We thank J. Garcia-Celma, I Smirnova and R. Kaback for contributions to the LacY measurements and E. Bamberg for support and helpful discussions.

1. Setting up an Apparatus for SSM-based Electrophysiology
Details are given in two of our technical publications, which also contain schematic drawings and photographs of our cuvettes and set up 1, 2. Different solution exchange configurations and flow protocols are also discussed in our method paper 2.
In the following we add a few recent improvements and technical details which are of direct relevance for the video presentation.
Valve control and data acquisition
The interface box contains a commercial USB digital output / analog input interface (NI USB 6009 National Instruments) and the valve drivers. It controls the solution flow and is responsible for data acquisition. Valves are normally driven with 12 V. However, for fast switching the valves can be driven with a voltage of up to 18 V. In the video we use a 12 V power supply. 
The valve driver circuit board can operate four valves. It was manufactured by the workshop of the Max Planck Institute of Biophysics. Valves can be controlled by computer or manually via switches at the front panel. The latter is convenient for flushing- and cleaning-procedures. During measurement, the interface box is computer-controlled using a valve control and data acquisition software (SURFE2R software, IonGate Biosciences).
2. Preparations
In this section the different protocols for the preparation of a SSM-based electrophysiology experiment are mentioned.
2.1 Making the Lipid solution for formation of the SSM
<ol>
 <li>Mix 25 &mu;l Octadecylamine (5 mg/ml in Chloroform) and 375 &mu;l Diphytanoylphosphatidylcholine (20 mg/ml in Chloroform) in a glas vial. </li>
 <li>Using a rotary evaporator and continuous nitrogen gas flow, evaporate the chloroform for about 30 min.</li>
 <li>Remove the lipid from the walls of the glass vial by shaking with 500 &mu;l of n-decane. The final lipid concentration is 15 mg/ml with 1:60 (w/w) octadecylamine.</li>
 <li>Transfer the solution to the glass storage vial. Store the lipid solution at -20 &deg;C. </li>
</ol>
2.2 Chlorination of the reference electrode
The reference electrodes have to be chlorinated in regular intervals due to abrasion. 
<ol>
 <li>Remove the rest of the old silverchloride layer using fine sandpaper before the chlorination process.
 </li>
 <li>Place the silver wire together with a platinum electrode into a 1 M hydrochloric acid solution and chlorinate for 15 min at 0.5 mA. After completing chlorination, the silver wire changes its color to a homogenous dark grey. </li>
</ol>
2.3 Preparation of the polyacrylamide gel bridge
<ol>
 <li>Prepare a solution containing 100 mM KPi at pH=7, 100 mM KCl and 6% Acrylamid. 
 </li>
 <li>Add 0.3% APS (10% Stock) and 0.6% TEMED and shortly mix the solution with a pipette. </li>
 <li>Immediately after mixing the solution, use a pipette to inject 30 &mu;l of the mixture into the empty gel bridge container. It is important to avoid air bubbles during the injection.
 </li>
 <li>During an incubation time of 20 min the gel polymerizes. </li>
 <li>After polymerization the gel bridge is stored in solution (100 mM KPi pH 7, 100 mM KCl). To prolong the lifespan of the gel bridge the solution is stored at 4 &deg;C. </li>
</ol>
2.4 Preparation of the measuring solutions
Due to the strong interaction of the solutes with the SSM, electrical artifacts are generated when solutions of different composition are exchanged. Therefore, the preparation of the solutions is a critical step. Take the following precautions during the solution preparation process to minimize solution exchange artifacts.
<ol>
 <li>Make non-activating and activating solutions from one batch, adjust the pH and ionic strength.
 </li>
 <li>High salt background helps to reduce solution exchange artifacts. </li>
 <li>Divide the batch solution into two volumes. </li>
 <li>Add the activating compound to the activating solution. Use a compensatory compound in the non-activating solution to keep the osmolarity and ionic strength of both solutions as similar as possible. </li>
 <li>If the solutions are stored at 4 &deg;C, make sure that all solutions reached room temperature before starting a measurement, because even small differences in temperature can produce artifacts. </li>
</ol>
3. A SSM-based Electrophysiology Experiment
Here we present a typical protocol for a SSM-based electrophysiological experiment. 
3.1 Preparing the SSM Setup
While the SSM Setup is not in use the tubes are filled with a 30% ethanol/water solution to avoid bacterial growth. 
<ol>
 <li>Wash the system with pure water to remove the ethanol from the tubing before mounting the cuvette. Replace the ethanol containers with water containers. Using high pressure (0.6 to 1.0 bar) clean the system with 20-30 ml of water. </li>
 <li>Repeat the cleaning procedure using a 100 mM KPi buffer at pH 7.6 or non-activating solution.</li>
 <li>Make sure that no air bubbles remain in the fluid system. </li>
</ol>
3.2 Mounting the cuvette
<ol>
 <li>Attach an o-ring to the reference electrode</li>
 <li>Take the gel bridge from the storage solution and connect the reference electrode. </li>
 <li>Prefill the outlet connector with non-activating buffer, insert an o-ring and connect the gel bridge to complete the reference electrode assembly. Take special care that no air bubbles are at the junction of the gel bridge and the outlet solution flow pathway. </li>
 <li>Preassemble the main part of the cuvette by adding the spring contact pin (connection to the amplifier), the inlet tube (connection to the terminal valve), the o-ring (sealing for the SSM) and the screws.</li>
 <li>Using a tweezers, take the sensor chip out of the storing solution (10 mM octadecanethiol in ethanol). </li>
 <li>Using a pipette, wash off the remaining solution with approx. 5 ml of pure ethanol. </li>
 <li>Dry the electrode under nitrogen gas. </li>
 <li>Place the sensor chip accurately on the base part of the cuvette so that the inlet bore of the main part of the cuvette is directed to the circular active area of the sensor. </li>
 <li>Add 1 &mu;l of the lipid solution to the active area of the sensor chip. Make sure, that the gold layer is fully covered by the lipid. </li>
 <li>Immediately after adding the lipid solution, close the cuvette by adding the preassembled main part.</li>
 <li>Before mounting the cuvette into the Faraday cage, be sure that all surfaces are completely dry.</li>
 <li>Connect the amplifier to the spring contact pin and the inlet tube to the terminal valve. Then fix the cuvette with the screws inside the faraday cage.</li>
 <li>Screw the outlet connector to the top of the cuvette. Finally connect the outlet tube to the outlet connector and the voltage generator to the reference electrode.</li>
 <li>Immediately after mounting the cuvette wash the system with buffer at 0.6 bar. This leads to spontaneous SSM formation. </li>
