging off the liposome.</li>
	<li>If the liposomes are too small, use less lipid.</li>
	<li>If there are many defects or few liposomes, this may be due to gel-phase lipids. Consider raising the electroformation temperature.</li>
	<li>If few or no giant liposomes formed at all from dehydrated small liposomes, reduce the osmotic strength of the liposome buffer, or increase the osmotic strength of the electroformation buffer. An inrush of buffer into the concentrated interstitial buffer solution can cause delamination of the deposited lipid film.</li>
	<li>If vesicle yield, quality, or size is poor, and you included salts or pH buffers in the electroformation or liposome buffers, consider alternative electroformation voltage protocols. For instance, Pott, et al.11, recommend a three step protocol using a 500 Hz sinewave, raising the voltage from 50-1,300 Vpp/m over 30 min, holding for 90 min, then decreasing the frequency to 50 Hz over 30-60 min. Platinum or titanium electrodes may be needed in this case, but this will not substantially alter the protocol.</li>
</ol>

<ol>
	<li>Dimova, R., Aranda, S., Bezlyepkina, N., Nikolov, V., Riske, K. A., Lipowsky, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+practical+guide+to+giant+vesicles.+Probing+the+membrane+nanoregime+via+optical+microscopy.">A practical guide to giant vesicles. Probing the membrane nanoregime via optical microscopy.</a> Journal of Physics-Condensed Matter. 18 (28), S1151-S1176 (2006).</li><li>Luisi, P. L., Walde, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Giant Vesicles. , (2000).</li><li>Riquelme, G., Lopez, E., Garcia-Segura, L. M., Ferragut, J. A., Gonzalez-Ros, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Giant+liposomes:+a+model+system+in+which+to+obtain+patch-clamp+recordings+of+ionic+channels.">Giant liposomes: a model system in which to obtain patch-clamp recordings of ionic channels.</a> Biochemistry. 29 (51), 11215-11222 (1990).</li><li>Walde, P., Cosentino, K., Engel, H., Stano, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Giant+vesicles:+preparations+and+applications.">Giant vesicles: preparations and applications.</a> Chembiochem. 11 (7), 848-865 (2010).</li><li>Aimon, S., Manzi, J., Schmidt, D., Poveda Larrosa, J. A., Bassereau, P., Toombes, G. E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+reconstitution+of+a+voltage-gated+potassium+channel+in+giant+unilamellar+vesicles.">Functional reconstitution of a voltage-gated potassium channel in giant unilamellar vesicles.</a> PLoS. One. 6 (10), e25529 (2011).</li><li>Girard, P., Pecreaux, J., Lenoir, G., Falson, P., Rigaud, J. L., Bassereau, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+method+for+the+reconstitution+of+membrane+proteins+into+giant+unilamellar+vesicles.">A new method for the reconstitution of membrane proteins into giant unilamellar vesicles.</a> Biophys. J. 87 (1), 419-429 (2004).</li><li>Manley, S., Gordon, V. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Making+giant+unilamellar+vesicles+via+hydration+of+a+lipid+film.">Making giant unilamellar vesicles via hydration of a lipid film.</a> Curr. Protoc. Cell. Biol. 24, 1-13 (2008).</li><li>Rodriguez, N., Pincet, F., Cribier, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Giant+vesicles+formed+by+gentle+hydration+and+electroformation:+a+comparison+by+fluorescence+microscopy.">Giant vesicles formed by gentle hydration and electroformation: a comparison by fluorescence microscopy.</a> Colloids. Surf. B. Biointerfaces. 42 (2), 125-130 (2005).</li><li>Angelova, M. I., Soleau, S., Meleard, P., Faucon, J. F., Bothorel, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+giant+vesicles+by+external+AC+electric+fields.+Kinetics+and+applications.">Preparation of giant vesicles by external AC electric fields. Kinetics and applications.</a> Progressin Colloid &amp; Polymer Science. 