This work was supported by F31NS086765 (KAC), F31NS076235 (MAP), and R01 NS052669 (GKP) and EIA9100012.&#160; The authors thank Eileen Kasperek for expertise and assistance with molecular biology and tissue culture; and Jason Myers for sharing data obtained from early prefrontal cortical neurons. &#160;

1. Cell Culture and Protein Expression
<ol>
	<li>Maintain HEK293 cells (ATCC number CRL-1573) between passages 22 and 40, which includes passages performed by ATCC, in monolayer culture in DMEM supplemented with 10% Fetal Bovine Serum (FBS) and 1% penicillin/streptomycin mixture at 5% CO2 and 37&#160;&#176;C. Between experiments, passage cells into T25 flasks at 5-20 fold dilutions in a final volume of 10 ml. Note: Using cells during these passages guarantees favorable cell health which will allow for optimal seal formation and patch stability.</li>
	<li>For transfection, plate HEK293 cells in 35 mm dishes at a density of ~105 cells/dish, which corresponds to ~0.5-0.6 ml of cell suspension per dish and grow cells in 2 ml medium for 18-24 hr.</li>
	<li>In a sterile 1.5 ml centrifuge tube, prepare the transfection mixture for four 35 mm dishes by adding (in this order): (1) 1 &#956;g of each cDNA (GluN1, GluN2A, and GFP); (2) 315 &#956;l of double distilled water (ddH2O); (3) 350 &#956;l of 42 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), and (4) 35 &#956;l of 2.5 M CaCl2 drop wise, which is used to form the precipitate.</li>
	<li>Vortex tube for 5 sec and add 175 &#956;l of transfection suspension in each of the four 35 mm dishes plated the day before, which now contain cells at 50-60% confluence, and incubate the cells at 37 &#176;C for 2 hr.</li>
	<li>Aspirate off the transfection medium, wash with PBS and replace with 2 ml of growth medium supplemented with 2 mM MgCl2 to prevent NMDA receptor mediated excitotoxicity9. Note: The cells can be used for electrophysiological recordings 24-48 hr post transfection.</li>
</ol>
2. Electrode Preparation
<ol>
	<li>Generate 2 symmetrical recording pipettes by pulling on borosilicate glass tubes with a vertical puller. Based on the size and geometry of the resulting tip, further shape pipettes using a polisher. Note: The tip size can be evaluated visually and electrically. Visually, the outer diameter should be in the 1.4-5.6 &#956;m range. Electrically, an optimum size tip, when filled with extracellular solution, should produce resistances in the 12-24 M&#8486; range (see Figure 2B).</li>
	<li>Use a pipette solution, which for cell attached experiments represents the extracellular milieu that will produce maximal channel activity and high current amplitudes. Note: For NMDA receptors, this corresponds to a solution containing saturating concentrations of agonists (glutamate and glycine), 1 mM EDTA, which chelates divalent cationic inhibitors and blockers, and has physiologic concentrations of sodium (150 mM) as the sole permeant ion (in mM: 1 glutamate, 0.1 glycine, 150 NaCl, 2.5 KCl, 1 EDTA, and 10 HEPBS, buffered at pH 8.0 with 1 N NaOH).</li>
</ol>
3. Cell-attached Patch-clamp Recording
<ol>
	<li>Open QuB data acquisition software (www.qub.buffalo.edu), and select the acquisition window under &#8216;layout&#8217;. Open a new QuB data file (QDF) by clicking &#8216;New Data&#8217; under the drop down menu entitled &#8216;File&#8217; and enter the following initial parameters: sampling rate, 40 kHz; A/D scaling, 3,000; output channel, 1; and A/D data size, 2. Adjust the amplitude scaling to 0.1 V/pA by right-clicking the data file, selecting properties, and clicking the &#8216;data&#8217; tab. Note: These parameters can be refined for each setup using a model-cell in the cell-attached mode. Additional instruction on data acquisition with QuB software and QDF properties are online at www.qub.buffalo.edu.</li>
