Our work was made possible by financial support from McKnight Endowment Fund for Neuroscience (PAS), the National Institute on Drug Abuse (R01 DA019022; PAS) and a pre-doctoral NIH NRSA fellowship to P.A. (F31AA017042) from the National Institute on Alcohol Abuse and Alcoholism. The GIRK2- L257W mutant was provided by Dr. Prafulla Aryal. Fluorescence microscope and sponsorship kindly provided by Leica Instruments.

1. Site Directed Mutagenesis
We use mammalian expression plasmid pcDNA3.1 encoding a cDNA for GIRK2 and introduce a point mutation using Stratagene (Agilent Technologies) Quikchange XL II site-directed mutagenesis kit, according to the manufacturer's instructions.
Primers sequence can be easily generated using online primer design program available at: <a href="https://www.genomics.agilent.com/CollectionSubpage.aspx?PageType=Tool&amp;SubPageType=ToolQCPD&amp;PageID=15">https://www.genomics.agilent.com/CollectionSubpage.aspx?PageType=Tool&amp;SubPageType=ToolQCPD&amp;PageID=15</a> Note: We use standard Eppendorf 1.5 mL tubes in place of BD 14 mL tubes described in the protocol. However, the use of XL10-Gold ultracompetent E. coli pretreated with &beta;-mercaptoethanol, 30s heat shock time and NZY+ (or NZYM+) media are critical for the successful mutagenesis. If possible, engineering an additional silent mutation that creates a new restriction site can be convenient for screening mutant colonies below.
2. Miniprep, DNA Sequencing and Maxiprep
2.1 Qiagen miniprep and sequencing.
Individual bacterial colonies are picked and grown in LB broth with ampicillin (AMP). We follow the manufacturer's instructions for purifying the cDNA. A simple restriction digest can be used confirm the successful introduction of the mutation, if a restriction site is incorporated in Step 1 above. Automated DNA sequencing is done off-site, and is essential to confirm the presence of the mutation in the cDNA, as well as assess whether there are any unwanted mutations. This step is very important. We sequence the entire GIRK coding sequence using forward T7 promoter and reverse BGH sequencing primers that flank multiple cloning site of the pcDNA3.1 plasmid. Note: It is important to carefully inspect the sequencing electropherogram to assure sufficient quality of the sequencing data. High quality electropherogram has sharp, non-overlapping peaks with little background noise.
2.2 Qiagen maxiprep.
We follow the manufacturer's instructions. Note: This step is included in our experiments to improve the purity of the cDNA, which we find is important for efficient transfection.
3. Culture and Transfection of HEK293T Cells
All of these steps should be done in a sterile tissue culture hood.
3.1 Cell culture
HEK293T cells are convenient to use because they are easy to culture (37&deg;C., 5%CO2 humidified incubator), have fast replication time, high transfection efficiency and are amenable to whole-cell patch-clamp electrophysiology. HEK293T cells express SV40 large T antigen that ensures episomal replication of pcDNA3.1 plasmid.
HEK293T cells are cultured in low-glucose DMEM media supplemented with L-glutamine and 10 % Fetal Bovine Serum (FBS). For passaging cells, add 0.25% trypsin/EDTA to confluent (90-95%) HEK293T cells, wait until cells detach from the tissue dish, then stop the enzyme activity by addition of fresh media containing FBS/. Plate ~1 X 105 cells/well into a 12 well cell culture dish together with 1 mL of antibiotic-free culture media per well. (1 X 105 cells is ~ 350 &mu;L of cell suspension obtained from confluent T25 flask resuspended with 10 mL media).
3.2 Transfection
After 8-24 hrs, HEK cells will adhere to the plastic and are ready for transfection. Transfect cells with 0.2 &mu;g channel cDNA, 0.4 &mu;g M2R receptor cDNA and 0.04 &mu;g YFP cDNA using Invitrogen Lipofectamine 2000. Note: we use a small amount of YFP cDNA to determine which cells are successfully transfected (green cells will likely contain both GIRK channels and M2 receptors). The concentration of cDNA to use will need to be adjusted for each clone.
3.3 Fluorescence visualization of transfected cells.
Place 12-well plate on stage of an inverted fluorescence microscope, and examine cells for YFP expression. Compare to DIC image and make note of approximate number of green cells. This provides an estimate of the transfection efficiency.
3.4 Preparation of 24-well dishes and Poly-D-lysine coated cover slips
Clean 12 mm glass cover slips (Warner, CS-12R) by shaking them overnight with Radiacwash, followed by washing with deionized water and overnight shaking in 95% ethanol. Store the clean cover slips in 70% ethanol until use.
Place cover slips in a Petri dish with ethanol, flame off ethanol using forceps and burner, and place in one well of a sterile 24 well plate.
Cover each well with 250 &mu;L of poly-D-lysine (PDL) solution [0.2 mg/mL] and let the cover slips sit overnight immersed in PDL solution under sterile hood. Wrap the 24 well plate in Parafilm to avoid evaporation.
Next morning, aspirate PDL and wash 2x with sterile water. Leave plates open to dry under the hood, then cover. The 24 well dishes with coated cover slips can be wrapped in with sterile aluminum foil and stored at room temperature for weeks.