89, 127-131 (1992).</li><li>Politano, T. J., Froude, V. E., Jing, B., Zhu, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=AC-electric+field+dependent+electroformation+of+giant+lipid+vesicles.">AC-electric field dependent electroformation of giant lipid vesicles.</a> Colloids. Surf. B. Biointerfaces. 79 (1), 75-82 (2010).</li><li>Pott, T., Bouvrais, H., M&#233;l&#233;ard, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Giant+unilamellar+vesicle+formation+under+physiologically+relevant+conditions.">Giant unilamellar vesicle formation under physiologically relevant conditions.</a> Chem. Phys. Lip. 154 (2), 115-119 (2008).</li><li>Sens, P., Isambert, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Undulation+Instability+of+Lipid+Membranes+under+an+Electric+Field.">Undulation Instability of Lipid Membranes under an Electric Field.</a> Phys. Rev. Lett. 88 (12), (2002).</li><li>Green, N. G., Ramos, A., Gonz&#225;lez, A., Morgan, H., Castellanos, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluid+flow+induced+by+nonuniform+ac+electric+fields+in+electrolytes+on+microelectrodes.+I.+Experimental+measurements.">Fluid flow induced by nonuniform ac electric fields in electrolytes on microelectrodes. I. Experimental measurements.</a> Phys. Rev. E. 61 (4), 4011-4018 (2000).</li><li>Baykal-Caglar, E., Hassan-Zadeh, E., Saremi, B., Huang, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+giant+unilamellar+vesicles+from+damp+lipid+film+for+better+lipid+compositional+uniformity.">Preparation of giant unilamellar vesicles from damp lipid film for better lipid compositional uniformity.</a> Biochim. Biophys. Acta. 1818 (11), 2598-2604 (2012).</li><li>Bakalyar, S. R., Bradley, M. P. T., Honganen, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+dissolved+gases+in+high+-performance+liquid+chromatography.">The role of dissolved gases in high -performance liquid chromatography.</a> J. Chromatogr. A. 158, 277-293 (1978).</li><li>Brown, J. N., Hewins, M., Van Der Linden, J. H. M., Lynch, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Solvent+degassing+and+other+factors+affecting+liquid+chromatographic+detector+stability.">Solvent degassing and other factors affecting liquid chromatographic detector stability.</a> J. Chromatogr. A. 204, 115-122 (1981).</li><li>Dolan, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mobile+Phase+Degassing-Why,+When,+and+How.">Mobile Phase Degassing-Why, When, and How.</a> LC-GC. 17 (10), 909-912 (1999).</li><li>Cheng, H., Jiang, X., Han, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Alterations+in+lipid+homeostasis+of+mouse+dorsal+root+ganglia+induced+by+apolipoprotein+E+deficiency:+a+shotgun+lipidomics+study.">Alterations in lipid homeostasis of mouse dorsal root ganglia induced by apolipoprotein E deficiency: a shotgun lipidomics study.</a> J. Neurochem. 101 (1), 57-76 (2007).</li><li>Greenspan, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Humidity+fixed+points+of+binary+saturated+aqueous+solutions.">Humidity fixed points of binary saturated aqueous solutions.</a> J. Res. Natl. Bur. Stand. 81 (1), 89-96 (1977).</li><li>Rockland, L. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Saturated+Salt+Solutions+for+Static+Control+of+Relative+Humidity+between+5+&#176;+and+40+&#176;C.">Saturated Salt Solutions for Static Control of Relative Humidity between 5 &#176; and 40 &#176;C.</a> Anal. Chem. 32 (10), 1375-1376 (1960).</li><li>Washburn, E. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> International Critical Tables of Numerical Data, Physics, Chemistry and Technology (1st Electronic Edition). , 216-249 (2003).</li><li>Estes, D. J., Mayer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Giant+liposomes+in+physiological+buffer+using+electroformation+in+a+flow+chamber.">Giant liposomes in physiological buffer using electroformation in a flow chamber.