The authors would like to thank Eric Fishback (Des Moines University, Des Moines, Iowa, USA) for his assistance with shooting and editing the video. The PS120 fibroblast-like cells stably expressing NHE3-wt or NHE3-YRK were kindly provided by Dr. Mark Donowitz (Johns Hopkins University School of Medicine, Baltimore, MD, USA).

Note: Two amplifiers are required for recording the activity of electroneutral transporters by a self-referencing ISE in conjunction with patch clamping, a patch clamp amplifier to maintain the cell in a whole-cell configuration and a high impendence amplifier (electrometer) to record the transporter activity via the ISE (see Table of Materials). The patch clamp amplifier is directly connected to the acquisition board terminal. The electrometer is connected to the differential amplifier utilized to filter and amplify the signal. The differential amplifier is connected to the acquisition board terminal. Capmeter, patch clamp software that monitors cell capacity, has been used in previous studies20 (see Table of Materials). The recording chamber and perfusion solutions are maintained at 37 &#176;C.
1. Preparation of Pipettes for ISEs
<ol>
	<li>Pull the pipettes from borosilicate glass capillaries to an outer diameter of 1.2 mm (see Table of Materials) using the pipette puller2 .</li>
	<li>Polish the pipette using the microforge. The diameter of the pipette tip should be 2-3 &#181;m.</li>
	<li>Place the pipettes in a jar holder for pipettes with the pipette tip facing up (see Table of Materials). 
	CAUTION: Perform steps 1.4-1.6 in a well-ventilated chemical fume hood.</li>
	<li>Mix the siliconization mixture that is composed of 1 drop (~150 &#181;L) of bis(dimethylamino)dimethyl silane and 10 drops of carbon tetrachloride (see Table of Materials).</li>
	<li>Pour the siliconization mixture into the jar and close the lid tightly.</li>
	<li>Incubate the jar with the siliconization mixture for 24-48 h in a chemical fume hood at room temperature until the mixture is completely evaporated.</li>
	<li>Backfill the pipette with the appropriate ion-selective resin to create a column of approximately 2-4 mm. 
	NOTE: Hydrogen ionophore I cocktail B can be usedfor recording NHE activity (see Table of Materials).</li>
	<li>Tap the tip of the pipette to distribute the ion-selective resin to the tip, and place the pipette vertically in the jar holder with the pipette tip down.</li>
	<li>Checkunder the dissecting microscope that the resin cocktail has reached the end of the tip. If needed, apply positive pressure to push the resin to the end.</li>
	<li>Backfill the half-pipette with the appropriate solution (~ 100 &#956;L). Here, use 100 mM KCl and 10 mM HEPES at pH 7.0 if using ionophore I cocktail B.</li>
	<li>Connect the ISE to the electrometer and place the tip of the ISE into the bath solution used for recording. Zero the electrometer.</li>
	<li>Test the H+-selective ISE response by changing the pH of the bath solution. The average H+-selective ISE response is ~ 58 mV per unit of pH.</li>
	<li>Periodically evaluate the ISE response to calibration.</li>
</ol>
2. Preparation of Patch Clamp Pipettes for Recording
NOTE: The method described by Donald Hilgemann in his article on Single-Channel Recording21 is recommended.
<ol>
	<li>Pull the pipettes for giant patch from borosilicate glass capillaries (2.0 mmOD / 1.5 mm ID) on the pipette puller2,22,23.</li>
	<li>Use a microforge mounted on the stage of an inverted microscope to make a wide pipette tip (10 - 22 &#956;m)23. Melt a bead (~ 0.5 mL) of soft glass onto the thick platinum wire of the microforge. 
	NOTE: A relatively thick platinum wire is required (~0.5 mm) to fabricate giant patch pipettes.</li>
	<li>Position the patch pipette close to the soft glass bead, and apply full heat until the tip begins to recede. The platinum wire will move slightly toward the centerduring heating.</li>
	<li>Turn off the heat, and push the pipette tip into the softened glass bead. The cooling wire will retract and break the pipette tip.</li>
	<li>Reapply the heat to smooth the pipette tip and create a pipette tip of the desired diameter. The diameter of the tip is dependent on the size of the cell to be patched (8 - 10 &#956;m in this study).</li>
</ol>
3. Preparation of Cells
<ol>
	<li>Warm up phosphate buffer saline (PBS) without Ca2+ and Mg2+, trypsin-EDTA solution and growth medium to at 37 &#176;C.</li>
	<li>Wash cells twice with PBS. Use 1 mL of the buffer for 35 mm diameter cell culture plate.</li>
	<li>Add 0.5 mL of trypsin to the cells for 35 mm diameter cell culture plate. 
	Note: When cells start to round-up, shake the plate to de-attach the cells.</li>
	<li>Stop the trypsin action by adding 5 mL (amount for 35 mm diameter cell culture plate) of complete growth medium supplemented with 10% fetal bovine serum. 
	NOTE: The reaction time in trypsin is cell-type dependent.</li>
	<li>Place the cells on a shaker with rocking to prevent formation of clumps.</li>
	<li>Use cells for up to 2 h after trypsinization. 
	NOTE: The time might vary depending on the cell line.</li>
</ol>
4. Preparation of an Intra-Pipette Perfusion System