</ol>
3.3 Measuring membrane parameters
To check the quality of the SSM, capacitance and conductance are measured using the function generator. 
<ol>
 <li>Using the function generator, apply 100 mV DC voltage to measure the conductance.</li>
 <li>Calculate the conductance: The current decay shows the charging of the membrane capacitor. After the capacitor is fully charged the measured current yields the membrane conductance using Ohm's law. For the sake of simplicity we use the current 1 sec after the voltage is applied to calculate the conductance G=I/U. </li>
 <li>Apply a triangular AC voltage of 50 mV peak to peak amplitude and a frequency of 0.5 Hz to measure the capacitance.</li>
 <li>Calculate the capacitance: The amplitude of the resulting square wave current represents the charging currents of the capacitor. The capacitance C is equal to the transferred charge Q=I&Delta;t divided by &Delta;U=100 mV. (&Delta;U is twice the applied voltage of 50 mV because I is the difference current positive minus negative slope of the triangular voltage.)</li>
 <li>Monitor the electrical parameters of the membrane repeatedly every 10 min for approx. 30 to 40 min, until constant values are reached. Wash in between with KPi buffer at high pressure in the range of about 0.6 to 1 bar. </li>
 <li>Estimate the quality of the SSM: Optimal parameters are in the range of 0.1 to 0.2 nS and 2 to 3.5 nF. A high conductance above 0.8 nS and low capacitance below 1 nF indicates that the SSM is not correctly formed. A high capacitance above 4 nF may imply that the active zone is not covered completely by the SSM. </li>
 <li>If the parameters are not in the optimal range, the membrane should be discarded and another SSM prepared, using a freshly prepared electrode chip. </li>
</ol>
3.4 Checking for solution exchange artifacts
After the membrane parameters have been checked, the measuring buffers should be tested for solution exchange artifacts. 
<ol>
 <li>Insert the respective solution containers to the fluid system.
 </li>
 <li>Remove air bubbles from the fluid system, using the manual valves.</li>
 <li>Using the data acquisition software chose the flow protocol you want to use in your experiment. </li>
 <li>Adjust the pressure to 0.6 bar and start the measurement. </li>
 <li>Apart from the mechanical valve switching artifacts, ideally no artifact currents should be measured or they should be much smaller than the expected protein transport signals. If solution exchange artifacts are observed, try to optimize the buffer compositions and/or the preparation procedure before continuing the experiment. </li>
</ol>
3.5 Adding the protein sample
Usually proteoliposomes or membrane fragments are stored frozen at -80 &deg;C. For one measurement an aliquot of approximately 30 &mu;l is needed. 
<ol>
 <li>Rinse the SSM with non-activating solution.</li>
 <li>Thaw the protein sample on ice. </li>
 <li>Three 10-sec sonication cycles are alternated with 10-sec cooling intervals on ice. Proteoliposomes are sonicated using a bath sonicator. For membrane fragments a tip sonicator (50W, 30 kHz, 1 mm sonotrode diameter, intensity 20%, cycle 0.5) is used. After the last sonication step do not put the sample back on ice, but inject it immediately into the cuvette.</li>
 <li>Unscrew the outlet connector together with the reference electrode assembly. </li>
 <li>Using a pipette, aspirate 30 &mu;l of the protein sample and mount the pipette tip to the outlet bore. 
 </li>
 <li>Open the manual valve in the non-activating pathway to the waste container and inject the proteoliposomes into the cuvette volume. Be careful not to inject air into the cuvette volume with the sensor. </li>
 <li>Before removing the pipette tip, close the manual valve to prevent further solution flow. </li>
 <li>Allow the proteoliposomes to adsorb to the SSM for an incubation time of 1 to 2 hr. It is also possible to incubate overnight. </li>
</ol>
3.6 General Procedure of an SSM-based experiment
<ol>
 <li>Warm up the measuring buffers in time, if stored at 4 &deg;C. All measuring buffers have to be at room temperature before starting the measurement to avoid artifacts. </li>
 <li>When changing solutions, first clean the tubes inside the solution containers with a non-fuzzing tissue, then insert the new bottles.</li>
 <li>Before starting the measurement, use the manual valves to remove air bubbles, which could have evolved from the changing procedure.</li>
 <li>Adjust the pressure just before starting a measurement. We routinely use a pressure of 0.6 bar which at our specific valve and tube configuration yields a flow rate of approximately 1.0 ml/sec. </li>
 <li>The first few measurements can be carried out straight one after another and should be rejected until the peak current remains constant. If the solutions differ in pH an incubation time of at least 3 min is needed to adjust the inner pH of the proteoliposomes before starting the measurement. </li>
 <li>Now the set up is ready for measurement. To reduce the noise at least 3 measurements should be made and averaged. </li>
</ol>
3.7 Control Measurements
A signal rundown during a series of measurements could occur due to protein degradation or a loss of adsorption. Each SSM experiment should include a rundown control. This is done by repeated measurements with identical solutions. The minimal rundown control means one measurement at the beginning and one at the end of the experiment, using the same solution. 
We recommend to do the rundown control under conditions where you expect the highest peak amplitude in the experiment. This makes it easier to quantify the rundown of the signal.
The signal rundown can be quantified by a comparison of the peak amplitudes of the two rundown controls. The resulting percentage value can be used to correct the measured signals by assuming a linear time dependence of the rundown. 
3.8 Artifact controls
If possible try to validate your results by inhibiting the protein of interest after the experiment. Remaining signals are probably solution exchange artifacts. The signals then have to be corrected for the measured artifacts.
3.9 After the experiment
<ol>
 <li> Wash the system with 20-30 ml of pure water and 20-30 ml of 30% ethanol/water. The ethanolic solution avoids bacterial growth when the system is not in use.</li>
 <li> After this cleaning procedure, release the pressure from the system and turn off all instruments. </li>
 <li> Remove the cuvette from the Faraday cage and dismount it. All parts of the cuvette can be cleaned with water and pure ethanol, if necessary, by using a bath sonicator.</li>
 <li> Place the sensor chip in a glas beaker with pure ethanol. Sonicate the beaker for about 1 to 2 min using a bath sonicator. </li>
 <li> Dry the sensor chip under nitrogen gas.</li>
 <li> Store the sensor chip away from light in a 10 mM Octadecanthiol ethanolic solution. After an incubation time of about 30 min the sensor chip is ready for reuse.</li>
</ol>