3.5 Plating transfected HEK293T cells on the cover slips
24h after transfection, split cells into 24-well dish (~4 X 103 cells/well) containing glass cover slips coated with poly-D-lysine. Note: transfected cells can be used up to 24-72 hrs post transfection. The cells are split to provide individual cover slips for patch clamp electrophysiology experiment.	
4. Electrophysiology
4.1 Preparing Solutions
Extracellular (bath) solution: 20 mM KCl, 140 mM NaCl, 0.5 mM CaCl2, 2 mM MgCl2 and 10 mM HEPES (pH 7.4 set with NaOH).
Aliquot 20K bath solution and add appropriate modulators
<ul>
	<li>20K + Ba2+: solution containing 1 mM BaCl2: a specific inhibitor of inward rectifying currents</li>
	<li>20K + Carbachol: solution with 5 &mu;M Carbachol: a specific M2R agonist; M2R couples to G-proteins that open GIRK channels</li>
	<li>20K + Ethanol: bath solution containing 100 mM EtOH to activate GIRK channels directly</li>
</ul>
Place bath solutions into syringes (solution containers) of the rapid exchange perfusion system. Note: We use a Warner pinch valve system to control the bath solutions. PE tubing is recommended.
Internal/electrode solution: 140 mM KCl, 20 mM NaCl, 5 mM EGTA, 5.4 mM MgCl2, 10 mM HEPES (pH 7.4) with 2.5 mM K2ATP and 0.3 &mu;M Li2GTP.
We make the electrode solution without ATP and GTP. We separately prepare aliquots of ATP and GTP, which are stored at -80&deg;C, and add to the electrode solution on the day of the experiment. We keep syringe with final electrode solution on ice to preserve the ATP/GTP. We sterile filter (0.2 &mu;m) the internal/external solutions to inhibit microbial growth and reduce possibility of clogging the tubes and the electrode.
4.2 Pulling electrodes
We use Narishige two-step vertical puller and borosilicate glass electrode from Warner Instrument (1.5 mm outer diameter, 0.86 mm inner diameter and 7.5 cm length) to obtain electrodes with 3-7 M&Omega; of resistance when filled with intracellular solution.
Note: It is necessary to experimentally determine parameters for the electrode puller and type of electrode glass in your lab. For our vertical puller, we use a heat setting of 60.2 for the first pull and ~43 for the second pull.
4.3 Whole-cell patch clamping
<ol>
	<li>Mount cover slip in a 35 mm dish using small amount of vaseline, cover with external solution and place dish on stage of an inverted fluorescence microscope. Use DIC to focus and find cells, switch to YFP fluorescence to locate YFP positive cells.</li>
	<li>Position tip of perfusion manifold near a green cell (20-30 &mu;m from the cell of interest=2-3 cell length apart).</li>
	<li>Fill electrodes with solution, attach it to the headstage of Axopatch 200B amplifier (Axon Instruments) with the following initial settings: Gain 1, Configuration: whole-cell &beta;=1, mode V-clamp.</li>
	<li>Apply positive pressure on electrode to prevent clogging of the pipette, and use the manipulator to place electrode tip just above the cell.</li>
	<li>To Zero the pipette potential, select the Seal Test button to deliver a test voltage step. The 5 mV rectangular voltage step will appear in the Seal Test window of Clampex. Adjust the Pipette Offset potentiometer on the amplifier to bring the base-line current to 0 nA. Alternatively, zero the pipette potential by switching the amplifier mode selector to V-track mode and the meter knob to Vtrack and use the Pipette Offset potentiometer on the amplifier to bring the voltage on the amplifier meter display to "0.00".</li>
	<li>Check resistance of electrode using seal test command of the Clampex 8.2 software (should be 3-7 M&Omega;).</li>
	<li>Open a saved protocol file or create a new protocol. To open protocol file select from the main menu Acquire then Open Protocol and choose a saved voltage protocol file. For this experiment, the protocol has been created to hold membrane potential at -40mV, deliver a single 50 ms voltage step to -100 mV and ramp to +40 mV over 200 ms. This protocol is executed every 2 seconds. The ramp protocol allows for observation of the reversal potential and inward rectification of GIRK channels. For voltage-gated ion channels, a rectangular voltage step may be more appropriate.</li>
	<li>Approach cell using fine adjustment of the manipulator, release positive pressure, after touching the cell surface apply negative pressure, and monitor increase in resistance in the Seal Test oscilloscope window. Once resistance is in the G&Omega; range, the electrode has sealed onto membrane (Gigaseal has formed).</li>
	<li>Apply a pulse of negative pressure to break the cell membrane and get access into the cell. Once in the whole-cell configuration, you will register increased capacitive currents.</li>
	<li>Switch to Membrane Test in Clampex and write down the access resistance Ra, membrane resistance Rm, membrane capacitance Cm and verify by eye the exponential fit of the theoretical curve to the real capacitative current. Membrane capacitance is proportional to cell size and will be used later to calculate current density.</li>
	<li>Switch back to Seal Test, set V-Out (membrane potential) to -40 mV and filter at 10 kHz</li>