	<li>Select a 35 mm dish containing ion channel expressing cells and replace growth medium with 2 ml PBS containing calcium and magnesium; mount the dish onto the microscope stage and focus on the cellular field using phase contrast microscopy to evaluate that cells are healthy and well attached to the dish in monolayer fashion (Figure 1A). Switch to fluorescence detection to verify that the transfection was successful (Figure 1B). Note: The range of fluorescence intensities can be used as a crude measure of protein expression.</li>
	<li>For single channel recording, set the amplifier output gain to x10, patch configuration to &#946; = 1, analog filter to 10 kHz, mode to voltage-clamp and applied voltage to +100 mV. With the voltage holding command in the OFF position, select the &#8216;seal test&#8217; button.</li>
	<li>Based on fluorescence and cell health, choose a cell to patch. Verify under phase contrast that the cell is well attached to the dish, has a large portion of the cell surface exposed for patching, and otherwise looks healthy. Note: Maintain phase contrast illumination to avoid bleaching the cell during the approach and patch-clamping.</li>
	<li>Fill a freshly polished pipette with just enough solution so it makes contact with the electrode and shake gently to dislodge air bubbles that may be trapped in the tip. Secure the recording pipette onto the amplifier headstage by tightening the sealing screw of the pipette holder and making sure that the silver wire is immersed in the pipette solution.</li>
	<li>Using a small (5 cc) plastic syringe, which is connected to the pipette holder by tubing, gently apply positive pressure to prevent impurities from entering and clogging the tip during the approach (Figure 2A).</li>
	<li>Using the micromanipulator, direct the pipette into the bath and position it directly over the cell selected for patching (Figure 1C), closing the electrical circuit. Take note of the oscilloscope, which should indicate a rectangular waveform corresponding to the amplifier&#8217;s seal test signal (Figure 2B). Note: Upon entry into the bath, the pipette resistance within the optimal range is 12-24 M&#8486;. For a 5 mV test signal, the measured current range corresponds to ~20-40 pA.</li>
	<li>While monitoring pipette position visually through the microscope and pipette resistance electrically on the oscilloscope, continue the approach in small increments until the pipette impinges gently on the cell, and the test signal decreases slightly to indicate increased resistance (Figure 2B).</li>
	<li>To form a seal, pick up the syringe and apply slight negative pressure through the lateral tubing by pulling the syringe plunger, which pulls the cellular membrane into the tip of the recording pipette and initiates the formation of a G&#937; resistance seal10. Take note of the test signal waveform on the oscilloscope, in which seal formation is indicated by its complete flattening, with only capacitive transients visible (Figure 2B). Note: If signal does not flatten completely or the baseline becomes noisy, this indicates a weak seal with substantial current leak around the seal. This will prevent resolving one channel currents. If this is the case, withdraw the pipette from the cell and away from the bath, remove it from the head stage and discard it. Repeat the process with a freshly polished pipette until obtaining a seal in the adequate range (&#8805; 1 G&#937;).</li>
	<li>On the amplifier, switch the external command toggle from &#8216;seal test&#8217; to &#8216;off&#8217;; switch the voltage holding command to positive (which was set previously to &#8216;off&#8217;); and increase the gain to x100. Observe the oscilloscope for channel activity, which, if present, will be displayed as square upward deflections from the previously flat baseline (Figure 2C).</li>
	<li>If channel activity is displayed on the oscilloscope, acquire data into the previously opened digital file in QuB, by pressing the &#8216;play&#8217; button followed by the &#8216;record&#8217; button. To stop acquiring data, press the stop button in QuB, and save the QDF file to an easily retrievable location on the computer hard drive by selecting &#8216;Save Data As&#8230;&#8217; in the drop down menu entitled &#8216;File&#8217;.</li>
</ol>
4. Data Preprocessing and Idealization