<ol>
	<li>Seifert, K., Fendler, K., Bamberg, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Charge+transport+by+ion+translocating+membrane+proteins+on+solid+supported+membranes.">Charge transport by ion translocating membrane proteins on solid supported membranes.</a> Biophys. J. 64, 384-391 (1993).</li><li>Schulz, P., Garcia-Celma, J. J., Fendler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SSM-based+electrophysiology.">SSM-based electrophysiology.</a> Methods. 46, 97-103 (2008).</li><li>Garcia-Celma, J. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+activation+of+the+melibiose+permease+MelB+immobilized+on+a+solid-supported+membrane.">Rapid activation of the melibiose permease MelB immobilized on a solid-supported membrane.</a> Langmuir. 24, 8119-8126 (2008).</li><li>Mager, T., Rimon, A., Padan, E., Fendler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transport+mechanism+and+pH+regulation+of+the+Na+/H++antiporter+NhaA+from+Escherichia+coli:+an+electrophysiological+study.">Transport mechanism and pH regulation of the Na+/H+ antiporter NhaA from Escherichia coli: an electrophysiological study.</a> J. Biol. Chem. 286, 23570-23581 (2011).</li><li>Zhou, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Charge+translocation+during+cosubstrate+binding+in+the+Na+/proline+transporter+of+E.coli.">Charge translocation during cosubstrate binding in the Na+/proline transporter of E.coli.</a> J. Mol. Biol. 343, 931-942 (2004).</li><li>Ganea, C., Fendler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bacterial+transporters:+charge+translocation+and+mechanism.">Bacterial transporters: charge translocation and mechanism.</a> Biochim. Biophys. Acta. 1787, 706-713 (2009).</li><li>Schulz, P., Werner, J., Stauber, T., Henriksen, K., Fendler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+G215R+mutation+in+the+Cl-/H+-antiporter+ClC-7+found+in+ADO+II+osteopetrosis+does+not+abolish+function+but+causes+a+severe+trafficking+defect.">The G215R mutation in the Cl-/H+-antiporter ClC-7 found in ADO II osteopetrosis does not abolish function but causes a severe trafficking defect.</a> PLoS ONE. 5, e12585 (2010).</li><li>Schulz, P., Dueck, B., Mourot, A., Hatahet, L., Fendler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measuring+ion+channels+on+solid+supported+membranes.">Measuring ion channels on solid supported membranes.</a> Biophys. J. 97, 388-396 (2009).</li><li>Garcia-Celma, J. J., Smirnova, I. N., Kaback, H. R., Fendler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+characterization+of+LacY.">Electrophysiological characterization of LacY.</a> Proc. Natl. Acad. Sci. U.S.A. 106, 7373-7378 (2009).</li><li>Garcia-Celma, J. J., Ploch, J., Smirnova, I., Kaback, H. R., Fendler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Delineating+electrogenic+reactions+during+lactose/H++symport.">Delineating electrogenic reactions during lactose/H+ symport.</a> Biochemistry. 49, 6115-6121 (2010).</li></ol>

No conflicts of interest declared.

