We thank Susanne Fenz for discussing the possibility of reconstituting proteins by agarose swelling, Feng Ching Tsai for current measurements, and present and former members of the Bassereau group for support and assistance. The project was funded by the Agence Nationale de la Recherche (grant BLAN-0057-01), by the European Commission (NoE SoftComp), by the Universit&#233; Pierre et Marie Curie (grant from the FED21, Dynamique des Syst&#232;mes Complexes). M.G. was supported by an Institut Curie International PhD Fellowship, S.A. by a fellowship from the Fondation pour la Recherche M&#233;dicale, G.E.S.T. by a Marie Curie Incoming International Fellowship from the European Commission and a grant from the Universit&#233; Pierre et Marie Curie. The publication fees were covered by the Labex &#8216;CelTisPhyBio&#8217; (ANR-11-LABX0038).

1. Solution Preparation
<ol>
	<li>Prepare 5 ml of &#8216;SUV buffer&#8217; containing 5 mM KCl, 1 mM HEPES (pH 7.4) or TRIS (pH 7.5), and 2 mM trehalose. Filter the buffer with a 0.2 &#181;m syringe filter and divide into 1 ml aliquots which can be stored at -20 &#176;C. 
	NOTE: Additional details for reagents and instruments are given in the materials list.</li>
	<li>Prepare 40 ml of GUV &#8216;Growth Buffer&#8217; that will fill the GUV interior during film rehydration. For a &#8216;low salt&#8217; growth, combine 5 mM KCl, 1 mM HEPES (pH 7.4) or 1 mM TRIS (pH 7.5), and ~400 mM sucrose. For a &#8216;physiological salt&#8217; growth, combine 100 mM KCl, 5 mM HEPES (pH 7.4) or 5 mM TRIS (pH 7.5), and ~200 mM sucrose.</li>
	<li>Prepare 40 ml of &#8216;Observation Buffer&#8217; for the external solution in the experimental chamber by combining 100 mM KCl, 5 mM HEPES (pH 7.4) or 5 mM TRIS (pH 7.5), and ~200 mM glucose. 
	NOTE: These buffers are only examples. See the discussion to adapt the buffers for other experiments.</li>
	<li>Measure the osmolarities of the growth and observation buffers with an osmometer. Add granules of sucrose or glucose to match them to within 1% so that GUVs will not lyse or collapse when transferred from the growth chamber to the observation chamber. 
	NOTE: In this concentration range, adding 1 mM of sucrose (13.7 mg per 40 ml) or glucose (7.2 mg per 40 ml) increases osmolarity by ~ 1 mOsm.</li>
	<li>Filter the growth and observation buffers with a 0.2 &#181;m filter and store them at 4 &#176;C to inhibit bacterial growth.</li>
	<li>Dissolve 50 mg of beta-casein in 10 ml of 20 mM TRIS (pH 7.5) buffer to form a 5 mg/ml beta-casein solution needed to passivate the surfaces of experimental chambers so that GUVs do not stick, spread and burst. Once the beta-casein is completely dissolved (up to several hours at 4 &#176;C), filter (0.2 &#181;m) it into 0.5 ml aliquots which can be flash frozen and stored at -20 &#176;C for later use (thawed aliquots stored at 4 &#176;C can typically be used for up to a 1 week).</li>
</ol>
2. SUV Preparation
<ol>
	<li>Prepare and freeze aliquots of proteo-SUVs following the previously published detailed protocol28. Use KvAP fluorescently labeled with Alexa-488 maleimide, reconstituted into DPhPC SUVs (10 mg/ml) at a protein to lipid ratio of 1:10 (by mass). 
	NOTE: Wild-type KvAP contains one cysteine per monomer located near the intra-cellular C-terminus (amino acid 247).</li>
	<li>Fluorescent, Lipid-only SUVs: 
	NOTE: Handle chloroform under a fume hood wearing nitrile gloves and safety glasses. Avoid the use of any plastic as chloroform can dissolve them. Chloroform solutions can be stored in amber glass vials with Teflon caps and transferred using glass syringes. Take care to rinse all glassware at least 5 to 10 times with chloroform before and after pipetting lipids.
	<ol>
		<li>Prepare 100 &#181;l of 10 mg/ml DPhPC SUVs containing 0.5 mol% of the red fluorescent lipid, Texas Red-DHPE, by mixing 100 &#181;l of DPhPC solution (10 mg/ml in chloroform) with 8.2 &#181;l of Texas Red-DHPE solution (1 mg/ml in methanol) in a 1.5 ml amber glass vial.</li>
		<li>Dry the lipids down under a stream of nitrogen in a chemical hood while rotating the vial. When the film appears to be dry, place the lipids under a vacuum for 3 hr to remove any residual solvent.</li>
		<li>Add 100 &#181;l of SUV buffer to the lipids, and vortex vigorously until no lipid remains stuck to the walls of the vial and the solution is uniformly milky.</li>
		<li>Sonicate the lipid solution to form SUVs. Adjust the vial position until the ultrasound causes the most movement and flow inside the vial, and take care not to heat the solution unnecessarily. Continue sonication until the solution becomes translucent, or when possible, transparent (2&#8211;5 min for tip sonication, ~20 min for bath sonication).</li>
		<li>Aliquot SUVs (e.g., 10 &#181;l or 20 &#181;l) and freeze (-20 &#176;C) for later use. 
		NOTE: Lipids, especially unsaturated lipids, can easily breakdown. Store lipid solutions at 20 &#176;C (or 80 &#176;C) under argon and use within 6 months. Lipid breakdown products can be detected with Thin Layer Chromatography.</li>
	</ol>
	</li>
</ol>
3. GUV Growth by Electroformation
<ol>
	<li>Prepare the electroformation chamber.
	<ol>
		<li>If the chamber has not been cleaned, remove the windows, wipe off all sealant and grease, extract the wires, and rinse and scrub the chamber with a tissue using water and ethanol (&#8805;70%) alternately.</li>
		<li>Rub the wires well, submerge the wires and chamber in acetone, and sonicate for 5 min. Wipe everything with a tissue again using acetone. Put the chamber in ethanol and sonicate for 5 min.</li>
