This work was supported by National Institutes of Health grants R01-HL70393 and R01-NS060706 to J.C. J.C. is a Professor of Biomedical Engineering on the Spencer T. Olin Endowment.

1. Injection of mRNA into Oocytes
<ol>
	<li>Inject 0.05 - 50 ng messenger RNA that was transcribed in vitro into Xenopus laevis oocytes (stage V-VI) using Nanoject II Auto-Nanoliter Injector (Drummond Scientific Company, model 3-000-204). Repeat the injection on a dozen oocytes for each mRNA.</li>
	<li>Rinse the injected oocytes twice using ND-96 solution (96 mM sodium chloride (NaCl, Mr 58.44 g/mol), 2 mM potassium chloride (KCl, Mr 74.56 g/mol), 1.8 mM calcium chloride (CaCl2&#8226;2H2O, Mr 147.02), 1 mM magnesium chloride (MgCl2&#8226;6H2O, Mr 203.31 g/mol), 5 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES, Mr 238.31 g/mol), 2.5 mM sodium pyruvate (Mr 110.04 g/mol), and 1x Penicillin-Streptomycin. pH 7.6), then place them in a culture plate and incubate at 18&deg;C for 2+ days.</li>
</ol>
2. Preparing the Perfusion System
<img src="/files/ftp_upload/2269/2269fig1.jpg" alt="Figure 1" /> Illustration of the Automate ValveLink 16 perfusion system. The perfusion system uses pressurized nitrogen to push perfusion solutions out of the solution reservoirs, through the perfusion tubings, and to the perfusion pencil and tip. The flow of each stream of perfusion solution is controlled by one reservoir valve and one electronic valve. In this protocol, one of the reservoirs is not pressurized and functions as a waste collector as well as a pressure-release mechanism. (Picture adapted from vendor's website http://www.autom8.com)
<ol>
	<li>Connect the tubings to the perfusion pencil. Turn on the electronic valve controller and open the electronic valves.</li>
	<li>To assure that the tubings and the perfusion pencil are free of clog and air bubbles, push deionized (DI) water through each tubing using a syringe filled with deionized water.</li>
	<li>Connect the tubings to the solution reservoirs and open the valve of gas cylinder to apply pressurized nitrogen.</li>
	<li>Open the reservoir valve to fill the tubing with reservoir solution. Flick the reservoir valve to remove bubbles in the solution if needed. Close the valve after the tubing is filled with solution.</li>
	<li>Fill the perfusion tip with water and screw it onto the perfusion pencil. Be careful not to trap any bubbles when connecting the tip.</li>
	<li>Fix the perfusion pencil to the bath stage using modeling clay. Make sure the perfusion tip is in the bath space. The perfusion system is now set up.</li>
	<li>Add DI water in the bath. Make sure the perfusion tip is submerged in the water.</li>
	<li>Test that each perfusion solution comes out of the perfusion tip correctly. Close the electronic valve connected to the tubing of waste collector and open the valve for one perfusion solution at a time (140 mM potassium hydroxide (KOH, Mr 56.1 g/mol), 20 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES, Mr 238.31 g/mol), 2 mM potassium chloride (KCl, Mr 74.56 g/mol), 1 mM Ethylene glycol-bis(2-aminoethylether)-N,N,N&#8242;,N&#8242;-tetraacetic acid (EGTA, Mr 380.35 g/mol), pH is adjusted to 7.1-7.2 by methanesulfonic acid(MeSO3, Mr 96.1 g/mol), Ca2+ or Mg2+ is added to desired concentration when needed). Under the microscope, observe the jet of perfusion fluid coming out of the perfusion tip. Close the valve for perfusion solution and open the valve for the waste collector. The perfusion jet should almost disappear. Repeat the same test for all the perfusion solutions.</li>
	<li>When you have verified that all the perfusion solutions flow correctly, replace DI water in the bath with bath solution (140 mM potassium hydroxide (KOH, Mr 56.1 g/mol), 20 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES, Mr 238.31 g/mol), 2 mM potassium chloride (KCl, Mr 74.56 g/mol), pH is adjusted to 7.1-7.2 by methanesulfonic acid(MeSO3, Mr 96.1 g/mol)).</li>
</ol>
3. Patch Clamping
<ol>
	<li>Recoat the Ag recording electrode with AgCl before each patch clamping session. First, remove the electrode wire from the pipette holder and submerge the tip half of the electrode wire in a vial containing fresh bleach for at least 15 minutes. This deposits a layer of AgCl on the wire.</li>
	<li>Rinse the electrode wire with deionized water and blot dry. Then install the Ag/AgCl electrode back in the pipette holder.</li>
	<li>Connect the bath (reference) electrode to the headstage and place it in the bath. This prepares the electrodes for recording.</li>
	<li>Now prepare for data acquisition. Turn on your computer and plug the hardware key into the printer port of your computer. The hardware key must be plugged for you to use the data acquisition software, which is HEKA pulse.</li>