1. Advantages of SSM-based electrophysiology compared to conventional methods
SSM-based electrophysiology has proven itself as a valuable tool of the electrophysiological toolbox. It is especially useful in cases where convention electrophysiology, namely patch clamp and voltage clamp methods, cannot be applied: Apart from a few rare exceptions bacterial transporters can not be investigated using voltage clamp or patch clamp methods because of the small size of bacteria and because they are difficult to express in mammalian cells or oocytes. But also physiologically relevant mammalian transporters can be investigated. In this case SSM-based electrophysiology is attractive for transporters from intracellular membranes and for screening applications in drug discovery because of its robustness and its potential for automation.
SSM-basedUsing conventional electrophysiology, time resolved characterization of transporters is challenging. Since the turnover of transporters is low a 'giant patch' or a 'whole cell' configuration is required, which have an inherently low time resolution in a solution exchange experiment. The complication can be overcome using photolytic substrate release. However, only a limited number of substrates are suited for this approach. Here the rapid solution exchange at the SSM offers the unique opportunity to perform electrophysiological studies with a high time resolution using arbitrary substrates.
2. Limitations and critical steps
In contrast to patch clamp and voltage clamp techniques, SSM-based electrophysiology cannot be used to apply a potential. Transporter characterization is therefore restricted to transport modes which do not rely on a membrane potential.
In general, SSM-based electrophysiology has no limitations concerning the type of the (electrogenic) transporter. But voltage clamp or patch clamp methods can have advantages, if intracellular components like binding proteins are required for protein functionality.
Limitations can arise, if solution exchange creates large artifact currents. This happens when the substrate interacts strongly with the SSM like in the case of lipophilic compounds. Artifact controls can be used to correct the measured signals. Moreover high salt background in all measuring buffers can be used to reduce the artifacts. But in cases, where the size of the artifact is comparable to the protein signal, it is almost impossible to isolate the protein related signal from the artifact. Fortunately, high artifacts are unusual in an optimized solution exchange.
There are a few steps which are critical for the successful realization of a SSM-based electrophysiology experiment. The preparation of the protein sample is the most important part. If proteoliposomes are used, be sure the reconstitution process yields a clean, reproducible sample of a sufficient LPR and the transporter is oriented in the correct way. The LPR can be checked by freeze fracture electron microscopy and orientation by an ELISA experiment if antibodies are available.
Only use a SSM which shows optimal parameters for incubating the protein sample. The injection of the protein is another critical step. Sonication is essential and air bubbles should be avoided during the injection. After sample incubation the measurements itself are critical, because air bubbles will remove the adsorbed protein sample from the sensor chip. Therefore always remove air bubbles after changing solutions. Nevertheless a signal rundown can occur. To correct a possible signal rundown, it is essential to accomplish rundown controls during the experiment.
3. Specialized Systems
The SSM-Setup can be modified according to its application. Moreover there are completely different, highly specialized setups available.
There is the possibility to measure protein signals under asymmetrical conditions, e.g. under a pH gradient. To establish asymmetrical buffer compositions inside and outside the proteoliposomes a third solution, the resting solution, has to be introduced and this requires a double exchange configuration. Here an additional three-way valve switching between non-activating and resting solutions is required.
To increase the time resolution of the system we developed an alternative flow pathway lacking the terminal valve, but using a different type of cuvette. Here the junction of activating and non-activating solution is located inside the cuvette, 3 mm in front of the SSM. This setup is well suited for kinetic analysis of fast transport processes. It could be shown that a time resolution as low as 2 msec is possible.
Commercial fully automated systems are available aiming at a significantly higher throughput for drug screening. A movable unit collects solutions and injects them on the sensor surface in 96-well plates in a standard microtiter plate format.

Here we demonstrate a new electrophysiological approach based on a solid supported membrane (SSM) for characterization of electrogenic membrane proteins.
The solid support consists of a thin gold layer on a glass slide, the sensor chip. The hydrophilic gold surface is used to bind the thiol group of an alcanethiol reagent. Afterwards, selfassembly of a phosphatidylcholine monolyer completes the formation of the SSM.
To measure electrogenic reactions of membrane proteins, proteoliposomes or membrane fragments are adsorbed to the SSM (Figure 1). The protein containing membrane and the SSM then form a capacitively coupled membrane system. Therefore, charge translocation at the protein containing membrane can be detected by capacitive coupling via the SSM. This method yields only transient currents. The peak current represents the stationary transport activity. The time dependent transporter currents can be reconstructed by circuit analysis.
The sensor chip is mounted into a cuvette system (Figure 2). The cuvette has a cylindrical cuvette volume of 17 &mu;l (net volume with o-ring mounted). A spring contact pin creates the contact to the amplifier. An outlet connector is screwed to the top of the main part and carries the reference electrode, a chlorinated silver wire. 
The cuvette is mounted in a Faraday cage. It is connected to a fluid pathway , which is used to induce the transport activity of the membrane protein in response to a fast solution exchange (Figure 3). In the single solution exchange protocol two solutions, one non-activating and one activating solution, are required. The flow is controlled by pressurized air using a valve control software on a computer or manual switches on an interface box.