</a> Biochim. Biophys. Acta. 1712 (2), 152-160 (2005).</li><li>Ayuyan, A. G., Cohen, F. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipid+Peroxides+Promote+Large+Rafts:+Effects+of+Excitation+of+Probes+in+Fluorescence+Microscopy+and+Electrochemical+Reactions+during+Vesicle+Formation.">Lipid Peroxides Promote Large Rafts: Effects of Excitation of Probes in Fluorescence Microscopy and Electrochemical Reactions during Vesicle Formation.</a> Biophys. J. 91 (6), 2172-2183 (2006).</li><li>Morales-Penningston, N. F., Wu, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GUV+preparation+and+imaging:+minimizing+artifacts.">GUV preparation and imaging: minimizing artifacts.</a> Biochim. Biophys. Acta. 1798 (7), 1324-1332 (2010).</li><li>Grit, M., de Smidt, J. H., Struijke, A., Crommelin, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hydrolysis+of+phosphatidylcholine+in+aqueous+liposome+dispersions.">Hydrolysis of phosphatidylcholine in aqueous liposome dispersions.</a> Int. J. Pharm. 50 (1), 1-6 (1989).</li><li>Zhou, Y., Berry, C. K., Storer, P. A., Raphael, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peroxidation+of+polyunsaturated+phosphatidyl-choline+lipids+during+electroformation.">Peroxidation of polyunsaturated phosphatidyl-choline lipids during electroformation.</a> Biomaterials. 28 (6), 1298-1306 (2007).</li><li>Farkas, E. R., Webb, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiphoton+polarization+imaging+of+steady-state+molecular+order+in+ternary+lipid+vesicles+for+the+purpose+of+lipid+phase+assignment.">Multiphoton polarization imaging of steady-state molecular order in ternary lipid vesicles for the purpose of lipid phase assignment.</a> J. Phys. Chem. B. 114 (47), 15512-15522 (2010).</li><li>Hauser, H. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+ultrasonic+irradiation+on+the+chemical+structure+of+egg+lecithin.">The effect of ultrasonic irradiation on the chemical structure of egg lecithin.</a> Biochem. Biophys. Res. Commun. 45 (4), 1049-1055 (1971).</li><li>Hauser, H., Cevc, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phospholipid+vesicles.">Phospholipid vesicles.</a> Phospholipids Handbook. , (1993).</li><li>Veatch, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electro-formation+and+fluorescence+microscopy+of+giant+vesicles+with+coexisting+liquid+phases.">Electro-formation and fluorescence microscopy of giant vesicles with coexisting liquid phases.</a> Meth. Mol. Biol. 398, 59-72 (2007).</li><li>Kim, R. S., LaBella, F. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+analytical+methods+for+monitoring+autoxidation+profiles+of+authentic+lipids.">Comparison of analytical methods for monitoring autoxidation profiles of authentic lipids.</a> J. Lipid. Res. 28 (9), 1110-1117 (1987).</li><li>Veatch, S. L., Leung, S. S. W., Hancock, R. E. W., Thewalt, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+probes+alter+miscibility+phase+boundaries+in+ternary+vesicles.">Fluorescent probes alter miscibility phase boundaries in ternary vesicles.</a> J. Phys. Chem. B. 111 (3), 502-504 (2007).</li><li>Juhasz, J., Davis, J. H., Sharom, F. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+probe+partitioning+in+GUVs+of+binary+phospholipid+mixtures:+implications+for+interpreting+phase+behavior.">Fluorescent probe partitioning in GUVs of binary phospholipid mixtures: implications for interpreting phase behavior.</a> Biochim. Biophys. Acta. 1818 (1), 19-26 (2012).</li><li>Herold, C., Chwastek, G., Schwille, P., Petrov, E. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+electroformation+of+supergiant+unilamellar+vesicles+containing+cationic+lipids+on+ITO-coated+electrodes.">Efficient electroformation of supergiant unilamellar vesicles containing cationic lipids on ITO-coated electrodes.</a> Langmuir. 28 (13), 5518-5521 (2012).</li></ol>