	<li>Switch on the amplifier (Axon Instruments, AXOPATCH 200B) and start HEKA Pulse. Load the protocol file and adjust the configuration settings, such as the Stimulus Scale. The goal for adjusting the configuration settings is to assure that the test potential is exactly equal to the command potential. This prepares the data acquisition equipment.</li>
	<li>Prepare oocytes. Submerge the oocyte in stripping solution (200 mM N-Methyl-D-glucamine (NMG, Mr 195.22 g/mol), 200 mM Aspartate (no K+) (Mr 133.1 g/mol), 2 mM potassium chloride (KCl, Mr 74.56 g/mol), 10 mM Ethylene glycol-bis(2-aminoethylether)-N,N,N&#8242;,N&#8242;-tetraacetic acid (EGTA, Mr 380.35 g/mol), 1 mM magnesium chloride (MgCl2&#8226;6H2O, Mr 203.31 g/mol), 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES, Mr 238.31 g/mol), pH is adjusted to 7.4 by sodium hydroxide (NaOH, Mr 40.0 g/mol) for 5 - 10 min. The stripping solution detaches the vitelline membrane from the plasma membrane which makes it possible to strip the vitelline membrane.</li>
	<li>Gently strip the vitelline membrane from the oocyte using two pairs of forceps. A devitellinized oocyte is extremely fragile and therefore should be moved as little as necessary.</li>
	<li>Taking care to prevent exposing the devitellinized oocyte to air or bubbles, use a glass pipette filled with enough solution to transfer the oocyte to the bath. This prepares the oocyte for recording.</li>
	<li>Prepare patch pipettes. Pull the glass pipettes after inserting a glass capillary tube (VWR International, Cat # 53432-921) into a Sutter P-97 Flaming/Brown Micropipette Puller. Observe the pipettes under microscope to determine the tip shape and opening diameter. The opening diameter should be 2-4 micrometers.</li>
	<li>To reduce capacitive current during recording and to ensure a smooth tip, coat the tip with wax and then fire-polish it.</li>
	<li>Fill the pipette tip by placing the tip of the pipette inside the pipette solution (140 mM potassium hydroxide (KOH, Mr 56.1 g/mol), 20 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES, Mr 238.31 g/mol), 2 mM potassium chloride (KCl, Mr 74.56 g/mol), 2 mM magnesium chloride (MgCl2&#8226;6H2O, Mr 203.31 g/mol), pH is adjusted to 7.1-7.2 by methanesulfonic acid(MeSO3, Mr 96.1 g/mol)) and drawing the plunger. This wets the tip. </li>
	<li>Fill pipette 1/3 full with pipette solution using a syringe and place the pipette in the pipette holder with the Ag/AgCl electrode inside and contacting with the pipette solution. This prepares the patch pipette.</li>
	<li>To excise an inside-out patch from the prepared oocyte, find the clear edge of oocyte under microscope. Move the patch pipette close to the oocyte using manipulator. Record the serial resistance of pipette. The ideal series resistance of the patch pipette is between 1-1.5 M&Omega; when filled with pipette solution and submerged in bath solution.</li>
	<li>Slowly push the pipette against the oocyte until the resistance approximately doubles.</li>
	<li>Apply gentle suction to the membrane by mouth through a suction tube which is connected to the pipette holder until a giga-ohm seal is obtained. Stabilize the patch by holding it at voltage -30 mV for a couple of minutes.</li>
	<li>To obtain an inside-out patch, excise the patch by quickly pushing the pipette further into the oocyte and then gently pulling it out.</li>
	<li>The inside-out patch is now ready for clamping. Move the pipette holding the inside-out patch to about 100 microns away from the opening of the perfusion tip.</li>
	<li>Close the electronic valve for waste collecting and open the reservoir valve for the desired perfusion solution</li>
	<li>Begin recording currents across the patch. Change the voltage and perfusion solutions as desired.</li>
	<li>When you are finished recording, clean the perfusion tubings, pencil and tip by pushing through deionized water to avoid clogs.</li>
</ol>
4. Representative Results
During the preparation of oocytes for patch clamp, the 5-10 min treatment of stripping solution would detach the vitelline membrane from the plasma membrane, which makes possible stripping the vitelline membrane.
The ideal series resistance of the patch pipette is between 1-1.5 M&Omega; when filled with pipette solution and submerged in bath solution.
Following is a representative recording of wild-type BK channels on an oocyte membrane patch. On the left top, the square waves indicate the voltage applied to the patch - the second step of voltage increases from 50 mV to 200 mV with increment of 50 mV. Below is the corresponding current traces when the perfusion solution has nominal 0 Ca2+ (free [Ca2+] is about 0.5 nM). The increase of current amplitude indicates that the open probability of BK channels increases with voltage. On the right side, the same patch is perfused with 1.0 &mu;M Ca2+ when applied with the same voltage protocol. The current amplitude increases more under the same voltage, indicating that the open probability increases with [Ca2+]. <img src="/files/ftp_upload/2269/2269fig2.jpg" alt="Figure 2" />