<table><tbody><tr><td colspan="4">&lt;strong&gt;Materials&lt;/strong&gt;</td></tr><tr><td>Waterbath Sonicator</td><td>Bandelin</td><td>RK 52 H</td><td></td></tr><tr><td>Tip Sonicator</td><td>Hielscher Ultrasonics GmbH</td><td>UP50H</td><td></td></tr><tr><td>2-way valve</td><td>NResearch, West Caldwell, USA</td><td>NR225T011</td><td></td></tr><tr><td>Terminal valve</td><td>NResearch, West Caldwell, USA</td><td>NR225T031</td><td></td></tr><tr><td>Manometer</td><td>Greisinger electronics</td><td>GDH 14 AN</td><td></td></tr><tr><td>Faraday cage</td><td>Max Planck Institute of Biophysics</td><td></td><td></td></tr><tr><td>Cuvette</td><td>Max Planck Institute of Biophysics</td><td></td><td></td></tr><tr><td>100 ml solution containers</td><td>Kartell</td><td>1623</td><td></td></tr><tr><td>O-rings</td><td>Seal Science Inc.</td><td></td><td></td></tr><tr><td>Oscilloscope</td><td>Tektronix</td><td>TDS 1002</td><td></td></tr><tr><td>Reference electrode</td><td>Max Planck Institute of Biophysics</td><td></td><td></td></tr><tr><td>Function generator</td><td>Max Planck Institute of Biophysics</td><td></td><td></td></tr><tr><td>Tubings</td><td>SAINT-GOBAIN Performance Plastics</td><td>AAC00006</td><td></td></tr><tr><td>Sensor chip</td><td>Fraunhofer Institut f&amp;#252;r Schicht und Oberfl&amp;#228;chentechnik</td><td></td><td></td></tr><tr><td>Interface box</td><td>Max Planck Institute of Biophysics</td><td></td><td></td></tr><tr><td>Amplifier</td><td>Keithley</td><td>427</td><td></td></tr><tr><td>Manual cog</td><td>Vygon GmbH</td><td>876</td><td></td></tr><tr><td>USB analog-to-digital converter</td><td>National Instruments</td><td>6009</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Regeants&lt;/strong&gt;</td></tr><tr><td>1,2&amp;#8209;Diphytanoyl-sn-glycero-3-Phosphatidylcholine</td><td>Avanti Polar Lipids, Inc.</td><td>850356</td><td></td></tr><tr><td>Acrylamide/Bis-acrylamide</td><td>Sigma Aldrich</td><td>A3574</td><td></td></tr><tr><td>Ammonium persulfate</td><td>Sigma Aldrich</td><td>A3678</td><td></td></tr><tr><td>D-(+)-Glucose </td><td>Sigma Aldrich</td><td>G8270</td><td></td></tr><tr><td>Dithiothreitol </td><td>Sigma Aldrich</td><td>43819</td><td></td></tr><tr><td>Ethanol absolut</td><td>VWR AnalaR NORMAPUR</td><td>603-002-00-5</td><td></td></tr><tr><td>Natural E. coli lipids polar extract</td><td>Avanti Polar Lipids, Inc.</td><td>100600</td><td>for LacY reconstitution</td></tr><tr><td>n-Decane</td><td>Sigma Aldrich</td><td>D901</td><td></td></tr><tr><td>Octadecanethiol</td><td>Sigma Aldrich</td><td>O1858</td><td></td></tr><tr><td>Octadecylamine</td><td>Sigma Aldrich</td><td>74750</td><td></td></tr><tr><td>Potassium chloride</td><td>Merck</td><td>1049360500</td><td></td></tr><tr><td>Potassium phosphate dibasic</td><td>Sigma Aldrich</td><td>P3786</td><td></td></tr><tr><td>Potassium phosphate monobasic</td><td>Sigma Aldrich</td><td>P9791</td><td></td></tr><tr><td>Tetramethylethylenediamine</td><td>BIO RAD</td><td>1610801</td><td></td></tr><tr><td>&amp;alpha;-Lactose monohydrate</td><td>Sigma Aldrich</td><td>L8783</td><td></td></tr></tbody></table>

introduction to solid supported membrane based electrophysiology

The electrophysiological method we present is based on a solid supported membrane (SSM) composed of an octadecanethiol layer chemisorbed on a gold coated sensor chip and a phosphatidylcholine monolayer on top. This assembly is mounted into a cuvette system containing the reference electrode, a chlorinated silver wire.
After adsorption of membrane fragments or proteoliposomes containing the membrane protein of interest, a fast solution exchange is used to induce the transport activity of the membrane protein. In the single solution exchange protocol two solutions, one non-activating and one activating solution, are needed. The flow is controlled by pressurized air and a valve and tubing system within a faraday cage.
The kinetics of the electrogenic transport activity is obtained via capacitive coupling between the SSM and the proteoliposomes or membrane fragments. The method, therefore, yields only transient currents. The peak current represents the stationary transport activity. The time dependent transporter currents can be reconstructed by circuit analysis.
This method is especially suited for prokaryotic transporters or eukaryotic transporters from intracellular membranes, which cannot be investigated by patch clamp or voltage clamp methods.