	<li>Compensate for membrane capacitance and series resistance on the amplifier with the following steps:
	<ol class="lalpha">
 	<li>Switch the WHOLE CELL CAP switch on. Adjust WHOLE CELL CAP and SERIES RESISTANCE dials back and forth, until you get a flat test pulse. You may also need to adjust pipette capacitance FAST CAP knob.</li>
		<li>Turn on % PREDICTION knob to about 60-80%, again re-adjusting WHOLE CELL CAP and the SERIES RESISTANCE dial (you may have to adjust pipette capacitance compensation FAST CAP knob as well).</li>
		<li>Turn on % COMP- to 100% without oscillating the cell, while trying to keep the pulse flat by re-adjusting WHOLE CELL CAP and SERIES RESISTANCE (you may have to adjust pipette capacitance compensation FAST CAP knob as well). Adjust the lag by reducing the time constant.</li>
		<li>Write down values for Cm (membrane capacitance), Rs (series resistance), %COMP, %PRED and Lag time set on the amplifier in the notebook. The Cm and Rs should be approximately the same as the Cm and Rm measured in the Membrane Test above.</li>
 </ol>
 </li>
	<li>Turn on external solution valve to control the bath solution. Set the amplifier 8-pole Bessel filter to 2kHz, open a voltage clamp protocol and begin recording currents.</li>
	<li>We record GIRK currents using a ramp protocol that steps to -100 mV for 50 ms then ramps from 100 mV to +40 mV over 200 ms and is delivered every 2 seconds. The ramp protocol allows for observation of the reversal potential and for inward rectification property of GIRK channels. At room temperature (22-25&deg; C), the GIRK current should reverse near -50 mV (with 20 mM KCl in the bath), which is near the calculated equilibrium potential for K (EK = -50 mV), then remain fairly flat at potentials positive to EK.</li>
	<li>Experimental Protocol: After recording a stable baseline, switch to solution 20K + Carbachol and wait for peak of response, switch back to 20K bath solution, then apply 20K + Ethanol and wait for peak response, then switch to 20K bath solution, and finally to 20K + Ba2+ solution. During application of these drugs you will see changes in the amplitude of GIRK currents (most pronounced at -100 mV). Note the trace number corresponding to the application of different extracellular solutions in the notebook. A single Clampfit file will contain all of the sweeps for the entire experiment.</li>
</ol>
5. Data Analysis
We use Clampfit 9.2 software to analyze data GIRK currents at -100 mV.
<ol>
	<li>Open the file with the recording, set cursor at -100 mV. By default you will see all the traces overlaid on the screen. By pressing "Write cursors" icon you will create Excel-like spreadsheet with numerical data.</li>
	<li>By assigning X to the trace number column and Y to the cursor column and pressing "Create Graph" icon you can quickly draw a time-course plot of the experiment.</li>
	<li>Calculate basal current by subtracting current in presence of Ba2+ from current recorded in external perfusion solution alone ("20K current" minus "20K + Ba2+ current"). Calculate induced currents by subtracting basal current in 20K from the activated currents ("20K + Carbachol current" minus "20K current"; "20K + Ethanol current" minus "20K current"). Calculate current density (pA/pF) by dividing the current by membrane capacitance (read directly from the amplifier). Calculate percent activation by dividing induced currents by basal currents and multiplying by 100.</li>
</ol>
6. Representative Results
You should be able to record current from cells transfected with wild-type channels. For GIRK channels measured in the extracellular solution containing 20 mM K+ (and 145 mM K+ intracellular) they should exhibit strong inward rectification and a reversal potential of ~ -50 mV, which is an essential property of potassium channels. The wild-type current will increase strongly upon carbachol application (3-5 fold increase over basal current) and respond similarly to the 100 mM ethanol. Note: the sign convention for electrophysiology is that inward current is negative and outward current is positive. For the L257W mutation in the alcohol binding pocket of GIRK2, you will see attenuated responses to both carbachol and ethanol. This suggests that L257 is involved in alcohol and G protein activation of GIRK2 channels. For further examples of the results of GIRK2 mutagenesis, see recent publication by Aryal et al.1.
<img src="/files/ftp_upload/2189/2189fig1.jpg" alt="Figure 1" /> Figure 1. Window of Clampfit 8.0 shows data file for wild-type GIRK2 recording. Upper left panel shows superimposed sweeps recorded in response to a ramp voltage protocol in the presence of different external bah solutions. Cursor 1 is positioned at -100 mV. The cursor values are written to the spreadsheet (Right panel). Lower left panel shows the plot of sweep number vs., current. Note the increase in current (downward deflection) with ethanol (EtOH) and carbachol (Carb) and current inhibition with Ba2+.
<img src="/files/ftp_upload/2189/2189fig2.jpg" alt="Figure 2" /> Figure 2. Window of Clampfit 8.0 shows data file for a mutant GIRK2-L257W recording. Descriptions are same as in Figure 1. Note the loss of carbachol (Carb) activation and reduction of EtOH-induced currents in the mutant channels.