<ol>
	<li>Methfessel, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patch+clamp+measurements+on+Xenopus+laevis+oocytes:+currents+through+endogenous+channels+and+implanted+acetylcholine+receptor+and+sodium+channels.">Patch clamp measurements on Xenopus laevis oocytes: currents through endogenous channels and implanted acetylcholine receptor and sodium channels.</a> Pfl&#252;gers Arch. 407, 577-588 (1986).</li><li>Toro, L., Wallner, M., Meera, P., Tanaka, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maxi-K+(Ca),+Unique+Member+of+the+Voltage-Gated+K+Channel+Superfamily.">Maxi-K (Ca), Unique Member of the Voltage-Gated K Channel Superfamily.</a> News Physiol. Sci. 13, 112-117 (1998).</li><li>Fettiplace, R., Fuchs, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+hair+cell+tuning.">Mechanisms of hair cell tuning.</a> Annu. Rev. Physiol. 61, 809-834 (1999).</li><li>Du, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium-sensitive+potassium+channelopathy+in+human+epilepsy+and+paroxysmal+movement+disorder.">Calcium-sensitive potassium channelopathy in human epilepsy and paroxysmal movement disorder.</a> Nat. Genet. 37, 733-738 (2005).</li><li>Ledoux, J., Werner, M. E., Brayden, J. E., Nelson, M. T., T, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium-activated+potassium+channels+and+the+regulation+of+vascular+tone.">Calcium-activated potassium channels and the regulation of vascular tone.</a> Physiology (Bethesda). 21, 69-78 (2006).</li><li>Salkoff, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-conductance+potassium+channels+of+the+SLO+family.">High-conductance potassium channels of the SLO family.</a> Nat. Rev. Neurosci. 7, 921-931 (2006).</li><li>Shi, J., Cui, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracellular+Mg2++enhances+the+function+of+BK-type+Ca2+-activated+K++channels.">Intracellular Mg2+ enhances the function of BK-type Ca2+-activated K+ channels.</a> J. Gen. Physiol. 118, 589-606 (2001).</li><li>Magleby, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gating+mechanism+of+BK+(Slo1)+channels:+so+near,+yet+so+far.">Gating mechanism of BK (Slo1) channels: so near, yet so far.</a> J. Gen. Physiol. 121, 81-96 (2003).</li><li>Latorre, R., Brauchi, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Large+conductance+Ca2+-activated+K++(BK)+channel:+activation+by+Ca2++and+voltage.">Large conductance Ca2+-activated K+ (BK) channel: activation by Ca2+ and voltage.</a> Biol. Res. 39, 385-401 (2006).</li><li>Cox, D. H., Cui, J., Aldrich, R. W., W, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Allosteric+gating+of+a+large+conductance+Ca-activated+K++channel.">Allosteric gating of a large conductance Ca-activated K+ channel.</a> J. Gen. Physiol. 110, 257-281 (1997).</li><li>Cui, J., Aldrich, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Allosteric+linkage+between+voltage+and+Ca2+-dependent+activation+of+BK-type+mslo1.">Allosteric linkage between voltage and Ca2+-dependent activation of BK-type mslo1.</a> K+ channels. Biochemistry. 39, 15612-15619 (2000).</li><li>Yang, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+Slo1+BK+channels+by+Mg2++coordinated+between+the+voltage+sensor+and+RCK1.">Activation of Slo1 BK channels by Mg2+ coordinated between the voltage sensor and RCK1.</a> 15, 1152-1159 (2008).</li><li>Lee, U. S., Cui, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term={beta}+subunit-specific+modulations+of+BK+channel+function+by+a+mutation+associated+with+epilepsy+and+dyskinesia.">{beta} subunit-specific modulations of BK channel function by a mutation associated with epilepsy and dyskinesia.</a> J. Physiol. , 587-1481 (2009).</li></ol>

