The authors would like to thank Glen Slocum (Medical College of Wisconsin) and Colleen A. Lavin (Nikon Instruments Inc) for excellent technical assistance with microscopy experiments. This study was supported by the National Institutes of Health grants R01 HL108880 (to AS), R01 DK095029 (to OPo) and K99 HL116603 (to TSP), National Kidney Foundation IG1724 (to TSP), American Heart Association 13GRNT16220002 (to OPo) and the Ben J. Lipps Research Fellowship from the American Society of Nephrology (to DVI).

The experimental procedures described below were approved by the Institutional Animal Care and Use Committee at the Medical College of Wisconsin and University of Texas Health Science Center at Houston and were in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Figure 1 demonstrates main steps of the tissue isolation and processing procedure. Briefly, kidneys from PCK or SD rats are used for manual isolation of epithelial monolayers of collecting ducts either from healthy non-dialed tubules or cysts. Here we studied kidneys from 4-16 weeks old PCK rats10,13.
1. Isolation of Renal Cysts and Connecting Tubules (CNT)/Collecting Ducts (CD) Segments
<ol>
	<li>Anesthetize experimental animal with isoflurane (5% induction, 1.5 to 2.5% maintenance)/medical grade O2 or another approved method. Animals must be continually monitored to ensure adequate level of anesthesia. Stable respiratory rate and toe pinch reaction are used to confirm proper anesthesia.</li>
	<li>Perform laparotomy and flush the kidneys with phosphate buffered saline (PBS) through the abdominal aorta17.
	<ol>
		<li>Prepare a polyethylene tubing catheter (PE50) and connect it to a syringe pump filled with PBS. Cut the skin and abdominal wall along the linea alba, then make a transverse abdominal incision across the abdomen from side to side and shift the abdominal organs&#160;with cotton swabs to get access to the descending aorta with branching mesenteric and celiac arteries. Moisten the internal organs with warm saline during further procedures to prevent their dryness.</li>
		<li>Place one ligature (#1) around the mesenteric and celiac arteries and another ligature (#2) around the aorta under the diaphragm (do not tie). Gently separate a. abdominalis by blunt dissecting connective tissues around it with thin forceps and place two ligatures ~3 mm below the left renal artery (#3) and above the iliac arteries bifurcation (#4).</li>
		<li>Tie ligature #4 and attach a vessel clamp around the aorta between the left renal artery and ligature #3. Make an incision between the clamp and ligature #4 to catheterize the aorta, use ligature #3 to firmly fix the catheter.</li>
		<li>Release the clamp and make sure that the blood pulse is visible in the catheter to ensure proper installation. Start perfusion at a rate of 6 ml/min, tie ligatures #1 and #2, and cut the left renal vein. Continue flushing for 1-2 min until the organs are completely blanched.</li>
		<li>Cut renal blood vessels, urethra and surrounding connective and adipose tissues with scissors to collect the kidneys. Then make a short tear in kidney capsule and peel the kidneys to decapsulate them. Place the kidneys into ice-cold PBS. Euthanasia is confirmed by a thoracotomy.</li>
	</ol>
	</li>
	<li>Prepare 5 x 5 mm cover glass chips coated with Poly-L-Lysine. Cut a cover glass with a diamond pencil and place approximately 20 &#181;l of 0.01% sterile filtered Poly-L-Lysine solution onto each glass chip. Prepare saline and two pairs of watchmaker forceps for dissection.</li>
	<li>Cut the kidneys with a razor blade along the frontal plane into slices of ~1-2 mm thickness. Place one of the slices under a stereomicroscope. Isolation of tissues should be done in ice-cold saline.</li>
	<li>Locate cysts under a stereomicroscope as round-shape cavities (Figure 2A); their walls often contain prominent network of proliferated blood vessels. Using fine-tipped forceps dissect the internal epithelial layer of a cyst as thin as possible to obtain a monolayer area. Attach it to a glass chip covered with Poly-L-Lysine. Sticky Poly-L-Lysine surface allows the researcher to firmly position the internal cyst layer on glass. Expose the cyst with the apical side up to provide access for the pipettes (Figure 2B).</li>
	<li>Use PCK or SD rat kidney central slices containing papilla for isolation of non-dilated CNT/CCDsegments18-21. 
	NOTE: Papillary tissues are more durable than cortex and by holding papillary tubular bands with forceps and tearing along radial axis the slice can be cleaved into thin sectors. Individual tubules are visible and can be pulled out to be placed on cover glass chips.</li>