Note: Important information can be extracted from single channel recordings by statistical analyses which assigns each data point to an appropriate conductance class (in the simplest case, closed or open). This process is referred to as data idealization and a brief description of data idealization with the segmental k means (SKM) method11 in QuB is described below.
<ol>
	<li>Open the data file in QuB, and view the current traces in the &#8216;Pre&#8217; interface under &#8216;layout&#8217;. Display the recorded file unfiltered (remove digital filter) by unchecking the box labeled &#8216;Fc.&#8217;</li>
	<li>Visually scan the record to spot irregularities and artifacts (Figure 4A). Correct brief current spikes that occur within the trace (Figure 4A) by selecting an adjacent clean region of the same conductance class, highlighting this region, right-clicking and selecting &#8216;set erase buffer.&#8217; Zoom in on the spike until individual sample points are visible, select the region to be replaced and erase by highlighting only this region and then right-click &#8216;erase&#8217;.</li>
	<li>Define the zero current baseline for the entire record by selecting an early portion of the record where the baseline is stable, highlight, right-click and select &#8216;set baseline&#8217;. Verify that the guideline which appears accurately represents the baseline level (purple in Figure 4B)</li>
	<li>Identify points in the record where the raw data baseline deviates visibly from the set baseline. Correct this by selecting a small section of baseline within the deviating region, right-click and select the &#8216;add a baseline node&#8217; command.</li>
	<li>Identify regions of the recording containing excess noise or artifacts that cannot be easily corrected (Figure 4C). Highlight the region to be discarded, right-click, and select &#8216;delete.&#8217; Note: In this case kinetic information will be lost: the processed record will be shorter and the points occurring after the splice point will appear to be continuous with the pre-noise region.</li>
	<li>To idealize records using the SKM algorithm11, highlight a portion of the trace containing both open and closed events and enter the &#8216;Mod&#8217; interface under &#8216;Layout.&#8217; In the &#8220;model&#8221; panel, highlight a clean portion of the trace that is representative of the baseline throughout the file, right-click the black square (closed state) and select &#8220;grab&#8221;. Do the same for channel openings by highlighting the open conductance, right click the red square in the &#8220;model&#8221; panel and select &#8220;grab&#8221;.</li>
	<li>Perform the idealization for the entire record by selecting the &#8216;File&#8217; button under &#8216;Data Source.&#8217; Right-click on the &#8216;Idealize&#8217; tab underneath the &#8216;Modeling&#8217; section to verify that the desired parameters for analysis are correct, and then click &#8216;Run.&#8217; The result of the idealization, along with amplitude histogram for the entire file, is overlaid with the data to allow for visual inspection of the idealization (Figure 5). Note: Inadequate idealization may lead to the generation of &#8216;false events,&#8217; in which QuB detects channel openings and closures that do not actually occur. Within the recording resolution, which is set by the amplifier&#8217;s analog filter and the sampling rate, the resolution with which SKM detects open and closed events can be controlled by setting a dead time for the analyses. Optimal dead times must be selected by trial and error and will depend on sampling rate, however, a good rule of thumb is to select a dead time that is 2-3 samples12.</li>
	<li>To verify that the idealization accurately represents the raw data, manually scan the idealized trace. Upon identification of errors in the idealization, highlight the trace over the false event, right click and select &#8216;Join Idl.&#8217; Note: For additional options and a detailed explanation of various commands and functions in QuB, consult the QuB Manual online (www.qub.buffalo.edu).</li>
</ol>