<ol>
	<li>Aryal, P., Dvir, H., Choe, S., Slesinger, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+discrete+alcohol+pocket+involved+in+GIRK+channel+activation.">A discrete alcohol pocket involved in GIRK channel activation.</a> Nature Neurosci. 12, 988-995 (2009).</li><li>Panchenko, V. A., Glasser, C. R., Mayer, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+similarities+between+glutamate+receptor+channels+and+K++channels+examined+by+scanning+mutagenesis.">Structural similarities between glutamate receptor channels and K+ channels examined by scanning mutagenesis.</a> J Gen Phys. 117, 345-360 (2001).</li><li>Doyle, D. A., Cabral, M. o. r. a. i. s., Pfuetzner, J., Kuo, R. A., Gulbis, A., Cohen, J. M., Chait, S. L., &amp;amp, B. T., MacKinnon, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+structure+of+the+potassium+channel:+molecular+basis+of+K++conduction+and+selectivity.">The structure of the potassium channel: molecular basis of K+ conduction and selectivity.</a> Science. 280, 69-77 (1998).</li><li>Nishida, M., Cadene, M., Chait, B. T., MacKinnon, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crystal+structure+of+a+Kir3.1-prokaryotic+Kir+channel+chimera.">Crystal structure of a Kir3.1-prokaryotic Kir channel chimera.</a> EMBO J. 26, 4005-4015 (2007).</li><li>Tao, X., Avalos, J. L., Chen, J., MacKinnon, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crystal+structure+of+the+eukaryotic+strong+inward-rectifier+K++channel+Kir2.2+at+3.1+&amp;Aring;+resolution.">Crystal structure of the eukaryotic strong inward-rectifier K+ channel Kir2.2 at 3.1 &amp;Aring; resolution.</a> Science. 326, 1668-1674 (2009).</li></ol>