		<li>Assemble the chamber by inserting the wires through the holes, and rotate and wipe the wires to make sure they are clean. Put the chamber in distilled water, sonicate for 5 min and dry the chamber with a stream of nitrogen or air.</li>
	</ol>
	</li>
	<li>Prepare 30 &#181;l of 3 mg/ml SUV suspension in SUV buffer. To form protein-containing GUVs, combine 8 &#181;l of proteo-SUVs (DPhPC 10 mg/ml KvAP 1:10), 2 &#181;l of fluorescent SUVs (10 mg/ml DPhPC, 0.5 mol% TexasRed-DHPE) and 20 &#181;l of SUV buffer in a 1.5 ml microcentrifuge tube for a final protein to lipid (mass) ratio of 1:12.5 and 0.1 mol% TexasRed-DHPE. Mix the solution vigorously.
	<ol>
		<li>Alternatively, to practice the protocol with lipid-only SUVs, simply combine 10 &#181;l of fluorescent SUVs (10 mg/ml DPhPC and 0.5 mol% TexasRed-DHPE) with 20 &#181;l SUV buffer.</li>
	</ol>
	</li>
	<li>Deposit the SUV Solution.
	<ol>
		<li>Use a 2 &#181;l pipette or 5 &#181;l glass syringe to deposit small (&#60;0.2 &#181;l) droplets of the SUV solution on the wires. Approximately 1 &#181;l of solution is needed to form a series of drops along 1 cm of wire. Make sure the drops are small enough and spaced far enough apart that they do not touch or fuse.</li>
		<li>Let the deposited SUVs dry for ~30 min in open air. When all the drops have settled, rotate the wire so the lipid deposits are easier to observe with the microscope. 
		NOTE: If the SUVs do not dry sufficiently, they can just wash off the wires when the growth buffer is added, while drying too much can damage the protein. Because air humidity influences the rate of drying, the drying time and/or air humidity can be adjusted for optimal results30. The lipid film on the wires should be visible under a microscope.</li>
	</ol>
	</li>
	<li>Assemble the chamber.
	<ol>
		<li>Seal the Chamber Bottom: Use a syringe to apply vacuum grease to the bottom of the chamber around the three wells and press a 40 mm x 22 mm coverslip gently against it to seal the chamber bottom so that it adheres without a gap. Seal the sides of the chamber (where the wires exit) with sealing paste. Apply vacuum grease on top of the chamber outlining the three wells.</li>
		<li>Slowly add growth buffer until each well is filled to the top. Avoid any rapid movement of the solution in the wells as this can strip the lipid film off the electrodes.</li>
		<li>Close the chamber by pressing the top cover slide gently onto the grease, taking care not to dislodge the bottom coverslip. Use a tissue to remove any drops of buffer at the edges of the top cover slip. 
		NOTE: This is a good time to examine the chamber under the microscope to confirm that the lipid film has remained on the wires.</li>
	</ol>
	</li>
	<li>Connect the signal generator to the wires using two alligator clips. Set the frequency (10 Hz/500 Hz sine wave for low/high salt buffer) and use a multimeter to measure and adjust the voltage across the wires to 0.7/0.35 V root mean square (Vrms) for the low/high salt buffer. Cover the chamber with aluminum foil to protect the fluorophores from light. Leave the GUVs to grow for 2 to 3 hr for the low salt buffer, and 12 hr or O/N for the high salt buffer.</li>
	<li>Disconnect the chamber from the generator and carefully place it on an inverted microscope to evaluate GUV growth. Use slow, steady movements or fluid flow in the wells may prematurely detach GUVs from the wires. 
	NOTE: GUVs on the wire edges are usually visible in phase contrast (40X long working distance objective), while GUVs anywhere on the lower half of the wires can be seen with epifluorescence. If no GUVs are visible, try rotating the wires to look at the upper surface. GUVs can be stored at 4 &#176;C in a growth chamber for several days.</li>
</ol>
4. GUV Growth by Gel-assisted Swelling
<ol>
	<li>Prepare 10 ml of a 1% agarose solution by mixing 100 mg of agarose with 10 ml of pure water. Heat it until it boils by placing it in a microwave at 480 W for ~20 sec. Stir to make sure the agarose is completely dissolved. 
	NOTE: The solution can be stored at 4 &#176;C and reheated when needed.</li>
	<li>Plasma-clean (air plasma) a cover-slide for 1 min so that the agarose solution will spread nicely on it. Use the cover-slides within the next 15 min as the effect of plasma cleaning wears off quickly.</li>
	<li>Apply 200 &#181;l of warm agarose solution to each 22 x 22 mm2 slide so the solution wets the entire surface. Tilt the slide vertically and touch the lower edge to a tissue to remove excess liquid and leave just a thin smooth layer of agarose on the slide.</li>
	<li>Place the slide on a hot plate or oven at 60 &#176;C and leave it to dry for at least 30 min. The agarose film is hardly visible by eye. After the slides cool to RT, use them immediately, or store them for up to one week in a closed container at 4 &#176;C.</li>
	<li>Place the agarose-coated coverslip in a standard 3.5 cm Petri dish.</li>
	<li>Prepare the SUV solution as in Section 3.2 and apply ~15 &#181;l of the SUV solution (3 mg/ml lipid) in ~30 very small drops gently onto the agarose surface. Take care not to distort the agarose layer too much.</li>
	<li>Place the slide under a gentle stream of nitrogen for about 10&#8211;15 min and follow the evaporation of the buffer by eye as the droplets dry.</li>
	<li>As soon as the SUVs have dried, add growth buffer to cover the slide surface. For a small 3.5 cm Petri dish use ~1 ml of buffer.</li>
	<li>Allow the swelling to proceed for ~30 min, and then examine the growth of GUVs in the chamber using an inverted microscope with phase-contrast or Differential Interference Contrast (DIC). 