<ol>
	<li>Cut an appropriate size of quartz tubing (150 &#181;m OD / 75 &#181;m ID) to prepare the intra-pipette perfusion line.</li>
	<li>Insert one side of the quartz tubing into a 200 &#181;L pipette tip. Glue the quartz tubing onto the tip, and cut the end of the tip with a sharp razor blade.</li>
	<li>Connect the perfusion line (quartz tubing glued to the tip) to a small piece of silicon tubing that would serve as a reservoir for the intra-pipette perfusion solution.</li>
	<li>Connect the silicon tubing to a syringe to create positive pressure.</li>
	<li>Insert the free end of the quartz tubing through the polyethylene tube of the pipette holder of the perfusion port (see Table of Materials).</li>
	<li>Evaluate the perfusion line with the pipette solution.</li>
</ol>
5. Preparation of Solutions
<ol>
	<li>Prepare a heavily buffered pipette solution for recording NHE (115 mM potassium aspartate (KAsp), 1 mM EGTA, 0.5 mM MgCl2, 10 mM Mg-ATP, 12.5 mM HEPES, 12.5 mM PIPES, 12.5 mM MOPS, and 12.5 mM MES, pH 6.0 adjusted with aspartic acid). 
	NOTE: Prepare the stock pipette solution and filter. Add Mg-ATP before recording.</li>
	<li>Prepare a fresh bath solution each time (140 mM NaCl, 2 mM CaCl2, 2 mM MgCl2, and 0.1 mM Tris-HCl pH 8.2).</li>
</ol>
6. Recording of Transporter Activity
<ol>
	<li>Maintain the recording chamber and all solutions at 37 &#176;C, because transporter activity is temperature-dependent.</li>
	<li>Place the cells into the recording chamber. Wait until they drop to the bottom of the chamber but do not let them attach to the chamber.</li>
	<li>Choose an appropriate cell. Measure the diameter of the cell with the ocular micrometer. Choose cells with similar diameters. 
	NOTE: Generally, a large cell will have more cellular surface and possibly more transporters expressed on its plasma membrane, thus also having more transporter activity. However, this is not necessarily correct, as transporter activity depends on the specific transporter and the cell type under study.</li>
	<li>Mount the ion-selective microelectrode pipette to the micromanipulator, backfill the patch-clamping pipette with the intracellular solution, make sure that the solution reaches the tip of the pipette2. 
	NOTE: It might require several gentle taps to eliminate bubbles in the tip of the pipette.</li>
	<li>Mount the patch-clamping pipette to the head stage of the electrophysiology set up2,22,23. 
	NOTE: It is important to keep the head stage at about a 45-degree angle to the horizontal axis, as this ensures an entry angle for the pipette that is suitable for the formation of a highly resistant seal.</li>
	<li>Apply a small positive pressure (~ 5 cm H2O) to the patch pipette to reduce the chance of blocking the tip, and place it into the bath solution.</li>
	<li>Stop moving the patch clamp pipette once the tip is close to the cell.</li>
	<li>Relieve the positive pressure while pipette touches the cell and apply slight negative pressure (~ 5 cm H2O) to obtain a &#62;1000 M&#937; (a Giga-ohm) seal on the cell membrane with the patch pipette23.</li>
	<li>Place the patch pipette with the cell in the same focal plane as the ISE, and move the patch pipette with the cell in proximity to the ISE (~ 5 - 10 &#181;m)2,5 but avoid allowing the cell to contact the ISE (Figure 1A).</li>
	<li>Assure that the holding potential is 0 mV.</li>
	<li>Rupture the seal with series of short suctions to obtain the whole-cell configuration.</li>
	<li>Move the ISE (or patch clamp pipette) slowly left-right or up-down to record the activity of the transporter (Figure 1B).</li>
	<li>Record at least 5-8 peaks (Figure 1C, from A to B).</li>
	<li>Apply square wave perturbations during the recording to monitor the quality of the seal and the cell membrane capacitance (Figure 1).</li>
	<li>Estimate the H+ flux (<img alt="Equation 1" src="/files/ftp_upload/56630/56630eq1.jpg" />) from the difference in free H+ concentration between the cell surface and the bulk solution (&#916;H+) 
	<img alt="Equation 2" src="/files/ftp_upload/56630/56630eq2.jpg" /> 
	<img alt="Equation 3" src="/files/ftp_upload/56630/56630eq3v2.jpg" /> 
	NOTE: <img alt="Equation 4" src="/files/ftp_upload/56630/56630eq4.jpg" /> and&#160;<img alt="Equation 5" src="/files/ftp_upload/56630/56630eq5.jpg" /> are H+ and buffer diffusion coefficients respectively, BT is the total buffer concentration, KD is the dissociation constant of the buffer, r is the cell radius, and <img alt="Equation 6" src="/files/ftp_upload/56630/56630eq6.jpg" /> determines the steady-state change in the concentration of bound H+ for a small change in free H+. H+ indicates a free proton in solution. To be consistent with previous studies, H+ fluxes are converted to electrophysiological flux units by calculating the current equivalents of the fluxes (<img alt="Equation 1" src="/files/ftp_upload/56630/56630eq1.jpg" />/faraday) in pA3,5].</li>
	<li>Normalize the proton flux to the cell surface that can be calculated from the cell size or to the membrane capacitance.</li>
</ol>

<ol>
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The authors have no conflicts of interest to disclose.