No conflicts of interest declared.

Site-directed mutagenesis is a classical approach to study structure-function relationships of ion channels. It is based on the premise that by changing a critical residue of the ion channel one should be able to observe pronounced changes in the channel function. In contrast, many mutations in non-critical position can be introduced without major perturbations to the channel function. Typically, the residue in question is first mutated to alanine, a small non-polar amino acid, followed by mutation to tryptophan, the largest of amino acids2. If the residue is important to the channel function, both mutations will change channel activity. If a less essential residue is mutated, the effects of mutation, especially to alanine are often unremarkable. Tryptophan mutations are more likely to cause changes due to steric hindrances associated with the tryptophan size. Often, when the structural information is lacking, systematic mutation to alanine or tryptophan within a fragment of channel (called alanine or tryptophan scan) can be used to locate critical residues. It is important to note that multiple clones with different point mutations can be prepared in parallel allowing for testing multiple residues within relatively short time. Recently, multiple crystal structures of the channels have become available3,4,5. They can be used to propose potential key residues to be tested using the approach described in this protocol.
Measurements on wild-type channel should always accompany the measurement of a mutant channel to serve as a positive control to assure that transfection and electrophysiological experiments were done correctly. Point mutations may lead to lack of a functional current, a current similar to the wild-type or altered regulation of the channel activity. Lack of measurable current may be due to improper folding or altered trafficking. In such case it may be useful to find out whether the channel traffics correctly to the surface using a biotinylation assay or immunofluorescence staining against extracellular tag (without cell permeabilization) to distinguish between non-functional channels on the plasma membrane and channels retained inside the cell. Finally, it is advisable to investigate not only basal activity of the channel but also consequences of mutation on the channel regulation. Different mechanisms of modulation of the channel function could be studied depending of the ion channel of interest including regulation by G proteins, PIP2, ions, ATP concentration, phosphorylation (using tyrosine, serine or threonine mutation), ubiquitination or sumoylation (using lysine mutation) and direct channel openers or blockers.

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en"> 
 
 <div>
 <table border="1">
 <thead>
 <tr>
 <td>Material Name</td>
 <td>Type</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comment</td>
 </tr>
 </thead>
 <tbody>
 
<tr>
<td>Chemicals</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>Varies</td>
<td>&nbsp;</td>
<tr>
<td>Mini Prep Kit</td>
<td>&nbsp;</td>
<td>Qiagen</td>
<td>27104</td>
<td>&nbsp;</td>
<tr>
<td>Radiacwash</td>
<td>&nbsp;</td>
<td>Biodex</td>
<td>005-100</td>
<td>&nbsp;</td>
<tr>
<td>Poly-D-lysine hydrobromide</td>
<td>&nbsp;</td>
<td>BD Biosciences</td>
<td>CB40210</td>
<td>Poly-L-lysine can also be used</td>
</tr>
<tr>
<td>Maxi Prep Kit</td>
<td>&nbsp;</td>
<td>Qiagen</td>
<td>12263</td>
<td>&nbsp;</td>
<tr>
<td>QuickChange XL II Site Directed Mutagenesis Kit</td>
<td>&nbsp;</td>
<td>Stratagene</td>
<td>200512-5</td>
<td>&nbsp;</td>
<tr>
<td>Lipofectamine 2000</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>11668-027</td>
<td>&nbsp;</td>
<tr>
<td>Cover slips &#x02C6;&#x2026; 12 mm</td>
<td>&nbsp;</td>
<td>Warner Instruments</td>
<td>CS-12R</td>
<td>&nbsp;</td>
<tr>
<td>Capillary Glass Tubing Borosilicate-standard wall</td>
<td>&nbsp;</td>
<td>Warner Instruments</td>
<td>G150-3</td>
<td>&nbsp;</td>
<tr>
<td><a href="http://www.leica-microsystems.com/products/light-microscopes/clinical/inverted-microscopes/details/product/leica-dmi3000-b/" target="_blank">Inverted fluorescence microscope</a></td>
<td>&nbsp;</td>
<td>Leica</td>
<td>DMI3000 B</td>
<td>&nbsp;</td>
<tr>
<td>Electrophysiology</td>
<td>&nbsp;</td>
<td>Nikon/ Leica/ Zeiss</td>
<td>Any inverted DIC (Hoffmann) microscope with fluorescence</td>
<td>&nbsp;</td>
<tr>
<td>Amplifier</td>
<td>&nbsp;</td>
<td>Molecular Devices</td>
<td>Axon Axopatch 200B amplifier</td>
<td>&nbsp;</td>
<tr>
<td>Perfusion system</td>
<td>&nbsp;</td>
<td>Warner Instruments</td>
<td>VC-6 Perfusion MM-6</td>
<td>&nbsp;</td>
<tr>
<td>Digitizer</td>
<td>&nbsp;</td>
<td>Molecular Devices</td>
<td>Digidata 1320 digitizer</td>
<td>&nbsp;</td>
<tr>
<td>Manipulators</td>
<td>&nbsp;</td>
<td>Sutter Instruments</td>
<td>Manipulator: MP-85</td>
<td>&nbsp;</td>
<tr>
<td>Electrode Puller</td>
<td>&nbsp;</td>
<td>Narishige</td>
<td>PC10 vertical electrode puller</td>
<td>&nbsp;</td>
<tr>
<td>Isolation Table</td>
<td>&nbsp;</td>
<td>Technical Manufacturing Corporation</td>
<td>Air table</td>
<td>&nbsp;</td> </tbody>
 </table>
 </div>

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.