	NOTE: Epifluorescence observation is difficult due to the strong background of fluorophores in the gel and the auto-fluorescence of the agarose.</li>
</ol>
5. Harvesting and Observing GUVs
<ol>
	<li>Passivate the observation chamber (e.g., small Petri dish or glass coverslip) so that GUVs do not stick, spread and burst on the chamber bottom. Cover the chamber bottom with beta-casein solution, incubate for 5 min, rinse out the casein solution with pure water, dry with a stream of air or nitrogen, and finally add observation buffer (e.g., ~5 mm depth for a small Petri dish).</li>
	<li>Harvest the GUVs. Cut the end of a 100 &#181;l pipette tips so the opening is larger (~2 mm diameter), and aspire slowly as the shear stress of pipetting can easily destroy GUVs.
	<ol>
		<li>For electro-formed GUVs, open the growth chamber by gently removing the top coverslip. Place the pipette tip directly above each wire and aspirate ~50 &#181;l while moving the pipette tip along the wire to detach the GUVs. 
		NOTE: It may help to rotate the wire to collect GUVs on the &#8220;other side&#8221; of the wire.</li>
		<li>For &#8220;gel-assisted swelling&#8221; GUVs, first tap the side of the petri dish a few times to help the GUVs detach from the coverslip surface. Position the pipette tip just above the coverslip and aspirate 50 &#181;l while pulling the tip back over the surface. Directly transfer harvested GUVs to an observation chamber, or store in a 1.5 ml microcentrifuge tube at 4 &#176;C for up to 1 week.</li>
	</ol>
	</li>
	<li>Place the observation chamber on an inverted microscope, add the GUVs to the observation chamber, and wait a few minutes for the GUVs to settle at the chamber bottom.</li>
	<li>Survey the chamber with phase contrast or DIC to quickly locate smooth, spherical (&#8216;defect free&#8217;) &#8220;GUV candidates&#8221;. Examine each &#8220;GUV candidate&#8221; in epifluorescence to exclude any containing smaller liposomes nested inside. Finally, check that the lipid fluorescence intensity is uniform and compatible with a single membrane (i.e., unilamellar). 
	NOTE: In some bilamellar (or multi-lamellar) vesicles, the membranes are too close together to be resolved so they appear unilamellar in phase contrast or DIC images. However, these objects can be distinguished from actual unilamellar GUVs by their lipid fluorescence, which is two times (or more) brighter.</li>
</ol>
6. Patch-clamping GUVs
<ol>
	<li>Make patch pipettes with a 1&#8211;2 &#181;m tip diameter from standard borosilicate capillary glass using the program recommended for the pipette puller. 
	NOTE: Special treatments such as fire polishing are not necessary, and pipettes can be used for several days after they have been pulled if they are kept in a closed box.</li>
	<li>Passivate the chamber by incubating with a beta-casein solution (5 mg/ml) to ensure that GUVs do not adhere, spread and rupture on chamber surfaces. Rinse the casein off after 5 min.</li>
	<li>Insert the ground electrode, fill the chamber with observation buffer, transfer 10 &#181;l of the GUV suspension as described in step 5.2 and 5.3, and wait a few minutes for the GUVs to settle at the bottom.</li>
	<li>Fill a fresh patch pipette with solution (observation buffer or another iso-osmotic solution) and mount it on the patch-clamp amplifier headstage.</li>
	<li>Search through the chamber to locate a &#8220;defect-free&#8221; GUV as described in section 5.4, and check that it contains fluorescent protein.</li>
	<li>Apply a constant positive pressure (&#62; 100 Pa, or roughly 1 cm H2O in a manometer) to keep the patch pipette interior clean, and insert the patch pipette into the chamber. Bring the patch pipette into the field of view, apply test pulses to measure/compensate the pipette voltage offset and resistance, and examine the pipette under fluorescent illumination to confirm that the tip is clean.</li>
	<li>Bring the patch pipette towards the GUV, and if necessary, simultaneously reduce the positive pressure so the outward flow from the patch pipette does not make the GUV &#8220;run away&#8221;. When the patch pipette is close to the GUV, apply a negative pressure (up to 5 cm H2O) to pull the GUV against the patch pipette. Monitor the resistance as the &#8220;tongue&#8221; of GUV membrane enters the patch pipette and the gigaseal forms.</li>
	<li>If a gigaseal did not form, remove the patch pipette from the chamber and return to step 6.4. If the membrane patch formed a gigaseal, but the GUV remains attached to the pipette, excise the patch by pulling away from the GUV, bursting the GUV against the chamber bottom, or briefly moving the pipette out of solution.</li>
	<li>When the inside-out membrane patch has been excised from the GUV and the gigaseal is stable, switch off the test pulses and apply a voltage protocol such as the one shown in Figure 13. 
	NOTE: Figure 13 follows the standard electrophysiological convention for an inside-out patch in which current flowing into the patch-electrode is &#8220;positive&#8221;, and V = Vbath&#8211; Vpipette. Holding the patch at a negative potential (e.g., V = -100 mV) for ~30 sec places KvAP in the resting state, while steps (100 msec to 5 sec) to more positive potentials (e.g., V = 100 mV) can then drive it into conducting (i.e., open) active states.</li>
	<li>After measurements on a membrane patch are finished, break the patch with a zap or pressure pulse and check that the voltage offset of the patch electrode has not drifted. Remove the patch pipette from the chamber, and return to step 6.4.</li>
</ol>