Despite the crucial functions of transporters, the methods available to study their activity are ineffective and inadequate. One limitation is that the available methods measure ion movement mediated by transporter activity without considering fluctuations in intracellular ion composition during the experiment4. The presented method ensures precise control of extracellular and intracellular ion compositions and offers a powerful tool for modifying intracellular ion, protein and lipid compositions3,4,5.
The most important step in this protocol is the preparation of an ISE that is stable for a long period25. In our experience, an unstable ISE is not well-siliconized (e.g., the ion-selective resin is redistributed on the ISE pipette sides), which can be detected by a constant and uncontrollable drift in the ISE signal. Additionally, the resin distribution in the ISE should be closely monitored during the experiment. The bath solution may push the ion-selective resin inside the ISE pipette, creating a column of bath solution before the resin25. In this case, the ISE will record the ion concentration from this column and become nonresponsive to changes in the ion gradients during the experiment. Adjusting the geometry of the ISE is key to obtain an ISE response in the range of 100 ms9 and overcome possible delays in ISE time-responses compared with changes in ion gradients.
The major limitation of the presented method is inherited from the patch clamping measurement of transporter activity in epithelial cells. When detached from the support and maintained in suspension, polarized epithelial cells might lose the polarized localization of transporters on the apical or basolateral membrane.
A modified patch clamping measurement should be used when studying channels and transporters in polarized epithelia. Considering that cells might approximate into a cylinder rather than a sphere is important when transforming the H+ gradient into flux via the formula provided. In this case, the formula should be changed to <img alt="Equation 7" src="/files/ftp_upload/56630/56630eq7.jpg" /> where &#955; is the length of the cylinder, r is the cylinder radius, ad x is the radial distance from the cylinder midpoint over which the gradient is measured5. Future development of this powerful technique for studying electroneutral transporters should focus on modifying the method to optimize recording cells grown in a monolayer.

Transport of ions and solutes across the plasma membrane is essential for the survival of cells and, hence, of organisms1. Selective transport of ions and solutes is achieved by an array of specialized channel and transporter proteins. Mutations in these proteins often result in a variety of clinical conditions, rendering channel and transporter proteins potential targets for pharmacological treatment1. Unfortunately, understanding the mechanisms underlying channel and transporter function and regulation is often limited by the approaches available to study their activity2,3,4.
Specifically, transporters can be roughly sub-divided into two large groups depending on whether they alter the cell transmembrane potential during transport of solutes: the altering electrogenic ion transporters [e.g., sodium-phosphate co-transporter 2a (NaPi2a), sodium-calcium exchanger (NCX), etc.] or the non-altering electroneutral ion transporters [e.g., sodium-proton exchanger (NHE), sodium-chloride co-transporter, NaPi2c, etc.]. The activities of both classes of transporters have been studied extensively using uptake of radioactive isotopes and fluorescent membrane-permeant dyes2. Both approaches estimate the activity of transporters by measuring changes in the bulk concentration of specific cytoplasmic ions, and both methods have limitations, such as moderate sensitivity and time resolution and inadequate control of the intracellular milieu. Indeed, the activity of many transporters is dependent on the cytoplasmic concentration of the carried ions (e.g., NHE3, NCX), and changes in these ion concentrations are expected to play a significant role in regulating transporter activity2,3,5. Precise measurement of these regulative mechanisms is limited using classical methods.
To overcome these limitations, patch clamp methods are used to study the transporter activity2,6. Specifically, the self-referencing ion-selective electrodes (ISEs)7,8 combined with the patch clamping system has recently allowed the measurement of electroneutral transporter activity3,4,5. ISEs are based on the fact that transporter activity creates an ion gradient in close proximity to the cell membrane. An ISE moving up to and away from the cell membrane in a repetitive, oscillatory fashion records a voltage difference (&#181;V). Voltage differences can be converted into ion flux values using a calibration method that applies Fick&#39;s first law of diffusion2,9. While ISEs are used to detect the flux of ions moving out of cells, the patch clamp method in both whole-cell or inside-out configurations is used to control the membrane potential and intracellular ion composition. Moreover, the application of the giant patch clamp technique allows modification of the intracellular composition of not only ions but also lipids and proteins3,5.
In summary, the versatility of the patch clamp method compared with that of other methods to study transporter activity has made patch clamping suitable to overcome the common limitations of these other methods. The combination of self-referencing ISEs and patch clamp techniques offers the unique possibility to measure the activity of electroneutral transporters in a tightly controlled experimental environment and to discover novel biophysical and molecular properties of cell membrane transport3,4,5. This approach has been successfully used to study the activity of the NHE. The mammalian NHE protein family catalyzes the electroneutral net exchange of extracellular sodium (Na+) for intracellular proton (H+)10,11 utilizing an inward Na+ gradient. In mammals, the NHE protein family includes 11 related proteins (NHE1-9 and NHA1-2) and a sperm-specific NHE10,12,13.
NHEs (SLC9a family) are found ubiquitously in most living organisms from simple prokaryotes to higher eukaryotes and are involved in a variety of vital cell functions10,11, including controlling the cell salinity defense in prokaryotes, maintaining acid-base homeostasis and cell volume, and regulating the absorption of salt and water in various specialized epithelia10,12,14,15. The key biological roles of NHEs and the significance of their functions have been determined through several studies; however, few studies have investigated the biophysical and molecular properties of mammalian NHEs because of methodological limitations4. Recently, the application of self-referencing ISEs during whole-cell patch clamping has revealed novel molecular mechanisms of NHE isoforms regulated by changes in the intracellular concentrations of ions, proteins and phospholipids3,4.
Specifically, the protocol provided in this manuscript outlines the methods and approaches for studying the activity and regulation of NHE isoform 3 (NHE3), a major player in the absorption of Na+, Cl-, HCO3- and fluid in the brush border membrane of renal and intestinal epithelia14,16. New insight into differences in the sensitivity of NHE3 activity to intracellular phosphoinositides (phosphatidylinositide 4,5-bisphosphate [PI(4,5)P2] and phosphatidylinositide 3,4,5-triphosphate [PI(3,4,5]P3]) is reported. Cell transport proteins, such as channels and transporters, are regulated by phosphoinositides17, and NHE3 directly binds both PI(4,5)P2 and PI(3,4,5)P318. Based on the current literature, either phosphoinositide could be relevant for the physiological or pathophysiological regulation of NHE35,18,19. Our findings support separate roles for PI(4,5)P2 and PI(3,4,5)P3 in the regulation of NHE3 activity. This distinction was possible because of the application of ISE techniques in combination with whole-cell patch clamp recording. This technique also allows control of the phosphoinositide cellular content via the intracellular perfusion of different phosphoinositides during the measurement of NHE3 activity.