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The authors of this manuscript declare that they have no competing financial interests.

In the ion channel field, an important area of research is dedicated to understanding the sequence of events that leads to channel opening or the channel&#8217;s gating mechanism. For most channels, this process is complex and involves several kinetic steps that cannot be deduced from a macroscopic multi-channel signal. In contrast, experiments can be designed where observing the sequence of open/closed events in single channel record can produce more detailed information about gating mechanisms. In the methods described...

Fast communication across biological membranes relies almost exclusively on oligomeric pore forming membrane proteins, commonly referred to as channels. These proteins differ widely in activation signals, gating mechanisms, and conductance properties. Channel proteins whose pores are selective to ions are classified as ion channels; their activation produces ionic currents across the membrane, and their responses can be recorded with high resolution in real time using electrophysiologic techniques. The activation signals span a broad array of chemical and physical inputs including concentration gradients, mechanical and electrical forces, and temperature; thus, further classifying ion channels into ligand gated, mechanosensitive, voltage gated, or heat sensitive types. In this article, protocols are described to record one-channel activity from a ligand gated channel, the NMDA receptor, and a mechanosensitive channel, PIEZO1, using the patch-clamp technique.&#160;
Patch-clamp electrophysiology is the first and most widely used experimental method sufficiently sensitive to permit the observation of single molecules1,2. In addition to this exquisite sensitivity, it has vastly expanded the biological preparations amenable to electrophysiologic recording and also has allowed the observation of ion channels in intact membranes. First, because both voltage clamping and current recording are accomplished with the same electrode, it can be used to record signals across small cells or membranes patches. The technique revealed that ion channels are not restricted to excitable membranes of frog muscles, eel electroplaques, or squid giant axons3,4, but rather that they represent ubiquitous fixtures of transmembrane signaling mechanisms and are intrinsic to all cellular membrane types of uni- or multicellular organisms, and also to intracellular membranes. Importantly, the capability to record transmembrane currents by simply attaching a glass pipette to an intact cell provided the unprecedented opportunity to record activity from ion channels in their native undisrupted membranes. Thus, the cell attached patch-clamp technique, which is described in this protocol, permits monitoring the activity of ion channels continuously for tens of min or longer in their native environment.
Under normal thermal fluctuations, all proteins, including ion channel proteins, undergo structural changes over a broad time scale, with the fastest and most frequent rearrangements represented most likely by side-chain movements and much slower, less frequent changes represented by the repositioning of entire domains or subunits, or in some cases by post translational modifications or protein-protein interactions5,6. Observing long periods of activity generated by one molecule can help to understand the functional dynamics of ion channels in intact physiological membranes and provides valuable information about the operational mechanism of the molecule observed.&#160;
In contrast to the growing understanding of the diversity of ion channels across cell types and developmental stages, knowledge about the molecular composition of ion channels in native membranes is still limited. All ion channels are multimeric proteins and the majority of native ion channels assemble from several types of subunits producing proteins of wide molecular diversity, which is often accompanied with diverse conductance and gating properties. For this reason, ion channels of defined molecular composition are studied upon expression in heterologous systems. In particular, HEK293 cells, which are a clonal line of immortalized human embryonic kidney cells7, gained widespread acceptance as the preferred system for heterologous expression of recombinant ion channels. Among the many advantages that elevated&#160; HEK293 cells as the choice system for ion channel electrophysiology are the ease and affordability of culturing and maintaining long-lived stable cultures, their ability to carry out post-translational folding, processing and trafficking of mammalian proteins, and in many cases, their low level or even absence of endogenous expression for the channel of interest7,8. Expressing recombinant ion channels and studying their functional properties in HEK293 cells continues to be a valuable approach to obtain information about structure-function properties of ion channels as well as the specific properties of ion channel isoforms and their roles in native tissue. The protocols described in this article can be applied equally well to recombinant ion channels expressed in HEK293 cells and to native ion channels.
In summary, the patch-clamp technique, through its unprecedented capacity to resolve signals from one molecule remains, to date, the most direct method for observing the behavior of single molecules. In its cell-attached mode, patch-clamp recording allows long observation periods which, when done for one molecule, can provide exceptional insight into the operation of ion channels. Below is presented a protocol for obtaining high resolution current recordings from cell attached patches containing one ion channel protein.

<table border="0" cellpadding="0" cellspacing="0" width="695">
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:189px;">Chemicals</td>
			<td style="width:173px;">Sigma</td>
			<td style="width:128px;">Various</td>
			<td style="width:205px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Borosillicate Glass</td>
			<td>Sutter</td>
			<td>BF-150-86-10</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Bright field inverted microscope</td>
			<td>Olympus</td>
			<td>1x51</td>
			<td>Nikon also has similar microscopes</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Fluroescent box</td>
			<td>X-cite</td>
			<td>Series 120</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Liquid Light Guide</td>
			<td>X-cite</td>
			<td>OEX-LG15</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Micromanipulator</td>
			<td>Sutter Instruments</td>
			<td>MP-225</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Oscilloscope</td>
			<td>Tektronix</td>
			<td>TDS1001</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Amplifier</td>
			<td>Molecular Devices</td>
			<td colspan="2">Axon Axopatch 200B&#160;</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Table</td>
			<td>TMC</td>
			<td align="right">63561</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">NIDAQ card</td>
			<td>National Instruments</td>
			<td>776844-01</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Puller</td>
			<td>Narishige</td>
			<td>PC-10</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Polisher</td>
			<td>Narishige</td>
			<td>Microforge MF-830</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Faraday Cage</td>
			<td>TMC</td>
			<td align="right">8133306</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">High Speed Pressure Clamp</td>
			<td>ALA Scientific Instruments</td>
			<td>ALA HSPC</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Pressue/Vaccuum Pump</td>
			<td>ALA Scientific Instruments</td>
			<td>ALA PV-PUMP</td>
			<td>For HSPC-1</td>
		</tr>
	</tbody>
</table>

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.

Recombinant NMDA Receptors
NMDA receptors bind and respond to the concomitant action of two co-agonists: glutamate and glycine. They assemble as heterotetramers of two glycine binding GluN1 subunits and two glutamate binding GluN2 subunits. GluN2 subunits are encoded by four genes (A-D) and of these the most widely transcribed forms in brain are GluN2A in adult and GluN2B in juvenile animals. Because of the diversity of NMDA receptor subtypes in native preparations, expressing...