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P., Schwille, P. <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+Lipids+in+VDAC+Oligomerization.">The Role of Lipids in VDAC Oligomerization.</a> Biophysical Journal. 102 (3), 523-531 (2012).</li><li>Ruta, V., Jiang, Y., Lee, A., Chen, J., MacKinnon, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+analysis+of+an+archaebacterial+voltage-dependent+K++channel.">Functional analysis of an archaebacterial voltage-dependent K+ channel.</a> Nature. 422 (6928), 180-185 (2003).</li><li>Jiang, Y., 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=X-ray+structure+of+a+voltage-dependent+K++channel.">X-ray structure of a voltage-dependent K+ channel.</a> Nature. 423 (6935), 33-41 (2003).</li><li>Aimon, S., 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=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>Lee, S., Zheng, H., Shi, L., Jiang, Q. -. X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reconstitution+of+a+Kv+Channel+into+Lipid+Membranes+for+Structural+and+Functional+Studies.">Reconstitution of a Kv Channel into Lipid Membranes for Structural and Functional Studies.</a> Journal of Visualized Experiments. (77), (2013).</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> Biochimica et Biophysica Acta (BBA) - Biomembranes. 1818 (11), 2598-2604 (2012).</li><li>Suchyna, T. M., Markin, V. S., Sachs, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biophysics+and+Structure+of+the+Patch+and+the+Gigaseal.">Biophysics and Structure of the Patch and the Gigaseal.</a> Biophysical Journal. 97 (3), 738-747 (2009).</li><li>Hille, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Ion Channels of Excitable Membranes. , (2001).</li><li>Finol-Urdaneta, R. K., McArthur, J. R., Juranka, P. F., French, R. J., Morris, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulation+of+KvAP+unitary+conductance+and+gating+by+1-alkanols+and+other+surface+active+agents.">Modulation of KvAP unitary conductance and gating by 1-alkanols and other surface active agents.</a> Biophysical journal. 98 (5), 762-772 (2010).</li><li>Schmidt, D., MacKinnon, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Voltage-dependent+K++channel+gating+and+voltage+sensor+toxin+sensitivity+depend+on+the+mechanical+state+of+the+lipid+membrane.">Voltage-dependent K+ channel gating and voltage sensor toxin sensitivity depend on the mechanical state of the lipid membrane.</a> Proceedings of the National Academy of Sciences. 105 (49), 19276-19281 (2008).</li><li>Quemeneur, F., 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=Shape+matters+in+protein+mobility+within+membranes.">Shape matters in protein mobility within membranes.</a> Proceedings of the National Academy of Sciences of the United States of America. , (2014).</li><li>Parc, A. L., Leonil, J., Chanat, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=&#945;S1-casein,+which+is+essential+for+efficient+ER-to-Golgi+casein+transport,+is+also+present+in+a+tightly+membrane-associated+form.">&#945;S1-casein, which is essential for efficient ER-to-Golgi casein transport, is also present in a tightly membrane-associated form.</a> BMC Cell Biology. 11 (1), 1-15 (2010).</li><li>Garten, M., Toombes, G. E. S., Aimon, S., 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=Studying+Voltage+Dependent+Proteins+with+Giant+Unilamellar+Vesicles+in+a+&#8220;Whole+Cell&#8221;+Configuration.">Studying Voltage Dependent Proteins with Giant Unilamellar Vesicles in a &#8220;Whole Cell&#8221; Configuration.</a> Biophysical Journal. 104 (2), 466a (2013).</li><li>Crowe, L. M., Reid, D. S., Crowe, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Is+trehalose+special+for+preserving+dry+biomaterials.">Is trehalose special for preserving dry biomaterials.</a> Biophysical Journal. 71 (4), 2087-2093 (1996).</li></ol>