	<li>Identify the CNT/CCD segments by bifurcations, higher transparency than proximal tubules and large prominently visible cells (Figure 2C). Place the cover glass chip with attached tubules under a microscope equipped with micromanipulators suitable for driving micropipettes. Using sharp micropipettes split-open tubules and attach their edges to the glass to make the apical surface accessible (Figure 2D).</li>
</ol>
2. Single Channel Patch-clamp Electrophysiology
<ol>
	<li>Fill the patch-clamp chamber with bath solution and transfer the cover glass chip with isolated tissues to the chamber. Ensure that the patch-clamp micropipettes have resistance of 7-10 M&#937; for reliable on-cell (cell-attached) measurements.</li>
	<li>For the cell-attached measurements, set the amplifier gain ratio to 20x and low-pass the currents at 300 Hz by an eight-pole Bessel filter. For ENaC channels monitoring, use a bath solution, in mM: 150 NaCl, 1 CaCl2, 2 MgCl2, 10 HEPES (pH 7.4); pipette: 140 LiCl, 2 MgCl2 and 10 HEPES (pH 7.4).</li>
	<li>Conduct a conventional patch-clamp experiment in a cell-attached mode10. Select a cell in the epithelial monolayer and approach the pipette to the apical membrane. Form a high-resistance seal between a pipette and cell membrane by applying gentle suction. Once a high resistant gigaOhm seal is formed, a gap-free protocol at a holding potential should be started for monitoring activity of the channel of interest.</li>
	<li>Store gap-free single channel current data from gigaOhm seals for subsequent analysis. Analyze the channel events using a software package generally supplied with patch-clamp setups18. Calculate channel activity as NPo where N refers to the number of active channels in the patch and Po is average open probability of the channels. 
	NOTE: Use isolated cysts or tubules in patch-clamp experiments for no more than 30 min.</li>
</ol>
3. Ratiometric Epifluorescence Measurements of Intracellular Calcium Concentration in the Epithelial Cells
<ol>
	<li>Use 5 mM Fura-2AM dissolved in DMSO. Aliquot stock solution into individual 500 &#181;l conical tubes (approximately 10 &#181;l). Protect from light and store in the freezer at -20 &#730;C for up to six months.</li>
	<li>Incubate isolated cysts and split-open tubules for 30-40 min in PSS (in mM: 145 NaCl, 4.5 KCl, 2 MgCl2, 2 CaCl2 and 10 HEPES at pH 7.35) containing 5 &#181;M Fura-2 AM dye and 0.05% pluronic acid to help disperse the acetoxymethyl esters. For that, place tissues in the 3.5 cm dish containing loading cocktail, protect from light and incubate on a slow shaker at room temperature.</li>
	<li>Change Fura-2 AM containing media to clean PSS after incubation and place the tissue under a microscope to further perform epifluorescence imaging.</li>
	<li>Turn on CCD camera and stable light source (monochromator system) equipped with filter wheel (each filter position can be associated with its own attenuation level, selected every time the filter is called).</li>
	<li>Find the tissue in bright field. Switch to detection of the fluorescence signal and adjust the intensity of the light source using neutral density filters and lamp power to avoid saturation of the signal.</li>
	<li>Monitor Fura-2 AM fluorescence in the tissue sample with ratiometric excitation at 340/380 nm with frequency of 0.125 Hz or higher. Use a 40&#215;/NA 1.3 or similar objective lens for proper resolution image.</li>
	<li>For image processing and calculations import the image sequence. Make sure to split the channels and use a hyperstack grayscale mode. Select several regions of interest (single cyst cells or selected areas of cystic tissue) and calculate intensity values for each channel (340 and 380 nm) into preferred data analysis software; subtract background intensity values from each data point.</li>
	<li>For each time point calculate the ratio of intensities of the Fura-2 AM 340 to 380 channels. Plot scatter/line point-time changes of Ca2+ transient for each region and calculate mean/SE values.</li>
</ol>

The authors have nothing to disclose.

We described here applications of conventional patch-clamp technique and epifluorescence calcium imaging to cystic epithelial monolayers derived from a murine genetic model of ARPKD. The protocol consist of three steps, of which the most attention should be paid to the isolation of the cysts (step 1.5 of the protocols section) and to the electrophysiological studies. These key procedures require extensive training and patience, and the reader should not be frustrated at once.
First of all, the...