The authors declare that they have no competing financial interests.

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....

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, &asymp;1-100 &mu;m 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).

<table><tbody><tr><td>Digital Multimeter</td><td>Agilent Technologies, &lt;a href=&quot;http://www.agilent.com&quot; target=&quot;_blank&quot;&gt;www.agilent.com&lt;/a&gt;</td><td>U1232A or similar</td><td>Any multimeter will do, but avoid old style analog ohmmeters which apply much more current to the resistance under test.</td></tr><tr><td></td><td>Fluke</td><td>117 or 177</td><td>Any multimeter will do, but avoid old style analog ohmmeters which apply much more current to the resistance under test.</td></tr><tr><td>Function Generator</td><td>Agilent Technologies, &lt;a href=&quot;http://www.agilent.com&quot; target=&quot;_blank&quot;&gt;www.agilent.com&lt;/a&gt;</td><td>33210A or similar</td><td>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 &amp;#937; load</td></tr><tr><td>ITO coated glass slides</td><td>Delta Technologies, Loveland, CO &lt;a href=&quot;http://www.delta-technologies.com&quot; target=&quot;_blank&quot;&gt;www.delta-technologies.com&lt;/a&gt;</td><td>CB-90IN-S107 or similar</td><td>Break these in half to make two slides, 25 mm x 37 mm</td></tr><tr><td>Temperature controller</td><td>Omega Engineering Stamford, CT &lt;a href=&quot;http://www.omega.com&quot; target=&quot;_blank&quot;&gt;www.omega.com&lt;/a&gt;</td><td>CNi3233 or similar</td><td></td></tr><tr><td>Hygrometer</td><td>Extech, Nashua, NH, &lt;a href=&quot;http://www.extech.com&quot; target=&quot;_blank&quot;&gt;www.extech.com&lt;/a&gt;</td><td>445815</td><td></td></tr><tr><td>Silicone rubber sheet</td><td>McMaster-Carr Elmhurst, IL &lt;a href=&quot;http://www.mcmaster.com&quot; target=&quot;_blank&quot;&gt;www.mcmaster.com&lt;/a&gt;</td><td>&lt;a href=&quot;http://www.mcmaster.com/#87315K64&quot; target=&quot;_blank&quot;&gt;87315K64&lt;/a&gt;</td><td>Use USP Grade VI silicone for its high purity</td></tr><tr><td>EMI gasket</td><td>Laird Technologies &lt;a href=&quot;http://www.lairdtech.com&quot; target=&quot;_blank&quot;&gt;www.lairdtech.com&lt;/a&gt;</td><td>4202-PA-51H-01800 or similar</td><td>Distributed by Mouser &lt;a href=&quot;http://www.mouser.com&quot; target=&quot;_blank&quot;&gt;www.mouser.com&lt;/a&gt;</td></tr><tr><td>TxR-DHPE</td><td>Life Technologies, Carlsbad, CA &lt;a href=&quot;http://www.lifetechnologies.com&quot; target=&quot;_blank&quot;&gt;www.lifetechnologies.com&lt;/a&gt;</td><td>T1395MP</td><td>Other fluorescently labeled lipids are available, but TxR-DHPE is one of the brightest and most photostable.</td></tr><tr><td>POPC</td><td>Avanti Polar Lipids, Alabaster, AL &lt;a href=&quot;http://www.avantilipids.com&quot; target=&quot;_blank&quot;&gt;www.avantilipids.com&lt;/a&gt;</td><td>850457P or 850457C</td><td>Lipids can be ordered as powders (P) or in chloroform (C)</td></tr><tr><td>POPS</td><td>Avanti Polar Lipids</td><td>840034P/C</td><td></td></tr><tr><td>Cholesterol</td><td>Sigma-Aldrich</td><td>C8667</td><td></td></tr></tbody></table>

giant liposome preparation for imaging and patch-clamp electrophysiology

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 &gt;10 &mu;m 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.

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 electrofo...

Watch this Scientific Journal Video about Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology at JoVE.com

Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

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.

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify ...

giant-liposome-preparation-for-imaging-patch-clamp

Research

JoVE Journal

Biology

22.1K Views.  University of Washington. 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.