Watch this Scientific Journal Video about Mutagenesis and Functional Analysis of Ion Channels Heterologously Expressed in Mammalian Cells at JoVE.com

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

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 ...

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17.3K Views.  Salk Institute for Biological Studies. We will demonstrate how to study the effect of a single point mutation on the function of an ion channel.

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

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 ...

Pulse-chase Analysis of N-linked Sugar Chains from Glycoproteins in Mammalian Cells

Attachment of the Glc3Man9GlcNAc2 precursor oligosaccharide to nascent polypeptides in the ER is a common modification for secretory proteins. Although this modification was implicated in several biological processes, additional aspects of its function are emerging, with recent evidence of its role in the production of signals for glycoprotein quality control and trafficking. Thus, phenomena related to N-linked glycans and their processing are being intensively investigated. Methods that have been recently developed for proteomic analysis have greatly improved the characterization of glycoprotein N-linked glycans. Nevertheless, they do not provide insight into the dynamics of the sugar chain processing involved. For this, labeling and pulse-chase analysis protocols are used that are usually complex and give very low yields. We describe here a simple method for the isolation and analysis of metabolically labeled N-linked oligosaccharides. The protocol is based on labeling of cells with [2-3H] mannose, denaturing lysis and enzymatic release of the oligosaccharides from either a specifically immunoprecipitated protein of interest or from the general glycoprotein pool by sequential treatments with endo H and N-glycosidase F, followed by molecular filtration (Amicon). In this method the isolated oligosaccharides serve as an input for HPLC analysis, which allows discrimination between various glycan structures according to the number of monosaccharide units comprising them, with a resolution of a single monosaccharide. Using this method we were able to study high mannose N-linked oligosaccharide profiles of total cell glycoproteins after pulse-chase in normal conditions and under proteasome inhibition. These profiles were compared to those obtained from an immunoprecipitated ER-associated degradation (ERAD) substrate. Our results suggest that most NIH 3T3 cellular glycoproteins are relatively stable and that most of their oligosaccharides are trimmed to Man9-8GlcNAc2. In contrast, unstable ERAD substrates are trimmed to Man6-5GlcNAc2 and glycoproteins bearing these species accumulate upon inhibition of proteasomal degradation.

We describe a method for analysis of the alteration of N-linked glycans through the early life of glycoproteins after their biosynthesis in mammalian cells. This is achieved by pulse-chase analysis of metabolically labeled glycans, enzymatic release from glycoproteins and examination by HPLC.

Attachment of the Glc3Man9GlcNAc2 precursor oligosaccharide to nascent polypeptides in the ER is a common modification for secretory proteins. Although ...

Analysis of DNA Double-strand Break (DSB) Repair in Mammalian Cells

DNA double-strand breaks are the most dangerous DNA lesions that may lead to massive loss of genetic information and cell death. Cells repair DSBs using two major pathways: nonhomologous end joining (NHEJ) and homologous recombination (HR). Perturbations of NHEJ and HR are often associated with premature aging and tumorigenesis, hence it is important to have a quantitative way of measuring each DSB repair pathway. Our laboratory has developed fluorescent reporter constructs that allow sensitive and quantitative measurement of NHEJ and HR. The constructs are based on an engineered GFP gene containing recognition sites for a rare-cutting I-SceI endonuclease for induction of DSBs. The starting constructs are GFP negative as the GFP gene is inactivated by an additional exon, or by mutations. Successful repair of the I-SceI-induced breaks by NHEJ or HR restores the functional GFP gene. The number of GFP positive cells counted by flow cytometry provides quantitative measure of NHEJ or HR efficiency.

Analysis of DNA Double-strand Break DSB Repair in Mammalian Cells

This article describes GFP-based fluorescence in vivo assays that separately quantify homologous recombination and nonhomologous end joining in mammalian cells.

DNA double-strand breaks are the most dangerous DNA lesions that may lead to massive loss of genetic information and cell death.  Cells repair DSBs using ...

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 ...