Ion channels play a significant role in many physiological functions, including cell growth and differentiation. Autosomal dominant and recessive polycystic kidney diseases (ADPKD and ARPKD, respectively) are genetic disorders characterized by the development of renal fluid-filled cysts of the tubular epithelial cell origin. ADPKD is caused by mutations of PKD1 or PKD2 genes encoding polycystins 1 and 2, membrane proteins involved in the regulation of cell proliferation and differentiation. PKD2 by itself or as a complex with PKD1 also function as a Ca2+-permeable cation channel1. Mutations of the PKHD1 gene encoding fibrocystin (a cilia-associated receptor-like protein involved in the tubulogenesis and/or maintenance of polarity of epithelium) are the genetic impetus of ARPKD2. Cyst growth is a complex phenomenon accompanied with disturbed proliferation3,4, angiogenesis5, dedifferentiation and loss of polarity of tubular cells6-8.
Defective reabsorption and augmented secretion in cystic epithelium contribute to fluid accumulation in the lumen and cyst expansion9,10. Impaired flow-dependent [Ca2+]i&#160;signaling has been also linked to cystogenesis during PKD11-15.
Here, we describe a method suitable for patch-clamp measurements of single channel activity and intracellular Ca2+ levels in cystic epithelial monolayers isolated from PCK rats. This method was successfully applied by us to characterize of activity of the epithelial Na+ channel (ENaC)10 and [Ca2+]i-dependent processes induced by Ca2+-permeable TRPV4 and purinergic signaling cascade13.
In these studies we used PCK rats, a model of ARPKD caused by a spontaneous mutation in the PKHD1 gene. The PCK strain was originally derived from Sprague-Dawley (SD) rats16 thereby SD rats are used as an appropriate control for comparison with the PCK strain. As a result, both SD rat nephron segments and non-dilated collecting ducts isolated from same PCK rats can serve as two different comparison groups for experiments on cystic epithelium.