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			<td height="34" style="height:34px;width:367px;">Patch clamp Amplifier</td>
			<td style="width:219px;">Molecular Devices</td>
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		</tr>
		<tr height="34">
			<td height="34" style="height:34px;">Dual Channel Differential Electrometer (HiZ-223)</td>
			<td style="width:219px;">Warner Instruments</td>
			<td style="width:101px;">64-1650</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;">Differential Amplifier (DP-301)</td>
			<td style="width:219px;">Warner Instruments</td>
			<td style="width:101px;">64-0044</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;">Patch Clamp Software Based on MatLab</td>
			<td style="width:219px;">MatLab with acquisition toolbox</td>
			<td style="width:101px;"></td>
			<td>The capmeter software is recommended</td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:367px;">ThermoClamp-1 Temperature Control System</td>
			<td>&#160;AutoMate Scientific</td>
			<td style="width:101px;">03-11-LL</td>
			<td>In-line heater</td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:367px;">Single-Channel Temperature Controller (TC-324C)</td>
			<td style="width:219px;">Warner Instruments</td>
			<td style="width:101px;">64-2400</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:367px;">Single-Barrel Standard Borosilicate Glass Tubing</td>
			<td>World Precision Instruments</td>
			<td style="width:101px;">1B120F-3&#160;&#160;</td>
			<td style="width:276px;">Used for ion selective electrodes</td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:367px;">Micropipette Storage Jar</td>
			<td>World Precision Instruments</td>
			<td>E212</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:367px;">Bis(dimethylamino)dimethylsilane</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td>14755-100ML</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:367px;">Carbon tetrachloride</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td colspan="2">319961-500ML</td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:367px;">Hydrogen ionophore I - cocktail B</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td style="width:101px;">95293</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:367px;">Thin Wall Borosilicate Tubing</td>
			<td style="width:219px;">&#160;Sutter Instrument</td>
			<td style="width:101px;">B200-156-15</td>
			<td style="width:276px;">Used for patch clamp pipette&#160;</td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:367px;">Soft glass (Corning 8161 Patch Glass)</td>
			<td style="width:219px;">Warner Instruments</td>
			<td style="width:101px;">64-0815</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;">Silica Capillary Tubing (150um OD/75um ID)</td>
			<td style="width:219px;">Molex (Polymicro Technologies)</td>
			<td>106815-0018</td>
			<td style="width:276px;">Used for intra-pipette perfusion system</td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;">Potassium chloride</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td>P9333</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">Potassium aspartate&#160;</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td>A6558</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">EGTA</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td>E3889</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">Magnesium chloride</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td style="width:101px;">M4880</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">Mg-ATP</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td style="width:101px;">A9187</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">HEPES</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td>H3375</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">PIPES</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td>P6757</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">MOPS</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td>M1254</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">MES</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td>M3671</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">Calcium chloride</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td>C5670</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">Tris</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td style="width:101px;">T1503</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">Hydrochloric acid</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td>H1758</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:367px;">Apyrase</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td>A6535</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;">Phosphatidylinositol(4,5) bisphosphate diC8</td>
			<td>Echelon Biosciences</td>
			<td>P-4508</td>
			<td></td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;">Phosphatidylinositol 3,4,5-trisphosphate diC8</td>
			<td>Echelon Biosciences</td>
			<td>P-3908</td>
			<td></td>
		</tr>
	</tbody>
</table>

application of electrophysiology measurement to study the activity of electro-neutral transporters