Video: Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

Site Directed Spin Labeling and EPR Spectroscopic Studies of Pentameric Ligand-Gated Ion Channels

Ion channel gating is a stimulus-driven orchestration of protein motions that leads to transitions between closed, open, and desensitized states. Fundamental to these transitions is the intrinsic flexibility of the protein, which is critically modulated by membrane lipid-composition. To better understand the structural basis of channel function, it is necessary to study protein dynamics in a physiological membrane environment. Electron Paramagnetic Resonance (EPR) spectroscopy is an important tool to characterize conformational transitions between functional states. In comparison to NMR and X-ray crystallography, the information obtained from EPR is intrinsically of lower resolution. However, unlike in other techniques, in EPR there is no upper-limit to the molecular weight of the protein, the sample requirements are significantly lower, and more importantly the protein is not constrained by the crystal lattice forces. Therefore, EPR is uniquely suited for studying large protein complexes and proteins in reconstituted systems. In this article, we will discuss general protocols for site-directed spin labeling and membrane reconstitution using a prokaryotic proton-gated pentameric Ligand-Gated Ion Channel (pLGIC) from Gloeobacter violaceus (GLIC) as an example. A combination of steady-state Continuous Wave (CW) and Pulsed (Double Electron Electron Resonance-DEER) EPR approaches will be described that will enable a complete quantitative characterization of channel dynamics.

This article describes methods for site-directed spin labeling and reconstitution of pentameric ligand-gated channels for Electron Paramagnetic Resonance studies. This protocol can be adapted for any membrane protein. The reconstitution method described here can also be used for patch-clamp measurements of macroscopic and single-channel currents in a defined lipid system.

Ion channel gating is a stimulus-driven orchestration of protein motions that leads to transitions between closed, open, and desensitized states. ...

Chemistry

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous number of imaging techniques currently available and the progress in instrumentation, there is still a need for highly sensitive probes that are suitable for in vivo imaging. One typical problem of available preclinical fluorescent probes is their rapid clearance in vivo, which reduces their imaging sensitivity. To circumvent rapid clearance, increase number of dye molecules at the target site, and thereby reduce background autofluorescence, encapsulation of the near-infrared fluorescent dye, DY-676-COOH in liposomes and verification of its potential for in vivo imaging of inflammation was done. DY-676 is known for its ability to self-quench at high concentrations. We first determined the concentration suitable for self-quenching, and then encapsulated this quenching concentration into the aqueous interior of PEGylated liposomes. To substantiate the quenching and activation potential of the liposomes we use a harsh freezing method which leads to damage of liposomal membranes without affecting the encapsulated dye. The liposomes characterized by a high level of fluorescence quenching were termed Lip-Q. We show by experiments with different cell lines that uptake of Lip-Q is predominantly by phagocytosis which in turn enabled the characterization of its potential as a tool for in vivo imaging of inflammation in mice models. Furthermore, we use a zymosan-induced edema model in mice to substantiate the potential of Lip-Q in optical imaging of inflammation in vivo. Considering possible uptake due to inflammation-induced enhanced permeability and retention (EPR) effect, an always-on liposome formulation with low, non-quenched concentration of DY-676-COOH (termed Lip-dQ) and the free DY-676-COOH were compared with Lip-Q in animal trials.

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

The use of fluorophores for in vivo imaging can be greatly limited by opsonization, rapid clearance, low detection sensitivity and cytotoxic effects on the host. Encapsulation of fluorophores in liposomes by film hydration and extrusion leads to fluorescence quenching and protection which enables in vivo imaging with high detection sensitivity.

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous ...

Bioengineering

Membrane Remodeling of Giant Vesicles in Response to Localized Calcium Ion Gradients

In a wide variety of fundamental cell processes, such as membrane trafficking and apoptosis, cell membrane shape transitions occur concurrently with local variations in calcium ion concentration. The main molecular components involved in these processes have been identified; however, the specific interplay between calcium ion gradients and the lipids within the cell membrane is far less known, mainly due to the complex nature of biological cells and the difficultly of observation schemes. To bridge this gap, a synthetic approach is successfully implemented to reveal the localized effect of calcium ions on cell membrane mimics. Establishing a mimic to resemble the conditions within a cell is a severalfold problem. First, an adequate biomimetic model with appropriate dimensions and membrane composition is required to capture the physical properties of cells. Second, a micromanipulation setup is needed to deliver a small amount of calcium ions to a particular membrane location. Finally, an observation scheme is required to detect and record the response of the lipid membrane to the external stimulation. This article offers a detailed biomimetic approach for studying the calcium ion-membrane interaction, where a lipid vesicle system, consisting of a giant unilamellar vesicle (GUV) connected to a multilamellar vesicle (MLV), is exposed to a localized calcium gradient formed using a microinjection system. The dynamics of the ionic influence on the membrane were observed using fluorescence microscopy and recorded at video frame rates. As a result of the membrane stimulation, highly curved membrane tubular protrusions (MTPs) formed inside the GUV, oriented away from the membrane. The described approach induces the remodeling of the lipid membrane and MTP production in an entirely contactless and controlled manner. This approach introduces a means to address the details of calcium ion-membrane interactions, providing new avenues to study the mechanisms of cell membrane reshaping.