The authors have nothing to disclose.

Biomimetic model systems are an important tool for studying the properties and interactions of proteins and membranes. Compared to other reconstituted systems like BLMs or supported lipid membranes, GUV based systems present several opportunities including considerable control of membrane composition, tension and geometry, as well as being truly oil-free. However, incorporating transmembrane proteins, such as KvAP, into GUVs requires significant adaptations of conventional protocols for lipid-only GUVs. The electroformat...

When studying the physical principles that govern living systems, bottom-up approaches allow an experimentalist to control system composition and other parameters that are not easily manipulated in cell-based systems1. For membrane-based processes, Giant Unilamellar Vesicles (GUVs, diameter ~ 1&#8211;100 &#181;m) have proven to be a very useful biomimetic system2&#8211;7 as they are well suited for microscopy studies and micromanipulation8&#8211;10. While there are many different protocols to produce GUVs, most fall into two categories &#8212; emulsion based approaches11,12 and techniques based on rehydrating a lipid film13&#8211;16. In emulsion-based methods, the inner and outer leaflets of the GUV membranes are assembled sequentially from lipid monolayers at water/oil interfaces. This approach is ideal for encapsulating soluble proteins within the GUVs, and for forming GUVs with asymmetric leaflet lipid composition. However, GUVs formed from emulsions can retain traces of solvent that change the membrane&#8217;s mechanical properties17, and the approach is not especially well-suited to trans-membrane protein reconstitution.
Film rehydration methods rely on the fact that drying (dehydration) causes many lipid mixtures to form a multi-lamellar stack of membranes. If this stack is then placed in contact with an aqueous buffer, membranes in the stack will move apart as solvent flows between them and at the surface of the stack, individual membranes can detach to form GUVs13,18 (as well a veritable zoo of other lipidic objects). However, even for optimal buffer and lipid compositions, this classical &#8220;spontaneous swelling&#8221; method has a relatively low yield of defect-free GUVs. One widely used method to boost the yield of defect-free GUVs is &#8220;electroformation&#8221;, in which an alternating current (AC) field is applied during film rehydration. While the mechanism remains poorly understood, &#8220;electroformation&#8221; can give spectacular GUV yields (&#62; 90% in favorable circumstances) for low salt concentration buffers (&#60; 5 mM)14,19, and can even work in physiological buffers (~100 mM) using a higher frequency (500 Hz versus 10 Hz) AC field and platinum electrodes15. An alternative approach to boost the yield of defect-free GUVs is &#8220;gel-assisted swelling&#8221;, in which the lipid solution is deposited onto a polymeric gel substrate rather than the passive (e.g., glass, PTFE) substrates used in classical &#8220;spontaneous swelling&#8221;. When the resultant lipid/gel film is rehydrated, GUVs can rapidly form even for physiological buffers16,20.
All these methods can produce lipid-only GUVs which can be used to study membrane associated phenomena such as the interaction between soluble proteins and membranes. However, to incorporate a trans-membrane protein into GUVs, significant modifications are needed to ensure that the protein remains in a functional state throughout the reconstitution procedure. While solutions of lipids in organic solvents (e.g., chloroform, cyclohexane) are ideal for producing lipid films, trans-membrane proteins are typically only stable when their hydrophobic trans-membrane domain is embedded in a lipid bilayer, or surrounded by a detergent micelle (e.g., during protein purification). Thus, the starting material for a reconstitution is typically native membranes, purified protein in a detergent solution, or small unilamellar protein-containing vesicles (proteo-SUVs) and/or multi-lamellar vesicles (proteo-MLVs) formed by detergent removal in the presence of lipids. Most methods to incorporate these membrane proteins into GUVs fall into three categories.
Direct Insertion: Trans-membrane protein suspended in detergent is mixed with pre-formed, lipid-only, mildly detergent solubilized GUVs, and the detergent then removed using biobeads21. While conceptually simple, this method requires precise control of the detergent concentration, as too high a detergent concentration can dissolve the GUVs while too low a concentration can cause the protein to unfold or aggregate.
GUV/Proteo-SUV Fusion: Protein in proteo-SUVs is combined with pre-formed, lipid-only GUVs and fusion is facilitated with special fusogenic peptides22 or detergent21. Typically the extent of fusion is limited leading to GUVs with low protein density.
Dehydration/Rehydration: A protein-containing lipid film is formed by partial dehydration of a proteo-SUV (or proteo-MLV) solution and GUVs are then grown as for a pure lipid film. The obvious challenge is to protect the protein during the partial dehydration step23, but the method has been successfully used to reconstitute trans-membrane proteins such as Bacteriorhodopsin, Calcium-ATPase, Integrin and VDAC into GUVs7,23&#8211;25.
This article describes dehydration/rehydration protocols to make GUVs containing the voltage-gated potassium channel, KvAP, from the hyper-thermophilic Archaea, Aeropyrum pernix. KvAP has a high degree of homology to eukaryotic voltage dependent potassium channels26 and a known crystal structure27, making it a good model for studying the mechanism of voltage gating. Production of the proteo-SUVs has been described in detail previously and is not part of this tutorial26,28,29. Importantly, KvAP proteo-SUVs do not have to be produced for each GUV preparation, as they can be stored in small (e.g., 10 &#181;l) aliquots at -80 &#176;C for extended periods of time (&#62; 1 year). Electroformation or gel-assisted swelling can then be used to grow GUVs from the KvAP proteo-SUVs (or proteo-MLVs).
The key steps for the electroformation protocol are illustrated in Figure 1. Droplets of a solution of SUVs containing the protein are deposited on platinum wires (shown in Figure 2). Partial dehydration of the SUV suspension leads to the formation of a lipid protein film through the fusion of SUVs. During rehydration, an AC field is applied to the electrodes to assist the lipid layers to delaminate and form GUVs. A 10 Hz field works well when using &#8220;low-salt&#8221; (&#60; 5 mM) rehydration buffer28 and GUVs take several hours to grow. In contrast, physiological buffers (containing ~100 mM salt) work well with a lower voltage, 500 Hz AC field but require a prolonged (~12 hr) swelling period15. This method is based upon an earlier protocol using ITO slides24, but uses a custom chamber containing two platinum wires as shown in Figure 2 (see the discussion for design details and suggestions for simpler, improvised chambers).
Figure 3 illustrates the gel-assisted swelling method. The protocol works well with buffers with physiological salt concentrations, is rapid, and produces GUVs with a more homogeneous protein distribution. However, the yield of isolated, apparently defect-free GUVs (i.e., the GUV membrane is uniform at optical length-scales and does not enclose any objects) is lower, although it provides a sufficient number for patch-clamp and micro-manipulation experiments. This method was based on a protocol using agarose gel to produce lipid-only GUVs16 and requires less specialized equipment than the electroformation method.
The characterization of GUVs with fluorescence microscopy is described, as well as procedures using a standard patch-clamp set-up to measure KvAP activity in &#8220;inside-out&#8221; excised membrane patches.
Growing protein-containing GUVs can be more difficult than lipid-only GUVs. In particular, the final GUV yield can depend sensitively on exactly how the SUV solution is deposited and dehydrated to form the membrane stack. For someone without any previous experience with GUVs, it may be helpful to first grow lipid-only GUVs following a conventional protocol15,16 in which the membrane film is formed by depositing lipids from an organic solvent. Once the conventional protocol works well, SUV deposition and partial dehydration can then be mastered using lipid-only SUVs, which are also very helpful when adjusting the protocol for a new lipid composition. When GUVs grow reliably from lipid-only SUVs, it is then only a small step to produce protein-containing GUVs from proteo-SUVs.