The transport of ions through cell membranes ensures the fine control of ion content within and outside the cell that is indispensable for cell survival. These transport mechanisms are mediated by the activities of specialized transporter proteins. Specifically,pH dynamics are finely controlled by plasma membrane proton (H+) extrusion systems, such as the Na+/H+ exchanger (NHE) protein family. Despite extensive efforts to study the mechanisms underlying NHE regulation, our current understanding of the biophysical and molecular properties of the NHE family is inadequate because of the limited availability of methods to effectively measure NHE activity. In this manuscript, we used H+-selective electrodes during whole-cell patch clamping recording to measure NHE-induced H+ flux. We proposed this approach to overcome some limitations of typically used methods to measure NHE activity, such as radioactive uptake and fluorescent membrane permeants. Measurement of NHE activity using the described method enables high sensitivity and time resolution and more efficient control of intracellular H+ concentrations. H+-selective electrodes are based on the fact that transporter activity creates an ion gradient in close proximity to the cell membrane. An H+-selective electrode moving up to and away from the cell membrane in a repetitive, oscillatory fashion records a voltage difference that is dependent on H+ flux. While H+-selective electrodes are used to detect H+ flux moving out of the cell, the patch clamp method in the whole-cell configuration is used to control the intracellular ion composition. Moreover, application of the giant patch clamp technique allows modification of the intracellular composition of not only ions but also lipids. The transporter activity of NHE isoform 3 (NHE3) was measured using this technical approach to study the molecular basis of NHE3 regulation by phosphoinositides.

A self-referencing ISE during whole-cell patch clamp recording was applied to study the regulation of NHE3 activity by phosphoinositides. PS120 fibroblast-like cells24, which lack the expression of endogenous plasma membrane NHEs, were used. NHE3 wild-type (NHE3-wt) or NHE3 mutants that do not bind phosphoinositides [tyrosine 501, arginine 503 and lysine 505 were substituted with alanine, Y501A/R503A/K505A (NHE3-YRK)] were stably expressed in PS120 fibroblast-like cells18.
The traces are presented in Figure 2. The cell was heldin a whole-cell configuration by a pipette for patch clamping, and the ISE was placed in front of the cell. In this position, the ISE recorded the concentration of free H+ close to the cell membrane (Figure 1A); then, the ISE (or patch clamp pipette) was manually moved laterally 50 &#181;m and recorded the concentration of free H+ in the solution (Figure 1B). The recorded voltage differences, corresponding to the two positions of the patch pipette with the cell in front of or far from the ISE, are representative of NHE3 activity (Figure 1C). The intra-pipette perfusion of apyrase (ATP-diphosphatase) decreases the intracellular concentration of ATP, subsequently decreasing the phosphoinositide content5. In agreement with previously reported studies5,18,19, the decreased phosphoinositide content mediated by apyrase treatment reduced NHE3 activity (~50%), which was recorded as a decrease in the voltage oscillation amplitude (Figure 2). The effect of apyrase (given at 5 units/ml) on NHE3 activity was completely reversed by the intra-pipette perfusion of apyrase in combination with 2 &#181;M PI(3,4,5)P3 (Figure 2 and 3A). These findings were supported by the NHE3 activity recorded in PS120 cells stably expressing NHE3 mutants (NHE3-YRK) that were unable to bind phosphatidylinositides18. The perfusion of apyrase alone or in combination with PIP3 did not affect the activity of NHE3 in NHE3-YRK mutants (Figure 3B). The activity of endogenous NHE3 was also inhibited by apyrase treatment in opossum kidney (OK) cells3. The intra-pipette perfusion of PI(3,4,5)P3 reversed the inhibition of NHE3 activity induced by apyrase in OK cells (Figure 3C). These findings suggest that the effect of PI(3,4,5)P3 on NHE3 activity is not cell type-specific.
Next, we aimed to test the specificity of PIP3 on regulating apyrase-mediated changes in NHE3 activity. We did this by assaying the effect on apyrase-dependent reduction on NHE3 activity after the perfusion of other phosphoinositides, such as PI(4,5)P2. PI(4,5)P2 (at 10 &#181;M) did not affect the decreased NHE3 activity mediated by apyrase (Figure 4A). Finally, the perfusion of AMP-PNP, a non-hydrolysable ATP analog, did not block theapyrase-dependent inhibition of NHE3 activity (Figure 4B). These findings suggest a role of PI(3,4,5)P3, but not PI(4,5)P2, in the regulation of NHE3 after ATP depletion. The identification of separate roles for PI(4,5)P2 and PI(3,4,5)P3 in NHE3 regulation was possible only because of the intercellular perfusion of phosphoinositides during ISE measurements combined with whole-cell patch clamp recording5. A specific action of PI(3,4,5)P3 rather than PI(4,5)P2 in the regulation of NHE3 activity is valuable to determine whether different intracellular phosphoinositides regulate the function of NHE3 differently.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56630/56630fig1.jpg" /> 
Figure 1. Schematic representation of H+ flux measurement and correlation between NHE activity and potential differences recorded by the pH microelectrode. 
Schematic representation of the recording setting. A. Diagram of the initial experimental setting. The cell is held in the whole-cell configuration by the patch clamp pipette and placed in front of the ISE. In this position, the ISE records the concentration of free H+ close to the cell membrane. B. The ISE (or patch clamp pipette) is manually moved laterally ~50 &#181;m and records the concentration of free H+ in the solution in this position. C. Graphic representation of voltage differences recorded using the ISE. The oscillations from position A (cell in front of the ISE) to B (cell far from the ISE) are representative of NHE activity. The software used registered the voltage differences recorded by the ISE and generated square wave perturbations (20 mV, 0.2 kHz) to monitor the cell membrane capacitance and quality of the patch clamp seal. <a href="https://cloudflare.jove.com/files/ftp_upload/56630/56630fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56630/56630fig2.jpg" /> 
Figure 2. Example of a recording set for NHE3 transporter activity in a single PS120 cell stably expressing NHE3-wt before and after perfusion with apyrase and PI(3,4,5)P3 (PIP3). 
After the initial recording, the cell was depleted of ATP by the intra-pipette perfusion of apyrase. NHE3 activity gradually decreased and was restored by PIP3 intra-pipette perfusion. Apyrase addition is indicated by the red bar, and PIP3 addition is indicated by the green bar. <a href="https://cloudflare.jove.com/files/ftp_upload/56630/56630fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56630/56630fig3.jpg" /> 
Figure 3. Effect of ATP depletion and PIP3 perfusion on NHE3 transporter activity in PS120 and OK cells. 
Effect of apyrase-mediated ATP depletion on NHE3 activity and reserving the effect of apyrase on NHE3 activity induced by PIP3 intra-pipette perfusion: A. PS120 cells expressing NHE3-wt or B. PS120 cells expressing NHE3-YRK, NHE3 mutants unable to bind phosphoinositides. C. OK cells expressing endogenous NHE3. Solid circles show average data from individual experiments (four experiments performed in identical conditions). Error bars represent the means &#177; standard errors (SE). *P &#60; 0.05 vs. control, ANOVA. $P&#60; 0.5, $$P &#60; 0.01 apyrase vs. apyrase + PIP3, ANOVA. <a href="https://cloudflare.jove.com/files/ftp_upload/56630/56630fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56630/56630fig4.jpg" /> 
Figure 4. Effect of PI(4,5)P2 (PIP2) and ANP-PNP perfusion on NHE3 activity after ATP depletion. 
Effect of A. PIP2 or B. ANP-PNP intra-pipette perfusion on the apyrase-dependent reduction of NHE3 activity in PS120 cells expressing NHE3-wt. Solid circles show average data from individual experiments (four experiments performed in identical conditions). Error bars represent the means &#177; SE. *P &#60; 0.05, **P &#60; 0.01 vs. control, ANOVA. <a href="https://cloudflare.jove.com/files/ftp_upload/56630/56630fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Application of Electrophysiology Measurement to Study the Activity of Electro-Neutral Transporters at JoVE.com