We present a technique for contactless micromanipulation of vesicles, using localized calcium ion gradients. Microinjection of a calcium ion solution, in the vicinity of a giant lipid vesicle, is utilized to remodel the lipid membrane, resulting in the production of membrane tubular protrusions.

In a wide variety of fundamental cell processes, such as membrane trafficking and apoptosis, cell membrane shape transitions occur concurrently with local ...

Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective imaging of fluorescent molecules that are closest to a high refractive index substance such as glass1. In this article, we apply this technique to image exocytosis of synaptic vesicles in retinal bipolar cells isolated from the goldfish retina. These neurons are very suitable for this kind of study due to their large axon terminals. By simultaneously patch clamping the bipolar cells, it is possible to investigate the relationship between pre-synaptic voltage and synaptic release2,3. Synaptic vesicles inside the bipolar cell terminals are loaded with a fluorescent dye (FM 1-43&reg;) by co-puffing the dye and a ringer solution containing a high K+ concentration onto the synaptic terminals. This depolarizes the cells and stimulates endocytosis and consequent dye uptake into the glutamatergic vesicles. After washing the excess dye away for around 30 minutes, cells are ready for being patch clamped and imaged simultaneously with a 488 nm laser. The patch pipette solution contains a rhodamine-based peptide that binds selectively to the synaptic ribbon protein RIBEYE4, thereby labeling ribbons specifically when terminals are imaged with a 561 nm laser. This allows the precise localization of active zones and the separation of synaptic from extra-synaptic events.

In this video, we demonstrate how to label and visualize single synaptic vesicle exocytosis and trafficking in goldfish retinal bipolar cells using total internal reflectance fluorescence (TIRF) microscopy.

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective ...

Super-resolution Imaging of the Natural Killer Cell Immunological Synapse on a Glass-supported Planar Lipid Bilayer

The glass-supported planar lipid bilayer system has been utilized in a variety of disciplines. One of the most useful applications of this technique has been in the study of immunological synapse formation, due to the ability of the glass-supported planar lipid bilayers to mimic the surface of a target cell while forming a horizontal interface. The recent advances in super-resolution imaging have further allowed scientists to better view the fine details of synapse structure. In this study, one of these advanced techniques, stimulated emission depletion (STED), is utilized to study the structure of natural killer (NK) cell synapses on the supported lipid bilayer. Provided herein is an easy-to-follow protocol detailing: how to prepare raw synthetic phospholipids for use in synthesizing glass-supported bilayers; how to determine how densely protein of a given concentration occupies the bilayer's attachment sites; how to construct a supported lipid bilayer containing antibodies against NK cell activating receptor CD16; and finally, how to image human NK cells on this bilayer using STED super-resolution microscopy, with a focus on distribution of perforin positive lytic granules and filamentous actin at NK synapses. Thus, combining the glass-supported planar lipid bilayer system with STED technique, we demonstrate the feasibility and application of this combined technique, as well as intracellular structures at NK immunological synapse with super-resolution.

We describe here a combination of the glass-supported lipid bilayer technique of forming immunological synapses with the super-resolution imaging technique of stimulated emission depletion (STED) microscopy. The goal of this protocol is to provide users with the instructions necessary to successfully carry out these two techniques.

The glass-supported planar lipid bilayer system has been utilized in a variety of disciplines. One of the most useful applications of this technique has ...

