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>

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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 border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments (optional)</td>
		</tr>
		<tr>
			<td>Digital Multimeter</td>
			<td>Agilent Technologies, <a href="http://www.agilent.com" target="_blank">www.agilent.com</a></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>&nbsp;</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, <a href="http://www.agilent.com" target="_blank">www.agilent.com</a></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 &Omega; load</td>
		</tr>
		<tr>
			<td>ITO coated glass slides</td>
			<td>Delta Technologies, Loveland, CO <a href="http://www.delta-technologies.com" target="_blank">www.delta-technologies.com</a></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 <a href="http://www.omega.com" target="_blank">www.omega.com</a></td>
			<td>CNi3233 or similar</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Hygrometer</td>
			<td>Extech, Nashua, NH, <a href="http://www.extech.com" target="_blank">www.extech.com</a></td>
			<td>445815</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Silicone rubber sheet</td>
			<td>McMaster-Carr Elmhurst, IL <a href="http://www.mcmaster.com" target="_blank">www.mcmaster.com</a></td>
			<td><a href="http://www.mcmaster.com/#87315K64" target="_blank">87315K64</a></td>
			<td>Use USP Grade VI silicone for its high purity</td>
		</tr>
		<tr>
			<td>EMI gasket</td>
			<td>Laird Technologies <a href="http://www.lairdtech.com" target="_blank">www.lairdtech.com</a></td>
			<td>4202-PA-51H-01800 or similar</td>
			<td>Distributed by Mouser <a href="http://www.mouser.com" target="_blank">www.mouser.com</a></td>
		</tr>
		<tr>
			<td>TxR-DHPE</td>
			<td>Life Technologies, Carlsbad, CA <a href="http://www.lifetechnologies.com" target="_blank">www.lifetechnologies.com</a></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 <a href="http://www.avantilipids.com" target="_blank">www.avantilipids.com</a></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>&nbsp;</td>
		</tr>
		<tr>
			<td>Cholesterol</td>
			<td>Sigma-Aldrich</td>
			<td>C8667</td>
			<td>&nbsp;</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

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Biology

22.1K Views.  University of Washington. إعادة تشكيل بروتينات الغشاء وظيفية في الجسيمات الشحمية العملاقة من تكوين المعرفة هو نهج قوية عندما يقترن الكهربية التصحيح، المشبك. ومع ذلك، قد يكون الإنتاج الحويصلية العملاقة التقليدية يتعارض مع استقرار البروتين. نحن تصف بروتوكولات لإنتاج الجسيمات الشحمية ال...

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

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 ...

Preparation of Artificial Bilayers for Electrophysiology Experiments

Planar lipid bilayers, also called artificial lipid bilayers, allow you to study ion-conducting channels in a well-defined environment. These bilayers can be used for many different studies, such as the characterization of membrane-active peptides, the reconstitution of ion channels or investigations on how changes in lipid bilayer properties alter the function of bilayer-spanning channels.  Here, we show how to form a planar bilayer and how to isolate small patches from the bilayer, and in a second video  will also demonstrate a procedure for using gramicidin channels to determine changes in lipid bilayer elastic properties.   We also demonstrate the individual steps needed to prepare the bilayer chamber, the electrodes and how to test that the bilayer is suitable for single-channel measurements.

Planar lipid bilayers, also called artificial lipid bilayers, allow you to study ion-conducting channels in a well-defined environment. Here, we demonstrate the individual steps needed to prepare the bilayer chamber, the electrodes and how to test that the bilayer is suitable for single-channel measurements.

Planar lipid bilayers, also called artificial lipid bilayers, allow you to study ion-conducting channels in a well-defined environment. These bilayers can ...

Spinal Cord Electrophysiology

The neonatal mouse spinal cord is a model for studying the development of neural circuitries and locomotor movement. We demonstrate the spinal cord dissection and preparation of recording bath artificial cerebrospinal fluid used for locomotor studies. Once dissected, the spinal cord ventral nerve roots can be attached to a recording electrode to record the electrophysiologic signals of the central pattern generating circuitry within the lumbar cord.

A demonstration of the isolation of neonatal mouse spinal cord for electrophysiologic studies.

The neonatal mouse spinal cord is a model for studying the development of neural circuitries and locomotor movement. We demonstrate the spinal cord ...