Watch this Scientific Journal Video about One-channel Cell-attached Patch-clamp Recording at JoVE.com

One-channel Cell-attached Patch-clamp Recording

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 ...

one-channel-cell-attached-patch-clamp-recording

Research

JoVE Journal

Biology

24.2K visualizações.  University at Buffalo, SUNY.  Descrito aqui é um procedimento para a obtenção de longos trechos de gravação atual de um canal iônico com a técnica de patch-clamp anexado à célula. Este método permite observar, em tempo real, o padrão de open-close conformações dos canais que estão na base do sinal biológico. Estes dados informar sobre as propriedades dos canais nas membranas biológicas não perturbadas. 

Vídeo: One-channel Cell-attached Patch-clamp Recording

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 ...

Dissecting and Recording from The C. Elegans Neuromuscular Junction

Neurotransmission is the process by which neurons transfer information via chemical signals to their post-synaptic targets, on a rapid time scale. This complex process requires the coordinated activity of many pre- and post-synaptic proteins to ensure appropriate synaptic connectivity, conduction of electrical signals, targeting and priming of secretory vesicles, calcium sensing, vesicle fusion, localization and function of postsynaptic receptors and finally, recycling mechanisms. As neuroscientists it is our goal to elucidate which proteins function in each of these steps and understand their mechanisms of action. Electrophysiological recordings from synapses provide a quantifiable read out of the underlying electrical events that occur during synaptic transmission. By combining this technique with the powerful array of molecular and genetic tools available to manipulate synaptic proteins in C. elegans, we can analyze the resulting functional changes in synaptic transmission. 
The C. elegans NMJs formed between motor neurons and body wall muscles control locomotion, therefore, mutants with uncoordinated locomotory phenotypes (known as unc s) often perturb synaptic transmission at these synapses 1. Since unc mutants are maintained on a rich supply of a bacterial food source, they remain viable as long as they retain some pharyngeal pumping ability to ingest food. This, together with the fact that C. elegans exist as hermaphrodites, allows them to pass on mutant progeny without the need for elaborate mating behaviors. These attributes, coupled with our recent ability to record from the worms NMJs 2,3,7 make this an excellent model organism in which to address precisely how unc mutants impact neurotransmission. 
The dissection method involves immobilizing adult worms using a cyanoacrylic glue in order to make an incision in the worm cuticle exposing the NMJs. Since C. elegans adults are only 1 mm in length the dissection is performed with the use of a dissecting microscope and requires excellent hand-eye coordination. NMJ recordings are made by whole-cell voltage clamping individual body wall muscle cells and neurotransmitter release can be evoked using a variety of stimulation protocols including electrical stimulation, light-activated channel-rhodopsin-mediated depolarization 4 and hyperosmotic saline, all of which will be briefly described.

Application of electrophysiology to accessible synapses provides a quantifiable measure of synaptic activity, useful in analyzing synaptic mutants. This article describes a dissection method used to expose the neuromuscular junctions (NMJ) of Caenorhabditis elegans (C. elegans) and briefly discusses some of the uses to which this preparation can be applied.

Neurotransmission is the process by which neurons transfer information via chemical signals to their post-synaptic targets, on a rapid time scale.   This ...

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

Ambulatory ECG Recording in Mice

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although the surface ECG is utilized for short-term measurements of waveform intervals, it is not practical for long-term studies of heart rate variability or the capture of rare episodes of arrhythmias. Implantable ECG telemeters offer the advantages of simple surgical implantation, long-term recording of electrograms in ambulatory mice, and scalability with simultaneous recordings of multiple animals. Here, we present a step-by-step guide to the implantation of telemeters for ambulatory ECG recording in mice. Careful adherence to aseptic technique is required for favorable survival results with the possibility of implantation and recording from weeks to months. Thus, implantable ECG telemetry is a valuable tool for detection of critical information on cardiac electrophysiology in ambulatory animal models such as the mouse. 

Telemetric ECG has emerged as an essential tool in evaluating animal models for cardiac arrhythmias and sudden cardiac death. Here, we present a stepwise guide to telemetric ECG recordings for application in long-term ambulatory ECG monitoring in mice.

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although ...

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 ...

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 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 ...