Immunology and Infection

Preparation, Purification, and Use of Fatty Acid-containing Liposomes

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due to the unique chemical properties of these amphiphiles. In particular, fatty acid liposomes enhance dynamic character, due to the relatively high solubility of single-chain amphiphiles. Similarly, liposomes containing free fatty acids are more sensitive to salt and divalent cations, due to the strong interactions between the carboxylic acid head groups and metal ions. Here we illustrate techniques for preparation, purification, and use of liposomes comprised in part or whole of single chain amphiphiles (e.g., oleic acids).

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due to the unique chemical properties of single chain amphiphiles. Here we describe techniques for the preparation, purification, and use of liposomes comprised in part or whole of these amphiphiles.

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due ...

SNARE-mediated Fusion of Single Proteoliposomes with Tethered Supported Bilayers in a Microfluidic Flow Cell Monitored by Polarized TIRF Microscopy

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. During neurotransmitter or hormone release via exocytosis, the fusion pore can transiently open and close repeatedly, regulating cargo release kinetics. Pore dynamics also determine the mode of vesicle recycling; irreversible resealing results in transient, &#34;kiss-and-run&#34; fusion, whereas dilation leads to full fusion. To better understand what factors govern pore dynamics, we developed an assay to monitor membrane fusion using polarized total internal reflection fluorescence (TIRF) microscopy with single molecule sensitivity and ~15 msec time resolution in a biochemically well-defined in vitro system. Fusion of fluorescently labeled small unilamellar vesicles containing v-SNARE proteins (v-SUVs) with a planar bilayer bearing t-SNAREs, supported on a soft polymer cushion (t-SBL, t-supported bilayer), is monitored. The assay uses microfluidic flow channels that ensure minimal sample consumption while supplying a constant density of SUVs. Exploiting the rapid signal enhancement upon transfer of lipid labels from the SUV to the SBL during fusion, kinetics of lipid dye transfer is monitored. The sensitivity of TIRF microscopy allows tracking single fluorescent lipid labels, from which lipid diffusivity and SUV size can be deduced for every fusion event. Lipid dye release times can be much longer than expected for unimpeded passage through permanently open pores. Using a model that assumes retardation of lipid release is due to pore flickering, a pore &#34;openness&#34;, the fraction of time the pore remains open during fusion, can be estimated. A soluble marker can be encapsulated in the SUVs for simultaneous monitoring of lipid and soluble cargo release. Such measurements indicate some pores may reseal after losing a fraction of the soluble cargo.

Here, we present a protocol to detect single, SNARE-mediated fusion events between liposomes and supported bilayers in microfluidic channels using polarized TIRFM, with single molecule sensitivity and ~15 msec time resolution. Lipid and soluble cargo release can be detected simultaneously. Liposome size, lipid diffusivity, and fusion pore properties are measured.

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. ...

Neuroscience

Lipid Bilayer Experiments with Contact Bubble Bilayers for Patch-Clampers

Lipid bilayers provide a unique experimental platform for functional studies of ion channels, allowing the examination of channel-membrane interactions under various membrane lipid compositions. Among them, the droplet interface bilayer has gained popularity; however, the large membrane size hinders the recording of low electrical background noise. We have established a contact bubble bilayer (CBB) method that combines the benefits of planar lipid bilayer and patch-clamp methods, such as the ability to vary the lipid composition and to manipulate the bilayer mechanics, respectively. Using the setup for conventional patch-clamp experiments, CBB-based experiments can be readily performed. In brief, an electrolyte solution in a glass pipette is blown into an organic solvent phase (hexadecane), and the pipette pressure is maintained to obtain a stable bubble size. The bubble is spontaneously lined with a lipid monolayer (pure lipids or mixed lipids), which is provided from liposomes in the bubbles. Next, the two monolayer-lined bubbles (~50 &#181;m in diameter) at the tip of the glass pipettes are docked for bilayer formation. Introduction of channel-reconstituted liposomes into the bubble leads to the incorporation of channels in the bilayer, allowing for single-channel current recording with a signal-to-noise ratio comparable to that of patch-clamp recordings. CBBs with an asymmetric lipid composition are readily formed. The CBB is renewed repeatedly by blowing out the previous bubbles and forming new ones. Various chemical and physical perturbations (e.g., membrane perfusion and bilayer tension) can be imposed on the CBBs. Herein, we present the basic procedure for CBB formation.