Application of Electrophysiology Measurement to Study the Activity of Electro-Neutral Transporters

This manuscript describes the applications of proton-selective electrodes and patch clamping methods to measure the activity of proton transport systems. These methods overcome some limitations of techniques commonly used to study proton transport activity, such as moderate sensitivity, time resolution and insufficient intracellular milieu control.

The transport of ions through cell membranes ensures the fine control of ion content within and outside the cell that is indispensable for cell survival. ...

application-electrophysiology-measurement-to-study-activity-electro

Nanyang Technological University

Research

JoVE Journal

Biology

6.9K Views.  Des Moines University. This manuscript describes the applications of proton-selective electrodes and patch clamping methods to measure the activity of proton transport systems. These methods overcome some limitations of techniques commonly used to study proton transport activity, such as moderate sensitivity, time resolution and insufficient intracellular milieu control.

Video: Application of Electrophysiology Measurement to Study the Activity of Electro-Neutral Transporters

Testing Visual Sensitivity to the Speed and Direction of Motion in Lizards

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of the visual system provide more empirical evidence for functional applications. Investigation into the sensory system provides information about the sensory capacity, learning and memory ability, and establishes known baseline behaviour in which to gauge deviations (Burghardt, 1977). However, unlike mammalian or avian systems, testing for learning and memory in a reptile species is difficult. Furthermore, using an operant paradigm as a psychophysical measure of sensory ability is likewise as difficult. Historically, reptilian species have responded poorly to conditioning trials because of issues related to motivation, physiology, metabolism, and basic biological characteristics. Here, I demonstrate an operant paradigm used a novel model lizard species, the Jacky dragon (Amphibolurus muricatus) and describe how to test peripheral sensitivity to salient speed and motion characteristics. This method uses an innovative approach to assessing learning and sensory capacity in lizards. I employ the use of random-dot kinematograms (RDKs) to measure sensitivity to speed, and manipulate the level of signal strength by changing the proportion of dots moving in a coherent direction. RDKs do not represent a biologically meaningful stimulus, engages the visual system, and is a classic psychophysical tool used to measure sensitivity in humans and other animals. Here, RDKs are displayed to lizards using three video playback systems. Lizards are to select the direction (left or right) in which they perceive dots to be moving. Selection of the appropriate direction is reinforced by biologically important prey stimuli, simulated by computer-animated invertebrates.