<table><tbody><tr><td>DPhPC (1,2-diphytanoyl-&lt;em&gt;sn&lt;/em&gt;-glycero-3-phosphocholine)</td><td>Avanti Polar Lipids&amp;nbsp;</td><td>850356P</td><td></td></tr><tr><td>Egg PC L-&amp;alpha;-phosphatidylcholine (Egg, Chicken)&amp;nbsp;</td><td>Avanti Polar Lipids&amp;nbsp;</td><td>840051P</td><td></td></tr><tr><td>Egg PA L-&amp;alpha;-phosphatidic acid (Egg, Chicken)</td><td>Avanti Polar Lipids&amp;nbsp;</td><td>840101P</td><td></td></tr><tr><td>18:1 (&amp;Delta;9-&lt;em&gt;cis&lt;/em&gt;) PC (DOPC) 1,2-dioleoyl-&lt;em&gt;sn&lt;/em&gt;-glycero-3-phosphocholine</td><td>Avanti Polar Lipids&amp;nbsp;</td><td>850375P</td><td></td></tr><tr><td>Cholesterol (ovine wool, &gt;98%)&amp;nbsp;</td><td>Avanti Polar Lipids&amp;nbsp;</td><td>700000P</td><td></td></tr><tr><td>TRed-DHPE</td><td>Invirtogen</td><td>T-1395MP</td><td>labeled lipid</td></tr><tr><td>BPTR-Ceramide</td><td>Invirtogen</td><td>D-7540&amp;nbsp;</td><td>labeled lipid</td></tr><tr><td>Choloroform</td><td>VWR</td><td>22711.290</td><td>AnalaR Normapur</td></tr><tr><td>Acetone</td><td>VWR</td><td>20066.296</td><td>AnalaR Normapur</td></tr><tr><td>Ethanol</td><td>VWR</td><td>20821.330</td><td>AnalaR Normapur</td></tr><tr><td>Kimwipe</td><td>Kimtech</td><td>7552</td><td></td></tr><tr><td>Hamilton syringes&amp;nbsp;</td><td>Hamilton Bonaduz AG</td><td></td><td>diverse</td></tr><tr><td>Amber vials and Teflon cups</td><td>Sigma</td><td>SU860083 and SU860076</td><td></td></tr><tr><td>Parafilm</td><td>VWR</td><td>291-1214</td><td></td></tr><tr><td>Microcentrifuge tube</td><td>Eppendorf</td><td></td><td>diverse</td></tr><tr><td>Agarose</td><td>Euromex</td><td>LM3 (1670-B, Tg 25.7C, Tm 64C)</td><td></td></tr><tr><td>Sucrose</td><td>Sigma</td><td>84097-1kg</td><td></td></tr><tr><td>Glucose&amp;nbsp;</td><td>Sigma</td><td>G8270-1kg</td><td></td></tr><tr><td>Trehalose</td><td>Sigma</td><td>T9531-25G</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>P9333-1kg</td><td></td></tr><tr><td>Hepes</td><td>Sigma</td><td>H3375-100G</td><td></td></tr><tr><td>TRIS</td><td>Euromex</td><td>EU0011-A</td><td></td></tr><tr><td>Osmometer</td><td>Wescor</td><td>Vapro</td><td></td></tr><tr><td>pH meter</td><td>Schott instruments</td><td>Lab850</td><td></td></tr><tr><td>Sonicator</td><td>Elmasonic</td><td>S 180 H</td><td></td></tr><tr><td>Syringe filter 0.2 &amp;micro;m</td><td>Sartorius steim</td><td>Minisart 16532</td><td></td></tr><tr><td>Function generator</td><td>TTi</td><td>TG315</td><td></td></tr><tr><td>Platinum wires</td><td>Goodfellow</td><td>LS413074</td><td>99.99+%, d = 0.5 mm</td></tr><tr><td>Polytetrafluoroethylene</td><td>Goodfellow</td><td></td><td></td></tr><tr><td>Dow Corning &#039;high vacuum grease&#039;</td><td>VWR</td><td>1597418</td><td></td></tr><tr><td>sealing paste</td><td>Vitrex medical A/S, Denmark</td><td>REF 140014</td><td>Sigillum Wax</td></tr><tr><td>Cover slides 22 x 40 mm No1.5</td><td>VWR</td><td>631-0136</td><td></td></tr><tr><td>Cover slides 22 x 22 mm No1.5</td><td>VWR&amp;nbsp;</td><td>631-0125</td><td></td></tr><tr><td>Plasma cleaner</td><td>Harrick</td><td>PDC-32G</td><td>airplasma, setting &#039;high&#039;</td></tr><tr><td>Petri dishes</td><td>Falcon BD</td><td>REF 353001</td><td>3.5 cm x 1 cm</td></tr><tr><td>patch clamp amplifier</td><td>Axon instruments</td><td>Multiclamp700B</td><td></td></tr><tr><td>DAQ Card</td><td>National Instruments</td><td>PCI-6221</td><td></td></tr><tr><td>Labview</td><td>National Instruments</td><td>version 8.6</td><td></td></tr><tr><td>Glass pipettes boro silicate OD 1&amp;nbsp;mm ID 0.58&amp;nbsp;mm</td><td>Harvard Apparatus</td><td>GC100-15</td><td></td></tr><tr><td>Electrode holder</td><td>Warner Instruments&amp;nbsp;</td><td>Q45W-T10P</td><td></td></tr><tr><td>Micromanipulator</td><td>Sutter&amp;nbsp;</td><td>MP-285</td><td></td></tr><tr><td>Pipette puller</td><td>Sutter&amp;nbsp;</td><td>P-2000&amp;nbsp;</td><td></td></tr><tr><td>Camera</td><td>ProSilica&amp;nbsp;</td><td>GC1380</td><td></td></tr><tr><td>Zeiss microscope</td><td>Zeiss</td><td>Axiovert 135</td><td></td></tr><tr><td>Objective</td><td>Zeiss</td><td></td><td>40X long working distance, Phase contrast</td></tr><tr><td>Objective&amp;nbsp;</td><td>Zeiss</td><td></td><td>100X Plan-Apochromat NA 1.3</td></tr><tr><td>Filterset GFP (for Alexa-488)</td><td>Horiba</td><td>XF100-3</td><td></td></tr><tr><td>Filterset Cy3 (for TexasRed)</td><td>Horiba</td><td>XF101-2</td><td></td></tr><tr><td>beta-casein</td><td>Sigma&amp;nbsp;</td><td>C6905-1G</td><td></td></tr><tr><td>Confocal microscope</td><td>Nikon Eclipse TE 2000-E</td><td></td><td>D-Eclipse C1 confocal head</td></tr><tr><td>Objective</td><td>Nikon</td><td></td><td>Plan Fluor 100X NA 1.3</td></tr><tr><td>Matlab</td><td>Mathworks</td><td></td><td>for image processing and analysis of the current traces</td></tr></tbody></table>