Here, we present a protocol for the formation of lipid bilayers using a contact bubble bilayer method. A water bubble is blown into an organic solvent, whereby a monolayer is formed at the water-oil interface. Two pipettes are manipulated to dock the bubbles to form a bilayer.

Lipid bilayers provide a unique experimental platform for functional studies of ion channels, allowing the examination of channel-membrane interactions ...

Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow GUVs must be modified in order to form GUVs containing functional transmembrane proteins. This article describes two dehydration-rehydration methods &#8212; electroformation and gel-assisted swelling &#8212; to form GUVs containing the voltage-gated potassium channel, KvAP. In both methods, a solution of protein-containing small unilamellar vesicles is partially dehydrated to form a stack of membranes, which is then allowed to swell in a rehydration buffer. For the electroformation method, the film is deposited on platinum electrodes so that an AC field can be applied during film rehydration. In contrast, the gel-assisted swelling method uses an agarose gel substrate to enhance film rehydration. Both methods can produce GUVs in low (e.g., 5 mM) and physiological (e.g., 100 mM) salt concentrations. The resulting GUVs are characterized via fluorescence microscopy, and the function of reconstituted channels measured using the inside-out patch-clamp configuration. While swelling in the presence of an alternating electric field (electroformation) gives a high yield of defect-free GUVs, the gel-assisted swelling method produces a more homogeneous protein distribution and requires no special equipment.

The reconstitution of the transmembrane protein, KvAP, into giant unilamellar vesicles (GUVs) is demonstrated for two dehydration-rehydration methods &#8212; electroformation, and gel-assisted swelling. In both methods, small unilamellar vesicles containing the protein are fused together to form GUVs that can then be studied by fluorescence microscopy and patch-clamp electrophysiology.

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow ...

Patch-Clamp Techniques for Single Endolysosomal Vesicle Analysis

Endolysosomal ion channels are critical for endolysosomal ion and pH homeostasis, membrane potential regulation, and vesicle trafficking. However, electrophysiologically accessing these channels within small intracellular vesicles has been a challenge. The development of endolysosomal patch-clamp techniques has been instrumental in overcoming this barrier, allowing for the direct measurement of ion channel activity in endolysosomal membranes.
Compared to existing planar patch-clamp techniques, endolysosomal patch-clamp can simultaneously record multiple cells and easily combine with other measurement methods. Manual operation offers the advantage of visualizing targeted vesicles. It also addresses the limitation of the indispensable presence of Ca2+ on one side of the endolysosomal membrane, increasing the flexibility of experimental design. Utilizing endolysosomal patch-clamp techniques enables the direct measurement and analysis of ion channel activity within endolysosomes.
Given the close link between aberrant endolysosomal ion channel function and diseases such as neurodegenerative diseases and metabolic disorders, investigating and modulating these channels may unveil new drug targets. By restoring intracellular ion balance, we may alleviate or cure related diseases. Therefore, this technique is pivotal for discovering new drug targets and developing relevant medications.

This protocol details a method for directly measuring ion channel activity on intracellular vesicles using a manual endolysosomal patch-clamp system. We illustrate the method that involves enlarging endolysosomes and manually isolating these vesicles. This approach ensures that researchers can accurately replicate and apply the procedure.

Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

Summary

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Chapters in this video

Site Directed Spin Labeling and EPR Spectroscopic Studies of Pentameric Ligand-Gated Ion Channels

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Membrane Remodeling of Giant Vesicles in Response to Localized Calcium Ion Gradients

Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

Super-resolution Imaging of the Natural Killer Cell Immunological Synapse on a Glass-supported Planar Lipid Bilayer

Preparation, Purification, and Use of Fatty Acid-containing Liposomes

SNARE-mediated Fusion of Single Proteoliposomes with Tethered Supported Bilayers in a Microfluidic Flow Cell Monitored by Polarized TIRF Microscopy

Lipid Bilayer Experiments with Contact Bubble Bilayers for Patch-Clampers

Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies

Patch-Clamp Techniques for Single Endolysosomal Vesicle Analysis