Up to now, SSM-based electrophysiology was used to characterize more than 20 transporters, mostly of prokaryotic origin, e.g. MelB 3, NhaA 4 and PutP 5 (summarized in 6). But also eukaryotic transporters from intracellular membranes, e.g. ClC7 7 as well as ion channels, e.g. the nicotinic acetylcholine receptor 8 have been investigated.
Here we present as an example measurements of the sugar/H+ cotransporter LacY from Escherichia coli using proteoliposomes with at least 85% right side out oriented LacY at a LPR of 5 9, 10. We focus on the main steps leading to a minimal kinetic model of this transport protein.
1. Electrophysiological characterization of wildtype LacY
The first step is a general characterization of the electrogenic reaction by measuring different pH values (Figure 4), substrate concentrations (Figure 5) and substrates. To validate the technique, KM and pK values given in literature were reproduced. 
The pH dependence of LacY shows two different electrogenic reactions. The capacitively coupled current increases between pH 5 and pH 8.5, but also changes its shape. Under alkaline conditions steady state transport is observed, while for acidic pH values only a rapid electrogenic reaction remains. To distinguish the steady state signals from the fast electrogenic reactions we used circuit analysis to reconstruct transporter currents (Figure 6). 
At pH 7 the monophasic signal becomes biphasic. This also indicates that there are two different electrogenic reactions. Estimated from the transferred charge (integral of the signals), the two reactions have about 6% and 94% of the total electrogenicity of the transport cycle.
2. Assignment of the electrogenic reactions
To assign the two electrogenic reactions to particular steps in the reaction cycle of LacY, the E325A LacY variant was measured (Figure 7). This variant only shows lactose exchange but is deficient for all active transport modes, because of the inhibition of proton release. The signal of E325A LacY represents a fast transient and is constant for all measured pH values. The shape and amplitude of the signal is similar to the signal of wildtype LacY under acidic conditions. In addition we observe a small negative phase after the declining peak current, which is characteristic for rapid transient currents measured in capacitively coupled systems. Because lactose binding and release is not electrogenic, the fast transient must correlate with an electrogenic conformational transition following lactose binding.
It is known that the proton release step is rate limiting for LacY turnover in the lactose concentration gradient driven transport mode. In this case we expect an increase of the transport rate with alkaline pH, because proton release is favored. This is the case for the steady state transport correlated signal of wildtype LacY. Moreover it could be shown by pH gradient measurement that only the inner pH influences the steady state transport of LacY (unpublished). Therefore the large electrogenic reaction could be assigned to the proton release step.
3. Measurements for kinetic analysis
After identification of the electrogenic reactions on and off transport rates were determined using a SSM Setup with a high time resolution of about 4.5 msec. This is only possible under conditions, when one reaction dominates the signal. In this case the rate constants can be derived from the transient currents under consideration of the time resolution of the signals using an iterative least-squares deconvolution algorithm (Figure 8). The determined rate constants as well as the proposed minimal kinetic model (Figure 9) finally were used for the simulation of the transporter kinetics. The simulated curves reproduce the SSM data which additionally proves the kinetic model. 
<img src="/files/ftp_upload/50230/50230fig1.jpg" alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/50230/50230fig1highres.jpg"/> 
Figure 1. Adsorption geometry of proteoliposomes on the SSM. The SSM is formed on a sensor chip, a structured gold coated glass slide. To measure electrogenic reactions of membrane proteins, proteoliposomes or membrane fragments are adsorbed to the SSM. The two membranes form a capacitively coupled system. This is why only transient currents are detected.
<img src="/files/ftp_upload/50230/50230fig2.jpg" alt="Figure 2" fo:content-width="5.5in" fo:src="/files/ftp_upload/50230/50230fig2highres.jpg"/> 
Figure 2. The SSM cuvette carries the sensor chip and the reference electrode. The sensor chip is sandwiched between the cuvette base and the cuvette head. The reference electrode is isolated from the flow pathway by a polyacrylamide gel salt bridge.
<img src="/files/ftp_upload/50230/50230fig3.jpg" alt="Figure 3" fo:content-width="4.5in" fo:src="/files/ftp_upload/50230/50230fig3highres.jpg"/> 
Figure 3. Fluid Pathway and Electrical Circuit of the SSM Setup. In the single solution exchange protocol only one non-activating (NA) and one activating (A) solution are required. The flow is controlled by two 2-way valves (V1) and one terminal valve (V2). The terminal valve switches between non-activating and activating solutions, directing one of the solutions to the cuvette and the other solution to a waste container (W). The sensor chip (SSM) is connected to the amplifier (I/U), while the reference electrode (AgCl) is connected to the function generator (F) in the external electrical circuit. A computer (PC) and an interface box are used for valve operation and data acquisition. 
<img src="/files/ftp_upload/50230/50230fig4.jpg" alt="Figure 4" fo:content-width="4in" fo:src="/files/ftp_upload/50230/50230fig4highres.jpg"/> 
Figure 4. Transient currents measured with wild-type LacY proteoliposomes after 100 mM lactose concentration jumps at different pH values. The non-activating solution contained 100 mM glucose while the activating solution contained 100 mM lactose. All solutions were prepared in 100 mM potassium phosphate buffer at the indicated pH with 1 mM DTT. For details of measurement conditions in this and the following figures please refer to 9 and 10.
<img src="/files/ftp_upload/50230/50230fig5.jpg" alt="Figure 5" fo:content-width="4in" fo:src="/files/ftp_upload/50230/50230fig5highres.jpg"/> 
Figure 5. Averaged normalized Peak currents measured with wild-type LacY proteoliposomes after lactose concentration jumps at different concentrations and pH values. The KM values are obtained from the hyperbolic fits.
<img src="/files/ftp_upload/50230/50230fig6.jpg" alt="Figure 6" fo:content-width="4.25in" fo:src="/files/ftp_upload/50230/50230fig6highres.jpg"/> 
Figure 6. Reconstruction of transporter currents. The transient currents (A) used to reconstruct the transporter currents (B) were measured with wild-type LacY proteoliposomes after 100 mM lactose concentration jumps at pH 8.5 (blue trace) and pH 5.2 (red trace). At pH 8.5 continuous turnover is observed while at pH 5.2 a fast electrogenic reaction occurs. This is clearly observed in the reconstructed current (B).
<img src="/files/ftp_upload/50230/50230fig7.jpg" alt="Figure 7" fo:content-width="4in" fo:src="/files/ftp_upload/50230/50230fig7highres.jpg"/> 
Figure 7. Transient current measured with E325A LacY proteoliposomes after a 100 mM lactose concentration jump at pH 5.2. This LacY variant is inhibited in the proton release step of the transport cycle and therefore transport-deficient. The small negative component observed is characteristic for rapid transient currents measured in capacitively coupled systems and is caused by the discharge of the membrane capacitance after a fast electrogenic reaction. Its time constant &pi;0 is determined by the capacitances and conductances of the system (Figure 1).
<img src="/files/ftp_upload/50230/50230fig8.jpg" alt="Figure 8" fo:content-width="4.25in" fo:src="/files/ftp_upload/50230/50230fig8highres.jpg"/> 
Figure 8. Transient currents measured with wild-type LacY proteoliposomes after different lactose concentration jumps at pH 5.2. Under these conditions no steady state transport is observed, but an electrogenic conformational transition upon lactose binding. The dashed line shows the transfer function representing the time resolution of the system. The inset shows the determination of the on and off rate constants from the observed rate constant kobs with a hyperbolic fit to the data. The observed rate constant was determined from the transient currents with an iterative least-squares deconvolution algorithm 10.
<img src="/files/ftp_upload/50230/50230fig9.jpg" alt="Figure 9" fo:content-width="4.25in" fo:src="/files/ftp_upload/50230/50230fig9highres.jpg"/> 
Figure 9. A kinetic model for the reaction cycle of LacY. Two electrogenic steps could be identified and characterized by SSM measurements. A strongly electrogenic reaction representing ~94% of the total charge displacement in the reaction cycle is only observed at neutral and basic pH values. It could be assigned to the proton release step (blue arrow). A weakly electrogenic step is observed at acidic pH when continuous turnover of wild-type LacY is inhibited. We assigned this reaction to an electrogenic conformational transition upon lactose binding, which represents 6% of the total charge displacement in the reaction cycle.