Optimized Transfection Strategy for Expression and Electrophysiological Recording of Recombinant Voltage-Gated Ion Channels in HEK-293T Cells

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) is a ubiquitous research strategy. HEK-293T cells must be plated onto glass coverslips at low enough density so that they are not in contact with each other in order to allow for electrophysiological recording without confounding effects due to contact with adjacent cells. Transfected channels must also express with high efficiency at the plasma membrane for whole-cell patch clamp recording of detectable currents above noise levels. Heterologous ion channels often require long incubation periods at 28&deg;C after transfection in order to achieve adequate membrane expression, but there are increasing losses of cell-coverslip adhesion and membrane stability at this temperature. To circumvent this problem, we developed an optimized strategy to transfect and plate HEK-293T cells. This method requires that cells be transfected at a relatively high confluency, and incubated at 28&deg;C for varying incubation periods post-transfection to allow for adequate ion channel protein expression. Transfected cells are then plated onto glass coverslips and incubated at 37&deg;C for several hours, which allows for rigid cell attachment to the coverslips and membrane restabilization. Cells can be recorded shortly after plating, or can be transferred to 28&deg;C for further incubation. We find that the initial incubation at 28&deg;C, after transfection but before plating, is key for the efficient expression of heterologous ion channels that normally do not express well at the plasma membrane. Positively transfected, cultured cells are identified by co-expressed eGFP or eGFP expressed from a bicistronic vector (e.g. pIRES2-EGFP) containing the recombinant ion channel cDNA just upstream of an internal ribosome entry site and an eGFP coding sequence. Whole-cell patch clamp recording requires specialized equipment, plus the crafting of polished recording electrodes and L-shaped ground electrodes from borosilicate glass. Drug delivery to study the pharmacology of ion channels can be achieved by directly micropipetting drugs into the recording dish, or by using microperfusion or gravity flow systems that produce uninterrupted streams of drug solution over recorded cells.

Reliable method for highly efficient in vitro expression and subsequent electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T).

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) ...

Analysis of mRNA Nuclear Export Kinetics in Mammalian Cells by Microinjection

In eukaryotes, messenger RNA (mRNA) is transcribed in the nucleus and must be exported into the cytoplasm to access the translation machinery. Although the nuclear export of mRNA has been studied extensively in Xenopus oocytes1 and genetically tractable organisms such as yeast2 and the Drosophila derived S2 cell line3, few studies had been conducted in mammalian cells. Furthermore the kinetics of mRNA export in mammalian somatic cells could only be inferred indirectly4,5. In order to measure the nuclear export kinetics of mRNA in mammalian tissue culture cells, we have developed an assay that employs the power of microinjection coupled with fluorescent in situ hybridization (FISH). These assays have been used to demonstrate that in mammalian cells, the majority of mRNAs are exported in a splicing dependent manner6,7, or in manner that requires specific RNA sequences such as the signal sequence coding region (SSCR) 6. In this assay, cells are microinjected with either in vitro synthesized mRNA or plasmid DNA containing the gene of interest. The microinjected cells are incubated for various time points then fixed and the sub-cellular localization of RNA is assessed using FISH. In contrast to transfection, where transcription occurs several hours after the addition of nucleic acids, microinjection of DNA or mRNA allows for rapid expression and allows for the generation of precise kinetic data.

Here we describe an assay that employs the power of microinjection coupled with fluorescent in situ hybridization in order to accurately measure the nuclear export kinetics of mRNA in mammalian somatic cells.

In eukaryotes, messenger RNA (mRNA) is transcribed in the nucleus and must be exported into the cytoplasm to access the translation machinery. Although ...

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular signatures, or to evolve proteins in the laboratory. Here, we present a protocol that allows the generation of large (>1011) mutant libraries for a given target sequence. This method is based on replication of a ColE1 plasmid encoding the desired sequence by a low-fidelity variant of DNA polymerase I (LF-Pol I). The target plasmid is transformed into a mutator strain of E. coli and plated on solid media, yielding between 0.2 and 1 mutations/kb, depending on the location of the target gene. Higher mutation frequencies are achieved by iterating this process of mutagenesis. Compared to alternative methods of mutagenesis, our protocol stands out for its simplicity, as no cloning or PCR are involved. Thus, our method is ideal for mutational labeling of plasmids or other Pol I templates or to explore large sections of sequence space for the evolution of activities not present in the original target. The tight spatial control that PCR or randomized oligonucleotide-based methods offer can also be achieved through subsequent cloning of specific sections of the library. Here we provide protocols showing how to create a random mutant library and how to establish drug-based selections in E. coli to identify mutants exhibiting new biochemical activities.

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

Here we demonstrate a simple protocol to create a random mutant library for a given target sequence. We show how this method, which is performed in vivo in Escherichia coli, can be coupled with functional selections to evolve new enzymatic activities.

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular ...