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			<td height="21" style="height:21px;width:283px;">Fura-2 AM</td>
			<td style="width:219px;">Life Technologies</td>
			<td style="width:183px;">F-14185</td>
			<td style="width:333px;"></td>
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			<td style="width:183px;">21091</td>
			<td style="width:333px;"></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Poly-L-lysine</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td style="width:183px;">P4707</td>
			<td style="width:333px;"></td>
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			<td height="21" style="height:21px;width:283px;">Pluronic acid</td>
			<td style="width:219px;">Sigma-Aldrich</td>
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			<td style="width:333px;">solution</td>
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			<td style="width:219px;">Boekel Scientific</td>
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			<td style="width:333px;">with integrated shutter/filter wheel driver</td>
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			<td height="21" style="height:21px;width:283px;">Neutral density filters</td>
			<td style="width:219px;">Nikon</td>
			<td style="width:183px;"></td>
			<td style="width:333px;">ND4, ND8</td>
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			<td height="21" style="height:21px;width:283px;">Objective</td>
			<td style="width:219px;">Nikon</td>
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			<td style="width:333px;">40/1.3 DIC WD 0.22&#160;&#160; oil</td>
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			<td style="width:333px;"></td>
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			<td height="23" style="height:23px;width:283px;">Nikon&#160; microscope (inverted)</td>
			<td style="width:219px;">Nikon</td>
			<td style="width:183px;">Nikon Eclipse TE2000-S</td>
			<td style="width:333px;"></td>
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			<td height="21" style="height:21px;width:283px;">Cover Glass</td>
			<td style="width:219px;">Thermo Scientific</td>
			<td style="width:183px;">6661B52</td>
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			<td height="21" style="height:21px;width:283px;">Diamond pencil</td>
			<td style="width:219px;">Fisher Scientific</td>
			<td style="width:183px;">22268912</td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Image acquisition software</td>
			<td style="width:219px;">Nikon</td>
			<td style="width:183px;">Nikon NIS-Elements&#160;</td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:283px;">Image analysis software</td>
			<td style="width:219px;">ImageJ</td>
			<td style="width:183px;">http://imagej.nih.gov/</td>
			<td style="width:333px;">ND Utility plugin allows to import images in the native Nikon Instruments .nd2 format</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Recording/perfusion chamber</td>
			<td style="width:219px;">Warner Instruments</td>
			<td style="width:183px;">RC-26</td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;"></td>
			<td style="width:219px;"></td>
			<td style="width:183px;"></td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Patch Clamp amplifier</td>
			<td style="width:219px;">Molecular Devices</td>
			<td style="width:183px;">MultiClamp 700B</td>
			<td style="width:333px;"></td>
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			<td height="21" style="height:21px;width:283px;">Data Acquisition System</td>
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			<td style="width:183px;">Digidata 1440A</td>
			<td style="width:333px;">Axon Digidata&#174; System</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Low Pass Filter</td>
			<td style="width:219px;">Warner Instruments</td>
			<td style="width:183px;">LPF-8</td>
			<td style="width:333px;">8 pole Bessel</td>
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			<td height="21" style="height:21px;width:283px;">Borosilicate glass capillaries</td>
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			<td style="width:183px;">1B150F-4</td>
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			<td style="width:219px;">Sutter Instrument Co</td>
			<td style="width:183px;">P-97</td>
			<td style="width:333px;">Flaming/Brown type micropipette puller</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Microforge</td>
			<td style="width:219px;">Narishige</td>
			<td style="width:183px;">MF-830</td>
			<td style="width:333px;">Japan</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Motorized Micromanipulator</td>
			<td style="width:219px;">Sutter Instrument Co</td>
			<td style="width:183px;">MP-225</td>
			<td style="width:333px;"></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Inverted microscope</td>
			<td style="width:219px;">Nikon</td>
			<td style="width:183px;">Eclipse Ti</td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Microvibration isolation table</td>
			<td style="width:219px;">TMC</td>
			<td style="width:183px;"></td>
			<td style="width:333px;">equipped with Faraday cage</td>
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			<td height="21" style="height:21px;width:283px;">Multichannel valve perfusion system</td>
			<td style="width:219px;">AutoMake Scientific</td>
			<td style="width:183px;">Valve Bank II</td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Recording/perfusion chamber</td>
			<td style="width:219px;">Warner Instruments</td>
			<td style="width:183px;">RC-26</td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Software</td>
			<td style="width:219px;">Molecular Devices</td>
			<td style="width:183px;">pClamp 10 . 2</td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;"></td>
			<td style="width:219px;"></td>
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		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Temperature controlled surgical table&#160;</td>
			<td style="width:219px;">MCW core</td>
			<td style="width:183px;"></td>
			<td style="width:333px;">for rodents</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Binocular stereomicroscope</td>
			<td style="width:219px;">Nikon</td>
			<td style="width:183px;">SMZ745</td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Syringe pump-based perfusion system</td>
			<td style="width:219px;">Harvard Apparatus</td>
			<td style="width:183px;"></td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">polyethylene tubing</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td style="width:183px;">PE50</td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Isofluorane anesthesia</td>
			<td style="width:219px;"><a href="http://www.vetequip.com/">http://www.vetequip.com/</a></td>
			<td style="width:183px;">911103</td>
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			<td style="width:219px;"></td>
			<td style="width:183px;"></td>
			<td style="width:333px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Other basic reagents</td>
			<td style="width:219px;">Sigma-Aldrich</td>
			<td style="width:183px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

implementing patch clamp and live fluorescence microscopy to monitor functional properties of freshly isolated pkd epithelium

Cyst initiation and expansion during polycystic kidney disease is a complex process characterized by abnormalities in tubular cell proliferation, luminal fluid accumulation and extracellular matrix formation. Activity of ion channels and intracellular calcium signaling are key physiologic parameters which determine functions of tubular epithelium. We developed a method suitable for real-time observation of ion channels activity with patch-clamp technique and registration of intracellular Ca2+ level in epithelial monolayers freshly isolated from renal cysts. PCK rats, a genetic model of autosomal recessive polycystic kidney disease (ARPKD), were used here for ex vivo analysis of ion channels and calcium flux. Described here is a detailed step-by-step procedure designed to isolate cystic monolayers and non-dilated tubules from PCK or normal Sprague Dawley (SD) rats, and monitor single channel activity and intracellular Ca2+ dynamics. This method does not require enzymatic processing and allows analysis in a native setting of freshly isolated epithelial monolayer. Moreover, this technique is very sensitive to intracellular calcium changes and generates high resolution images for precise measurements. Finally, isolated cystic epithelium can be further used for staining with antibodies or dyes, preparation of primary cultures and purification for various biochemical assays.

Potential ENaC involvement in cystogenesis has been demonstrated by several studies that observed disrupted epidermal growth factor (EGF) signaling in PKD progression22-25 and abnormal sodium reabsorption in ARPKD murine models and tissue cultures26-28. For example, Veizis et al. showed that amiloride-sensitive Na+&#160;absorption is decreased in CD cells from the non-orthologous BPK mouse model of ARPKD29. We recently demonstrated that impaired sodium and water reabso...