No conflicts of interest declared.

The oocyte expression system is ideal for electrophysiological characterization of voltage-dependent ion channels due to relatively low background of endogenous channels. Furthermore, since it is a transient expression system, it provides an efficient method of performing mutagenesis study of these channels. However, it should be noted that the oocyte expression system is different from that in mammalian cells, thus there may be differences in post-translational modification and association with different subunits. Furth...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Nanoject II Auto-Nanoliter Injector</td>
<td>&nbsp;</td>
<td>Drummond Scientific Company</td>
<td>3-000-204</td>
<td>&nbsp;</td>
<tr>
<td>P-97 Flaming/Brown Micropipette Puller</td>
<td>&nbsp;</td>
<td>Sutter Instruments</td>
<td>P-97</td>
<td>&nbsp;</td>
<tr>
<td>Perfusion System and Electronic Controller</td>
<td>&nbsp;</td>
<td>AutoMate Scientific, Inc.</td>
<td>ValveLink 16</td>
<td>Inner diameter of perfusion tip: 100 microns</td>
</tr>
<tr>
<td>Inverted Microscope</td>
<td>&nbsp;</td>
<td>Olympus</td>
<td>CKX31</td>
<td>&nbsp;</td>
<tr>
<td>Glass Pipettes</td>
<td>&nbsp;</td>
<td>VWR International</td>
<td>53432-921</td>
<td>&nbsp;</td>
<tr>
<td>Flaming/Brown Micropipette Puller</td>
<td>&nbsp;</td>
<td>Sutter Instrument Co.</td>
<td>P-97</td>
<td>&nbsp;</td>
<tr>
<td>Amplifier</td>
<td>&nbsp;</td>
<td>Axon Instruments</td>
<td>AXOPATCH 200B</td>
<td>&nbsp;</td>
<tr>
<td>Computer Interface</td>
<td>&nbsp;</td>
<td>INSTRUTECH Corporation</td>
<td>ITC-18</td>
<td>&nbsp;</td>
<tr>
<td>Headstage</td>
<td>&nbsp;</td>
<td>Axon Instruments</td>
<td>CV 203BU</td>
<td>&nbsp;</td>		</tbody>
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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.

Watch this Scientific Journal Video about Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes at JoVE.com

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.