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 ...

Exploring Arterial Smooth Muscle Kv7 Potassium Channel Function using Patch Clamp Electrophysiology and Pressure Myography

Contraction or relaxation of smooth muscle cells within the walls of resistance arteries determines the artery diameter and thereby controls flow of blood through the vessel and contributes to systemic blood pressure. The contraction process is regulated primarily by cytosolic calcium concentration ([Ca2+]cyt), which is in turn controlled by a variety of ion transporters and channels. Ion channels are common intermediates in signal transduction pathways activated by vasoactive hormones to effect vasoconstriction or vasodilation. And ion channels are often targeted by therapeutic agents either intentionally (e.g. calcium channel blockers used to induce vasodilation and lower blood pressure) or unintentionally (e.g. to induce unwanted cardiovascular side effects).
Kv7 (KCNQ) voltage-activated potassium channels have recently been implicated as important physiological and therapeutic targets for regulation of smooth muscle contraction. To elucidate the specific roles of Kv7 channels in both physiological signal transduction and in the actions of therapeutic agents, we need to study how their activity is modulated at the cellular level as well as evaluate their contribution in the context of the intact artery.
The rat mesenteric arteries provide a useful model system. The arteries can be easily dissected, cleaned of connective tissue, and used to prepare isolated arterial myocytes for patch clamp electrophysiology, or cannulated and pressurized for measurements of vasoconstrictor/vasodilator responses under relatively physiological conditions. Here we describe the methods used for both types of measurements and provide some examples of how the experimental design can be integrated to provide a clearer understanding of the roles of these ion channels in the regulation of vascular tone.

Measurements of Kv7 (KCNQ) potassium channel activity in isolated arterial myocytes (using patch clamp electrophysiological techniques) in parallel with measurements of constrictor/dilator responses (using pressure myography) can reveal important information about the roles of Kv7 channels in vascular smooth muscle physiology and pharmacology.

Contraction  or relaxation of smooth muscle cells within the walls of resistance arteries  determines the artery diameter and thereby controls flow of ...

Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ring-supported hydrophilized PTFE membrane to provide structural support to both living and fixed tissue during immunohistochemical processing, which allows for the use of a variety of protocols that would otherwise cause damage to the tissue. First, this is demonstrated with bolus loading of fluorescent markers into living retinal tissue. This method allows for quick visualization of targeted structures, while the membrane support maintains tissue integrity during the injection and allows for easy transfer of the preparation for further imaging or processing.
Second, a procedure for antibody staining in tissue fixed with carbodiimide is described. Though paraformaldehyde fixation is more common, carbodiimide fixation provides a superior substrate for the visualization of synaptic proteins. A limitation of carbodiimide is that the resulting fixed tissue is relatively fragile; however, this is overcome with the use of the supporting membrane. Retinal tissue is used to demonstrate these techniques, but they may be applied to any fragile tissue.

The protocol described here is for structural assessment of a wholemount retinal preparation. This includes descriptions of tissue dissection, mounting onto a hydrophilized polytetrafluoroethylene (PTFE) membrane insert, bolus loading with fluorescent markers, and a comparison of fixation with carbodiimide and paraformaldehyde for immunohistochemical analysis of cellular and synaptic components.

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ...

One-channel Cell-attached Patch-clamp Recording

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate depends on properties intrinsic to each channel protein as well as the mechanism by which it is generated and controlled and represents an important area of current research. Information about the operational dynamics of ion channel proteins can be obtained by observing long stretches of current produced by a single molecule. Described here is a protocol for obtaining one-channel cell-attached patch-clamp current recordings for a ligand gated ion channel, the NMDA receptor, expressed heterologously in HEK293 cells or natively in cortical neurons. Also provided are instructions on how to adapt the method to other ion channels of interest by presenting the example of the mechano-sensitive channel PIEZO1. This method can provide data regarding the channel&#8217;s conductance properties and the temporal sequence of open-closed conformations that make up the channel&#8217;s activation mechanism, thus helping to understand their functions in health and disease.

Described here is a procedure for obtaining long stretches of current recording from one ion channel with the cell-attached patch-clamp technique. This method allows for observing, in real time, the pattern of open-close channel conformations that underlie the biological signal. These data inform about channel properties in undisturbed biological membranes.

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate ...

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 ...