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 growth of GUVs can be quickly evaluated by examining the growth chamber under the microscope. For electroformation, the GUVs tend to grow in bunches along the platinum wires, as shown in Figure 4. During gel-assisted swelling, GUVs appear as spherical structures that rapidly grow and fuse together (Figure 5).
Defect-free GUVs are more easily identified and evaluated after transferring to an observation chamber. Calibration measurements are needed to rigoro...

Watch this Scientific Journal Video about Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies at JoVE

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

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 ...

reconstitution-transmembrane-protein-voltage-gated-ion-channel-kvap

Research

JoVE Journal

Biology

19.0K Views.  Université Pierre et Marie Curie.  De reconstitutie van het transmembraan eiwit, KvAP, in grote unilamellaire vesicles (GUVs) is aangetoond voor twee dehydratatie-rehydratatie methode - electroformation en gel ondersteunde zwelling. In beide werkwijzen worden kleine unilamellaire blaasjes die het eiwit gefuseerd aan GUVs die vervolgens kan worden onderzocht door fluorescentie microscopie en patch-clamp elektrofysiologie vormen. 

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

Methods for Patch Clamp Capacitance Recordings from the Calyx

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal. Electrical recordings from the presynaptic terminal allow the measurement of action potentials, calcium channel currents, vesicle fusion (exocytosis) and subsequent membrane uptake (endocytosis). The fusion of vesicles containing neurotransmitter causes the vesicle membrane to be added to the cell membrane of the calyx. This increase in the amount of cell membrane is measured as an increase in capacitance. The subsequent reduction in capacitance indicates endocytosis, the process of membrane uptake or removal from the calyx membrane. Endocytosis, is necessary to maintain the structure of the calyx and it is also necessary to form vesicles that will be filled with neurotransmitter for future exocytosis events. Capacitance recordings at the calyx of Held have made it possible to directly and rapidly measure vesicular release and subsequent endocytosis in a mammalian CNS nerve terminal. In addition, the corresponding postsynaptic activity can be simultaneously measured by using paired recordings. Thus a complete picture of the presynaptic and postsynaptic electrical activity at a central nervous system synapse is achievable using this preparation. Here, the methods for slice preparation, morphological features for identification of calyces of Held, basic patch clamping techniques, and examples of capacitance recordings to measure exocytosis and endocytosis are presented.

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal.

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal. ...

Patch Clamp Recording of Ion Channels Expressed in  Xenopus Oocytes

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological measurement of single or multiple ion channels in cells. This technique can be applied to ion channels in both their native environment and expressed in heterologous cells, such as oocytes harvested from the African clawed frog, Xenopus laevis. Here, we describe the well-established technique of patch clamp recording from Xenopus oocytes. This technique is used to measure the properties of expressed ion channels either in populations (macropatch) or individually (single-channel recording). We focus on techniques to maximize the quality of oocyte preparation and seal generation. With all factors optimized, this technique gives a probability of successful seal generation over 90 percent. The process may be optimized differently by every researcher based on the factors he or she finds most important, and we present the approach that have lead to the greatest success in our hands.

This is intended as an introduction to patch clamp recording from Xenopus laevis oocytes. It covers vitelline membrane removal, formation of a gigaohm seal (gigaseal), and the optional conversion of the patch to the outside-out topology.

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological ...

Pressure-polishing Pipettes for Improved Patch-clamp Recording

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording. We first developed this method for fabricating pipettes suitable for recording from small (&lt;3 m) neuronal cell bodies. The basic principal is similar to glass-blowing and combines air pressure and heat to modify the shape of patch pipettes prepared by a conventional micropipette puller. It can be applied to so-called soft (soda lime) and hard (borosilicate) glasses. Generally speaking, pressure polishing can reduce pipette resistance by 25% without decreasing the diameter of the tip opening (Goodman and Lockery, 2000). It can be applied to virtually any type of glass and requires only the addition of a high-pressure valve and fitting to a microforge. This technique is essential for recording from ultrasmall cells (&lt;5 m) and can also improve single-channel recording by minimizing pipette resistance. The blunt shape is also useful for perforated-patch clamp recording since this tip shape results in a larger membrane bleb available for perforation.

This is a guide to modifying the shape of glass micropipettes. Specifically, by using heat and air pressure the taper is widened without increasing the tip opening, leading to lower pipette resistance. This is critical to obtain low noise recordings of small cells but is useful in many applications.

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording.  We first developed this method for fabricating pipettes suitable for ...

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 ...