Testing visual sensitivity in lizards using an operant conditioning paradigm that employs video playback of random-dot kinematograms and computer-generated invertebrates.

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of ...

Computer-Generated Animal Model Stimuli

Communication between animals is diverse and complex. Animals may communicate using auditory, seismic, chemosensory, electrical, or visual signals. In particular, understanding the constraints on visual signal design for communication has been of great interest. Traditional methods for investigating animal interactions have used basic observational techniques, staged encounters, or physical manipulation of morphology. Less intrusive methods have tried to simulate conspecifics using crude playback tools, such as mirrors, still images, or models. As technology has become more advanced, video playback has emerged as another tool in which to examine visual communication (Rosenthal, 2000). However, to move one step further, the application of computer-animation now allows researchers to specifically isolate critical components necessary to elicit social responses from conspecifics, and manipulate these features to control interactions. Here, I provide detail on how to create an animation using the Jacky dragon as a model, but this process may be adaptable for other species. In building the animation, I elected to use Lightwave  3D to alter object morphology, add texture, install bones, and provide comparable weight shading that prevents exaggerated movement. The animation is then matched to select motor patterns to replicate critical movement features. Finally, the sequence must rendered into an individual clip for presentation. Although there are other adaptable techniques, this particular method had been demonstrated to be effective in eliciting both conspicuous and social responses in staged interactions.

Computer-generated stimuli using the Jacky dragon as a model.

Communication between animals is diverse and complex. Animals may communicate using auditory, seismic, chemosensory, electrical, or visual signals. In ...

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 ...

Semi-automated Optical Heartbeat Analysis of Small Hearts

We have developed a method for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts (Fink, et. al., 2009). Our Semi-automatic Optical Heartbeat Analysis (SOHA) uses a novel movement detection algorithm that is able to detect cardiac movements associated with individual contractile and relaxation events. The program provides a host of physiologically relevant readouts including systolic and diastolic intervals, heart rate, as well as qualitative and quantitative measures of heartbeat arrhythmicity. The program also calculates heart diameter measurements during both diastole and systole from which fractional shortening and fractional area changes are calculated. Output is provided as a digital file compatible with most spreadsheet programs. Measurements are made for every heartbeat in a record increasing the statistical power of the output. We demonstrate each of the steps where user input is required and show the application of our methodology to the analysis of heart function in all three genetically tractable heart models.

We have developed a Semi-automated Optical Heartbeat Analysis method (SOHA) for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts. We demonstrate the application of our methodology to the analysis of heart function in fruit fly and embryonic mouse hearts.

We have developed a method for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts (Fink, et. al., 2009). Our ...

Spinal Cord Electrophysiology

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

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

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

Application of MassSQUIRM for Quantitative Measurements of Lysine Demethylase Activity

Recently, epigenetic regulators have been discovered as key players in many different diseases 1-3. As a result, these enzymes are prime targets for small molecule studies and drug development 4. Many epigenetic regulators have only recently been discovered and are still in the process of being classified. Among these enzymes are lysine demethylases which remove methyl groups from lysines on histones and other proteins. Due to the novel nature of this class of enzymes, few assays have been developed to study their activity. This has been a road block to both the classification and high throughput study of histone demethylases. Currently, very few demethylase assays exist. Those that do exist tend to be qualitative in nature and cannot simultaneously discern between the different lysine methylation states (un-, mono-, di- and tri-). Mass spectrometry is commonly used to determine demethylase activity but current mass spectrometric assays do not address whether differentially methylated peptides ionize differently. Differential ionization of methylated peptides makes comparing methylation states difficult and certainly not quantitative (Figure 1A). Thus available assays are not optimized for the comprehensive analysis of demethylase activity.
Here we describe a method called MassSQUIRM (mass spectrometric quantitation using isotopic reductive methylation) that is based on reductive methylation of amine groups with deuterated formaldehyde to force all lysines to be di-methylated, thus making them essentially the same chemical species and therefore ionize the same (Figure 1B). The only chemical difference following the reductive methylation is hydrogen and deuterium, which does not affect MALDI ionization efficiencies. The MassSQUIRM assay is specific for demethylase reaction products with un-, mono- or di-methylated lysines. The assay is also applicable to lysine methyltransferases giving the same reaction products. Here, we use a combination of reductive methylation chemistry and MALDI mass spectrometry to measure the activity of LSD1, a lysine demethylase capable of removing di- and mono-methyl groups, on a synthetic peptide substrate 5. This assay is simple and easily amenable to any lab with access to a MALDI mass spectrometer in lab or through a proteomics facility. The assay has ~8-fold dynamic range and is readily scalable to plate format 5.

We present a method for using MALDI mass spectrometry and reductive methylation chemistry to quantify changes in lysine methylation.

Recently, epigenetic regulators have been discovered as key players in many different diseases 1-3. As a result, these enzymes are prime targets for small ...

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.

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 ...