Watch this Scientific Journal Video about Introduction to Solid Supported Membrane Based Electrophysiology at JoVE.com

Introduction to Solid Supported Membrane Based Electrophysiology

Here we present an electrophysiological method based on solid supported membranes with focus on its applications for the characterization of electrogenic membrane transporters.

The electrophysiological method we present is based on a solid supported membrane (SSM) composed of an octadecanethiol layer chemisorbed on a gold coated ...

introduction-to-solid-supported-membrane-based-electrophysiology

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

15.0K Views.  Max Planck Institute of Biophysics. Here we present an electrophysiological method based on solid supported membranes with focus on its applications for the characterization of electrogenic membrane transporters.

Video: Introduction to Solid Supported Membrane Based Electrophysiology

Applying Microfluidics to Electrophysiology

Microfluidics can be integrated with standard electrophysiology techniques to allow new experimental modalities.  Specifically, the motivation for the microfluidic brain slice device is discussed including how the device docks to standard perfusion chambers and the technique of passive pumping which is used to deliver boluses of neuromodulators to the brain slice.  By simplifying the device design, we are able to achieve a practical solution to the current unmet electrophysiology need of applying multiple neuromodulators across multiple regions of the brain slice.  This is achieved by substituting the standard coverglass substrate of the perfusion chamber with a thin microfluidic device bonded to the coverglass substrate.  This was then attached to the perfusion chamber and small holes connect the open-well of the perfusion chamber to the microfluidic channels buried within the microfluidic substrate.  These microfluidic channels are interfaced with ports drilled into the edge of the perfusion chamber to access and deliver stimulants.  This project represents how the field of microfluidics is transitioning away from proof-of concept device demonstrations and into practical solutions for unmet experimental and clinical needs.

Microfluidics can be integrated with standard electrophysiology techniques to allow new experimental modalities.  Specifically, the motivation for the ...

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 ...

Measuring the Induced Membrane Voltage with Di-8-ANEPPS

Placement of a cell into an external electric field causes a local charge redistribution inside and outside of the cell in the vicinity of the cell membrane, resulting in a voltage across the membrane. This voltage, termed the induced membrane voltage (also induced transmembrane voltage, or induced transmembrane potential difference) and denoted by &Delta;&Phi;, exists only as long as the external field is present. If the resting voltage is present on the membrane, the induced voltage superimposes (adds) onto it. By using one of the potentiometric fluorescent dyes, such as di-8-ANEPPS, it is possible to observe the variations of &Delta;&Phi; on the cell membrane and to measure its value noninvasively. di-8-ANEPPS becomes strongly fluorescent when bound to the lipid bilayer of the cell membrane, with the change of the fluorescence intensity proportional to the change of &Delta;&Phi;. This video shows the protocol for measuring &Delta;&Phi; using di-8-ANEPPS and also demonstrates the influence of cell shape on the amplitude and spatial distribution of &Delta;&Phi;.

External electric field induces a voltage on the membrane of a cell, termed the induced membrane voltage (&Delta;&Phi;). By using the potentiometric dye di-8-ANEPPS, it is possible to measure the &Delta;&Phi; noninvasively. This video shows the protocol for measuring &Delta;&Phi; using di-8-ANEPPS.

Placement of a cell into an external electric field causes a local charge redistribution inside and outside of the cell in the vicinity of the cell ...

Real-time Electrophysiology: Using Closed-loop Protocols to Probe Neuronal Dynamics and Beyond

Experimental neuroscience is witnessing an increased interest in the development and application of novel and often complex, closed-loop protocols, where the stimulus applied depends in real-time on the response of the system. Recent applications range from the implementation of virtual reality systems for studying motor responses both in mice1 and in zebrafish2, to control of seizures following cortical stroke using optogenetics3. A key advantage of closed-loop techniques resides in the capability of probing higher dimensional properties that are not directly accessible or that depend on multiple variables, such as neuronal excitability4 and reliability, while at the same time maximizing the experimental throughput. In this contribution and in the context of cellular electrophysiology, we describe how to apply a variety of closed-loop protocols to the study of the response properties of pyramidal cortical neurons, recorded intracellularly with the patch clamp technique in acute brain slices from the somatosensory cortex of juvenile rats. As no commercially available or open source software provides all the features required for efficiently performing the experiments described here, a new software toolbox called LCG5 was developed, whose modular structure maximizes reuse of computer code and facilitates the implementation of novel experimental paradigms. Stimulation waveforms are specified using a compact meta-description and full experimental protocols are described in text-based configuration files. Additionally, LCG has a command-line interface that is suited for repetition of trials and automation of experimental protocols.

Closed-loop protocols are becoming increasingly widespread in modern day electrophysiology. We present a simple, versatile and inexpensive way to perform complex electrophysiological protocols in cortical pyramidal neurons in vitro, using a desktop computer and a digital acquisition board.

Experimental neuroscience is witnessing an increased interest in the development and application of novel and often complex, closed-loop protocols, where ...

Neuroscience

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 ...

Modeling Biological Membranes with Circuit Boards and Measuring Electrical Signals in Axons: Student Laboratory Exercises

This is a demonstration of how electrical models can be used to characterize biological membranes. This exercise also introduces biophysical terminology used in electrophysiology. The same equipment is used in the membrane model as on live preparations. Some properties of an isolated nerve cord are investigated: nerve action potentials, recruitment of neurons, and responsiveness of the nerve cord to environmental factors.