Chemistry

Mutagenesis and Functional Analysis of Ion Channels Heterologously Expressed in Mammalian Cells

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying potassium (referred to as GIRK) channels, which are important for regulating the excitability of neurons. There are four different mammalian GIRK channel subunits (GIRK1-GIRK4) - we focus on GIRK2 because it forms a homotetramer. Stimulation of different types of G protein-coupled receptors (GPCRs), such as the muscarinic receptor (M2R), leads to activation of GIRK channels. Alcohol also directly activates GIRK channels. We will show how to mutate one amino acid by specifically changing one or more nucleotides in the cDNA for the GIRK channel. This mutated cDNA sequence will be amplified in bacteria, purified, and the presence of the point mutation will be confirmed by DNA sequencing. The cDNAs for the mutated and wild-type GIRK channels will be transfected into human embryonic kidney HEK293T cells cultured in vitro. Lastly, whole-cell patch-clamp electrophysiology will be used to study the macroscopic potassium currents through the ectopically expressed wild-type or mutated GIRK channels. In this experiment, we will examine the effect of a L257W mutation in GIRK2 channels on M2R-dependent and alcohol-dependent activation.

We will demonstrate how to study the effect of a single point mutation on the function of an ion channel.

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

The actin cytoskeleton, the principal mechanical machinery in the cell, mediates numerous essential physical cellular activities, including cell deformation, division, migration, and adhesion. However, studying the dynamics and structure of the actin network in vivo is complicated by the biochemical and genetic regulation within live cells. To build a minimal model devoid of intracellular biochemical regulation, actin is encapsulated inside giant unilamellar vesicles (GUVs, also called liposomes). The biomimetic liposomes are cell-sized and facilitate a quantitative insight into the mechanical and dynamical properties of the cytoskeleton network, opening a viable route for bottom-up synthetic biology. To generate liposomes for encapsulation, the inverted emulsion method (also referred to as the emulsion transfer method) is utilized, which is one of the most successful techniques for encapsulating complex solutions into liposomes to prepare various cell-mimicking systems. With this method, a mixture of proteins of interest is added to the inner buffer, which is later emulsified in a phospholipid-containing mineral oil solution to form monolayer lipid droplets. The desired liposomes are generated from monolayer lipid droplets crossing a lipid/oil-water interface. This method enables the encapsulation of concentrated actin polymers into the liposomes with desired lipid components, paving the way for in vitro reconstitution of a biomimicking cytoskeleton network.

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

In this manuscript, we demonstrate the experimental techniques to encapsulate the F-actin cytoskeleton into giant unilamellar lipid vesicles (also called liposomes), and the method to form a cortex-biomimicking F-actin layer at the inner leaflet of the liposome membrane.

The actin cytoskeleton, the principal mechanical machinery in the cell, mediates numerous essential physical cellular activities, including cell ...

Bioengineering

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.

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 ...

Reconstitution of a Kv Channel into Lipid Membranes for Structural and Functional Studies

To study the lipid-protein interaction in a reductionistic fashion, it is necessary to incorporate the membrane proteins into membranes of well-defined lipid composition. We are studying the lipid-dependent gating effects in a prototype voltage-gated potassium (Kv) channel, and have worked out detailed procedures to reconstitute the channels into different membrane systems. Our reconstitution procedures take consideration of both detergent-induced fusion of vesicles and the fusion of protein/detergent micelles with the lipid/detergent mixed micelles as well as the importance of reaching an equilibrium distribution of lipids among the protein/detergent/lipid and the detergent/lipid mixed micelles. Our data suggested that the insertion of the channels in the lipid vesicles is relatively random in orientations, and the reconstitution efficiency is so high that no detectable protein aggregates were seen in fractionation experiments. We have utilized the reconstituted channels to determine the conformational states of the channels in different lipids, record electrical activities of a small number of channels incorporated in planar lipid bilayers, screen for conformation-specific ligands from a phage-displayed peptide library, and support the growth of 2D crystals of the channels in membranes. The reconstitution procedures described here may be adapted for studying other membrane proteins in lipid bilayers, especially for the investigation of the lipid effects on the eukaryotic voltage-gated ion channels.

Procedures for complete reconstitution of a prototype voltage-gated potassium channel into lipid membranes are described. The reconstituted channels are suitable for biochemical assays, electrical recordings, ligand screening and electron crystallographic studies. These methods may have general applications to the structural and functional studies of other membrane proteins.

To study the lipid-protein interaction in a reductionistic fashion, it is necessary to incorporate the membrane proteins into membranes of well-defined ...

Pulling Membrane Nanotubes from Giant Unilamellar Vesicles

The reshaping of the cell membrane is an integral part of many cellular phenomena, such as endocytosis, trafficking, the formation of filopodia, etc. Many different proteins associate with curved membranes because of their ability to sense or induce membrane curvature. Typically, these processes involve a multitude of proteins making them too complex to study quantitatively in the cell. We describe a protocol to reconstitute a curved membrane in vitro, mimicking a curved cellular structure, such as the endocytic neck. A giant unilamellar vesicle (GUV) is used as a model of a cell membrane, whose internal pressure and surface tension are controlled with micropipette aspiration. Applying a point pulling force on the GUV using optical tweezers creates a nanotube of high curvature connected to a flat membrane. This method has traditionally been used to measure the fundamental mechanical properties of lipid membranes, such as bending rigidity. In recent years, it has been expanded to study how proteins interact with membrane curvature and the way they affect the shape and the mechanics of membranes. A system combining micromanipulation, microinjection, optical tweezers, and confocal microscopy allows measurement of membrane curvature, membrane tension, and the surface density of proteins, concurrently. From these measurements, many important mechanical and morphological properties of the protein-membrane system can be inferred. In addition, we lay out a protocol of creating GUVs in the presence of physiological salt concentration, and a method of quantifying the surface density of proteins on the membrane from fluorescence intensities of labeled proteins and lipids.

Many proteins in the cell sense and induce membrane curvature. We describe a method to pull membrane nanotubes from lipid vesicles to study the interaction of proteins or any curvature-active molecule with curved membranes in vitro.

The reshaping of the cell membrane is an integral part of many cellular phenomena, such as endocytosis, trafficking, the formation of filopodia, etc. Many ...

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

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