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 ...

patch-clamp-perfusion-techniques-for-studying-ion-channels-expressed

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20.8K Views.  Washington University in St. Louis. 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.

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

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 ...

Microinjection of Xenopus Laevis Oocytes

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic transport. Because of its large nucleus and high density of nuclear pore complexes (NPCs), nuclear transport can be easily visualized in the Xenopus oocyte. Much insight into the mechanisms of nuclear import and export has been gained through use of this system (reviewed by Pant&eacute;, 2006). In addition, we have used microinjection of Xenopus oocytes to dissect the nuclear import pathways of several viruses that replicate in the host nucleus. 
Here we demonstrate the cytoplasmic microinjection of Xenopus oocytes with a nuclear import substrate. We also show preparation of the injected oocytes for visualization by thin-sectioning EM, including dissection, dehydration, and embedding of the oocytes into an epoxy embedding resin. Finally, we provide representative results for oocytes that have been microinjected with the capsid of the baculovirus Autographa californica nucleopolyhedrovirus (AcMNPV) or the parvovirus Minute Virus of Mice (MVM), and discuss potential applications of the technique.

Here we demonstrate cytoplasmic microinjection of Xenopus laevis oocytes with a nuclear import substrate, as well as preparation of the injected oocytes for visualization by thin-sectioning electron microscopy.

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic ...

Visualizing RNA Localization in Xenopus Oocytes

RNA localization is a conserved mechanism of establishing cell polarity. Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to spatially restrict gene expression of Vg1 protein. Tight control of Vg1 distribution in this manner is required for proper germ layer specification in the developing embryo. RNA sequence elements in the 3' UTR of the mRNA, the Vg1 localization element (VLE) are required and sufficient to direct transport. To study the recognition and transport of Vg1 mRNA in vivo, we have developed an imaging technique that allows extensive analysis of trans-factor directed transport mechanisms via a simple visual readout. 
To visualize RNA localization, we synthesize fluorescently labeled VLE RNA and microinject this transcript into individual oocytes. After oocyte culture to allow transport of the injected RNA, oocytes are fixed and dehydrated prior to imaging by confocal microscopy. Visualization of mRNA localization patterns provides a readout for monitoring the complete pathway of RNA transport and for identifying roles in directing RNA transport for cis-acting elements within the transcript and trans-acting factors that bind to the VLE (Lewis et al., 2008, Messitt et al., 2008). We have extended this technique through co-localization with additional RNAs and proteins (Gagnon and Mowry, 2009, Messitt et al., 2008), and in combination with disruption of motor proteins and the cytoskeleton (Messitt et al., 2008) to probe mechanisms underlying mRNA localization.

Visualizing RNA Localization in Xenopus Oocytes

Visualization of in vivo RNA transport is accomplished by microinjection of fluorescently labeled RNA transcripts into Xenopus oocytes, followed by confocal microscopy.

RNA localization is a conserved mechanism of establishing cell polarity.  Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to ...

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 ...

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 ...

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 ...

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

Chromosome segregation in human oocytes is error prone, resulting in aneuploidy, which is the leading genetic cause of miscarriage and birth defects. The study of chromosome behavior in oocytes from model organisms holds much promise to uncover the molecular basis of the susceptibility of human oocytes to aneuploidy. Drosophila melanogaster is amenable to genetic manipulation, with over 100 years of research, community, and technique development. Visualizing chromosome behavior and spindle assembly in Drosophila oocytes has particular challenges, however, due primarily to the presence of membranes surrounding the oocyte that are impenetrable to antibodies. We describe here protocols for the collection, preparation, and imaging of meiosis I spindle assembly and chromosome behavior in Drosophila oocytes, which allow the molecular dissection of chromosome segregation in this important model organism.

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

We present protocols for the collection, preparation, and imaging of mature Drosophila oocytes. These methods allow the visualization of chromosome behavior and spindle assembly and function during meiosis.

Chromosome segregation in human oocytes is error prone, resulting in aneuploidy, which is the leading genetic cause of miscarriage and birth defects. The ...