Analysis of Cell Cycle Position in Mammalian Cells

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of cell proliferation underlies the pathology of diseases like cancer. As such there is great need to be able to investigate cell proliferation and quantitate the proportion of cells in each phase of the cell cycle. It is also of vital importance to indistinguishably identify cells that are replicating their DNA within a larger population. Since a cell&prime;s decision to proliferate is made in the G1 phase immediately before initiating DNA synthesis and progressing through the rest of the cell cycle, detection of DNA synthesis at this stage allows for an unambiguous determination of the status of growth regulation in cell culture experiments.
DNA content in cells can be readily quantitated by flow cytometry of cells stained with propidium iodide, a fluorescent DNA intercalating dye. Similarly, active DNA synthesis can be quantitated by culturing cells in the presence of radioactive thymidine, harvesting the cells, and measuring the incorporation of radioactivity into an acid insoluble fraction. We have considerable expertise with cell cycle analysis and recommend a different approach. We Investigate cell proliferation using bromodeoxyuridine/fluorodeoxyuridine (abbreviated simply as BrdU) staining that detects the incorporation of these thymine analogs into recently synthesized DNA. Labeling and staining cells with BrdU, combined with total DNA staining by propidium iodide and analysis by flow cytometry1 offers the most accurate measure of cells in the various stages of the cell cycle. It is our preferred method because it combines the detection of active DNA synthesis, through antibody based staining of BrdU, with total DNA content from propidium iodide. This allows for the clear separation of cells in G1 from early S phase, or late S phase from G2/M. Furthermore, this approach can be utilized to investigate the effects of many different cell stimuli and pharmacologic agents on the regulation of progression through these different cell cycle phases.
In this report we describe methods for labeling and staining cultured cells, as well as their analysis by flow cytometry. We also include experimental examples of how this method can be used to measure the effects of growth inhibiting signals from cytokines such as TGF-&beta;1, and proliferative inhibitors such as the cyclin dependent kinase inhibitor, p27KIP1. We also include an alternate protocol that allows for the analysis of cell cycle position in a sub-population of cells within a larger culture5. In this case, we demonstrate how to detect a cell cycle arrest in cells transfected with the retinoblastoma gene even when greatly outnumbered by untransfected cells in the same culture. These examples illustrate the many ways that DNA staining and flow cytometry can be utilized and adapted to investigate fundamental questions of mammalian cell cycle control.

Determining the cell cycle position of a population of cells, or understanding how signals affect proliferation, can be readily measured by flow cytometry using this protocol. We report a simple experimental approach to staining cells and quantifying their position in the cell cycle.

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of ...

Induction and Analysis of Epithelial to Mesenchymal Transition

Epithelial to mesenchymal transition (EMT) is essential for proper morphogenesis during development. Misregulation of this process has been implicated as a key event in fibrosis and the progression of carcinomas to a metastatic state. Understanding the processes that underlie EMT is imperative for the early diagnosis and clinical control of these disease states. Reliable induction of EMT in vitro is a useful tool for drug discovery as well as to identify common gene expression signatures for diagnostic purposes. Here we demonstrate a straightforward method for the induction of EMT in a variety of cell types. Methods for the analysis of cells pre- and post-EMT induction by immunocytochemistry are also included. Additionally, we demonstrate the effectiveness of this method through antibody-based array analysis and migration/invasion assays.

A straightforward method is described for the induction of epithelial to mesenchymal transition (EMT) in a variety of cell types. Methods for the analysis of cells in EMT states by immunocytochemistry are included.

Epithelial  to mesenchymal transition (EMT) is essential for proper morphogenesis during  development. Misregulation of this  process has been implicated ...

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of cell heath due to their role in both cell metabolism and energy production as well as control of apoptosis. Therefore, direct evaluation of isolated mitochondria and mitochondrial perturbation offers the ability to determine if organelle-specific (dys)function is occurring. The methods described in this protocol include isolation of intact, functional mitochondria from HEK cultured cells and mouse liver and spinal cord, but can be easily adapted for use with other cultured cells or animal tissues. Mitochondrial function assessed by TMRE and the use of common mitochondrial uncouplers and inhibitors in conjunction with a fluorescent plate reader allow this protocol not only to be versatile and accessible to most research laboratories, but also offers high throughput.

Comparison of mitochondrial membrane potential between samples yields valuable information about cellular status. Detailed steps for isolating mitochondria and assessing response to inhibitors and uncouplers using fluorescence are described. The method and utility of this protocol are illustrated by use of a cell culture and animal model of cellular stress.

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of ...