Reconstruction of 3-Dimensional Histology Volume and its Application to Study Mouse Mammary Glands

Histology volume reconstruction facilitates the study of 3D shape and volume change of an organ at the level of macrostructures made up of cells. It can also be used to investigate and validate novel techniques and algorithms in volumetric medical imaging and therapies. Creating 3D high-resolution atlases of different organs1,2,3 is another application of histology volume reconstruction. This provides a resource for investigating tissue structures and the spatial relationship between various cellular features. We present an image registration approach for histology volume reconstruction, which uses a set of optical blockface images. The reconstructed histology volume represents a reliable shape of the processed specimen with no propagated post-processing registration error. The Hematoxylin and Eosin (H&#38;E) stained sections of two mouse mammary glands were registered to their corresponding blockface images using boundary points extracted from the edges of the specimen in histology and blockface images. The accuracy of the registration was visually evaluated. The alignment of the macrostructures of the mammary glands was also visually assessed at high resolution.
This study delineates the different steps of this image registration pipeline, ranging from excision of the mammary gland through to 3D histology volume reconstruction. While 2D histology images reveal the structural differences between pairs of sections, 3D histology volume provides the ability to visualize the differences in shape and volume of the mammary glands.

We present an image registration approach for 3-dimensional (3D) histology volume reconstruction, which facilitates the study of the changes of an organ at the level of macrostructures made up of cells . Using this approach, we studied the 3D changes between wild-type and Igfbp7-null mammary glands.

Histology volume reconstruction facilitates the study of 3D shape and volume change of an organ at the level of macrostructures made up of cells. It can ...

High-throughput Image Analysis of Tumor Spheroids: A User-friendly Software Application to Measure the Size of Spheroids Automatically and Accurately

The increasing number of applications of three-dimensional (3D) tumor spheroids as an in vitro model for drug discovery requires their adaptation to large-scale screening formats in every step of a drug screen, including large-scale image analysis. Currently there is no ready-to-use and free image analysis software to meet this large-scale format. Most existing methods involve manually drawing the length and width of the imaged 3D spheroids, which is a tedious and time-consuming process. This study presents a high-throughput image analysis software application &#8211; SpheroidSizer, which measures the major and minor axial length of the imaged 3D tumor spheroids automatically and accurately; calculates the volume of each individual 3D tumor spheroid; then outputs the results in two different forms in spreadsheets for easy manipulations in the subsequent data analysis. The main advantage of this software is its powerful image analysis application that is adapted for large numbers of images. It provides high-throughput computation and quality-control workflow. The estimated time to process 1,000 images is about 15 min&#160;on a minimally configured laptop, or around 1 min&#160;on a multi-core performance workstation. The graphical user interface (GUI) is also designed for easy quality control, and users can manually override the computer results. The key method used in this software is adapted from the active contour algorithm, also known as Snakes, which is especially suitable for images with uneven illumination and noisy background that often plagues automated imaging processing in high-throughput screens. The complimentary &#8220;Manual Initialize&#8221; and &#8220;Hand Draw&#8221; tools provide the flexibility to SpheroidSizer in dealing with various types of spheroids and diverse quality images. This high-throughput image analysis software remarkably reduces labor and speeds up the analysis process. Implementing this software is beneficial for 3D tumor spheroids to become a routine in vitro model for drug screens in industry and academia.

We present a high-throughput image analysis software application to measure the size of three-dimensional tumor spheroids imaged with bright-field microscopy. This application provides a fast and effective way to examine the effects of therapeutic drugs on spheroids, which is beneficial for researchers who wish to use spheroids in drug screens.

The increasing number of applications of three-dimensional (3D) tumor spheroids as an in vitro model for drug discovery requires their adaptation to ...

Measurement of Larval Activity in the Drosophila Activity Monitor

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This protocol repurposes an instrument often used to measure adult activity, the TriKinetics Drosophila activity monitor (MB5 Multi-Beam Activity Monitor) to study larval activity. The instrument can monitor the movements of animals in 16 individual 8 cm glass assay tubes, using 17 infrared detection beams per tube. Logging software automatically saves data to a computer, recording parameters such as number of moves, times sensors were triggered, and animals&#8217; positions within the tubes. The data can then be analyzed to represent overall locomotion and/or position preference as well as other measurements. All data are easily accessible and compatible with basic graphing and data manipulation software. This protocol will discuss how to use the apparatus, how to operate the software and how to run a larval activity assay from start to finish.

Measurement of Larval Activity in the Drosophila Activity Monitor

This report describes a method for measuring Drosophila larval activity using the TriKinetics Drosophila Activity Monitor. The device employs infrared beams to detect movements of up to 16 individual animals. Data can be analyzed to represent motion parameters including rates and the positions of the animals within the assay chambers.

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This ...

Application of Electrophysiology Measurement to Study the Activity of Electro-Neutral Transporters

Summary

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Chapters in this video

Testing Visual Sensitivity to the Speed and Direction of Motion in Lizards

Computer-Generated Animal Model Stimuli

Applying Microfluidics to Electrophysiology

Semi-automated Optical Heartbeat Analysis of Small Hearts

Spinal Cord Electrophysiology

Application of MassSQUIRM for Quantitative Measurements of Lysine Demethylase Activity

Introduction to Solid Supported Membrane Based Electrophysiology

Reconstruction of 3-Dimensional Histology Volume and its Application to Study Mouse Mammary Glands

High-throughput Image Analysis of Tumor Spheroids: A User-friendly Software Application to Measure the Size of Spheroids Automatically and Accurately

Measurement of Larval Activity in the Drosophila Activity Monitor