This is a demonstration of how biological membranes can be understood using electrical models. We also demonstrate procedures for recording action potentials from the ventral nerve cord of the crayfish for student orientated laboratories.

This is a demonstration of how electrical models can be used to characterize biological membranes. This exercise also introduces biophysical terminology ...

Murine Short Axis Ventricular Heart Slices for Electrophysiological Studies

Murine cardiomyocytes have been extensively used for in vitro studies of cardiac physiology and new therapeutic strategies. However, multicellular preparations of dissociated cardiomyocytes are not representative of the complex in vivo structure of cardiomyocytes, non-myocytes and extracellular matrix, which influences both mechanical and electrophysiological properties of the heart. Here we describe a technique to prepare viable ventricular slices of adult mouse hearts with a preserved in vivo like tissue structure, and demonstrate their suitability for electrophysiological recordings. After excision of the heart, ventricles are separated from the atria, perfused with Ca2+-free solution containing 2,3-butanedione monoxime and embedded in a 4% low-melt agarose block. The block is placed on a microtome with a vibrating blade, and tissue slices with a thickness of 150-400 &#181;m are prepared keeping the vibration frequency of the blade at 60-70 Hz and moving the blade forward as slowly as possible. Thickness of the slices depends on the further application. Slices are stored in ice cold Tyrode&#39;s solution with 0.9 mM Ca2+ and 2,3-butanedione monoxime (BDM) for 30 min. Afterwards, slices are transferred to 37 &#176;C DMEM for 30 min to wash out the BDM. Slices can be used for electrophysiological studies with sharp electrodes or micro electrode arrays, for force measurements to analyze contractile function or to investigate the interaction of transplanted stem cell-derived cardiomyocytes and host tissue. For sharp electrode recordings, a slice is placed into a 3 cm cell culture dish on the heating plate of an inverted microscope. The slice is stimulated with a unipolar electrode, and intracellular action potentials of cardiomyocytes within the slice are recorded with a sharp glass electrode.

Here, we describe the preparation of viable ventricular slices from adult mice and their use for sharp electrode action potential recordings. These multicellular preparations provide a preserved in vivo like tissue structure, which makes them a valuable model for electrophysiological and pharmacological studies in vitro.

Murine cardiomyocytes have been extensively used for in vitro studies of cardiac physiology and new therapeutic strategies. However, multicellular ...

Medicine

Electrophysiological Recordings of Single-cell Ion Currents Under Well-defined Shear Stress

Fluid shear stress is well known to play a major role in endothelial function. In most vascular beds, elevated shear stress from acute increases in blood flow triggers a signaling cascade resulting in vasodilation thereby alleviating mechanical stress on the vascular wall. The pattern of shear stress is also well known to be a critical factor in the development of atherosclerosis with laminar shear stress being atheroprotective and disturbed shear stress being pro-atherogenic. While we have a detailed understanding of the various intermediate cell signaling pathways, the receptors that first translate the mechanical stimulus into chemical mediators are not completely understood. Mechanosensitive ion channels are critical to the response to shear and regulate shear-induced cell signaling thereby controlling the production of vasoactive mediators. These channels are among the earliest activated signaling components to shear and have been linked to shear-induced vasodilation through promoting nitric oxide production (e.g., inwardly rectifying K+ [Kir] and transient receptor potential [TRP] channels) and endothelium hyperpolarizing factor (e.g., Kir and calcium-activated K+ [KCa] channels) and shear-induced vasoconstriction through an undetermined mechanism that involves piezo channels. Understanding the biophysical mechanism by which these channels are activated by shear forces (i.e., directly or through a primary mechano-receptor) could provide potential new targets to resolve the pathophysiology associated with endothelial dysfunction and atherogenesis. It is still a major challenge to record flow-induced activation of ion channels in real time using electrophysiology. The standard methods to expose cells to well-defined shear stress, such as the cone and plate rheometer and closed parallel plate flow chamber do not allow real time study of ion channel activation. The goal of this protocol is to describe a modified parallel plate flow chamber that allows real time electrophysiological recording of mechanosensitive ion channels under well-defined shear stress.

The goal of this protocol is to describe a modified parallel plate flow chamber for use in investigating real time activation of mechanosensitive ion channels by shear stress.

Fluid shear stress is well known to play a major role in endothelial function. In most vascular beds, elevated shear stress from acute increases in blood ...

Immunology and Infection

Lipid Bilayer Experiments with Contact Bubble Bilayers for Patch-Clampers

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

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

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

Patch Clamp Electrophysiology

Neuron cell membranes are populated with ion channels that control the movement of charge into and out of the cell, thereby regulating neuron firing. One extremely useful technique for investigating the biophysical properties of these channels is called patch clamp recording. In this method, neuroscientists place a polished glass micropipette against a cell and apply suction to form a high resistance seal. This process isolates a small &#8220;patch&#8221; of membrane that contains one or more ion channels. Using an electrode within the micropipette, researchers can &#8220;clamp&#8221; or control the electrical properties of the membrane, which is important for analysis of channel activity. The electrode also allows for changes in the voltage across the membrane, or the flow of ions through the membrane, to be recorded. This video begins with an overview of the principles behind patch clamp electrophysiology, an introduction to the necessary equipment, and descriptions of the various patch configurations, including whole cell, cell-attached, perforated, inside-out, and outside-out patches. Next, the key steps of a typical whole-cell patch clamp experiment are outlined, in which a current-voltage (IV) curve is generated. Finally, applications of patch clamp recording are provided to demonstrate how the biophysical properties of ion channels, cell excitability, and neuroactive compounds are evaluated in neurophysiology labs today.

Patch Clamp Electrophysiology: Principle &amp; Applications

Neuron cell membranes are populated with ion channels that control the movement of charge into and out of the cell, thereby regulating neuron firing. One ...