Watch this Scientific Journal Video about Implementing Patch Clamp and Live Fluorescence Microscopy to Monitor Functional Properties of Freshly Isolated PKD Epithelium at JoVE.com

Implementing Patch Clamp and Live Fluorescence Microscopy to Monitor Functional Properties of Freshly Isolated PKD Epithelium

Ion channels expressed in renal tubular epithelium play a significant role in the pathology of polycystic kidney disease. Here we describe experimental protocols used to perform patch-clamp analysis and intracellular calcium level measurements in cystic epithelium freshly isolated from rodent kidneys.

Cyst initiation and expansion during polycystic kidney disease is a complex process characterized by abnormalities in tubular cell proliferation, luminal ...

implementing-patch-clamp-live-fluorescence-microscopy-to-monitor

Research

JoVE Journal

Biology

9.7K Views.  Medical College of Wisconsin. Ion channels expressed in renal tubular epithelium play a significant role in the pathology of polycystic kidney disease. Here we describe experimental protocols used to perform patch-clamp analysis and intracellular calcium level measurements in cystic epithelium freshly isolated from rodent kidneys.

Video: Implementing Patch Clamp and Live Fluorescence Microscopy to Monitor Functional Properties of Freshly Isolated PKD Epithelium

Methods for Patch Clamp Capacitance Recordings from the Calyx

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal. Electrical recordings from the presynaptic terminal allow the measurement of action potentials, calcium channel currents, vesicle fusion (exocytosis) and subsequent membrane uptake (endocytosis). The fusion of vesicles containing neurotransmitter causes the vesicle membrane to be added to the cell membrane of the calyx. This increase in the amount of cell membrane is measured as an increase in capacitance. The subsequent reduction in capacitance indicates endocytosis, the process of membrane uptake or removal from the calyx membrane. Endocytosis, is necessary to maintain the structure of the calyx and it is also necessary to form vesicles that will be filled with neurotransmitter for future exocytosis events. Capacitance recordings at the calyx of Held have made it possible to directly and rapidly measure vesicular release and subsequent endocytosis in a mammalian CNS nerve terminal. In addition, the corresponding postsynaptic activity can be simultaneously measured by using paired recordings. Thus a complete picture of the presynaptic and postsynaptic electrical activity at a central nervous system synapse is achievable using this preparation. Here, the methods for slice preparation, morphological features for identification of calyces of Held, basic patch clamping techniques, and examples of capacitance recordings to measure exocytosis and endocytosis are presented.

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal.

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal. ...

Patch Clamp Recording of Ion Channels Expressed in  Xenopus Oocytes

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological measurement of single or multiple ion channels in cells. This technique can be applied to ion channels in both their native environment and expressed in heterologous cells, such as oocytes harvested from the African clawed frog, Xenopus laevis. Here, we describe the well-established technique of patch clamp recording from Xenopus oocytes. This technique is used to measure the properties of expressed ion channels either in populations (macropatch) or individually (single-channel recording). We focus on techniques to maximize the quality of oocyte preparation and seal generation. With all factors optimized, this technique gives a probability of successful seal generation over 90 percent. The process may be optimized differently by every researcher based on the factors he or she finds most important, and we present the approach that have lead to the greatest success in our hands.

This is intended as an introduction to patch clamp recording from Xenopus laevis oocytes. It covers vitelline membrane removal, formation of a gigaohm seal (gigaseal), and the optional conversion of the patch to the outside-out topology.

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological ...

Pressure-polishing Pipettes for Improved Patch-clamp Recording

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording. We first developed this method for fabricating pipettes suitable for recording from small (&lt;3 m) neuronal cell bodies. The basic principal is similar to glass-blowing and combines air pressure and heat to modify the shape of patch pipettes prepared by a conventional micropipette puller. It can be applied to so-called soft (soda lime) and hard (borosilicate) glasses. Generally speaking, pressure polishing can reduce pipette resistance by 25% without decreasing the diameter of the tip opening (Goodman and Lockery, 2000). It can be applied to virtually any type of glass and requires only the addition of a high-pressure valve and fitting to a microforge. This technique is essential for recording from ultrasmall cells (&lt;5 m) and can also improve single-channel recording by minimizing pipette resistance. The blunt shape is also useful for perforated-patch clamp recording since this tip shape results in a larger membrane bleb available for perforation.

This is a guide to modifying the shape of glass micropipettes. Specifically, by using heat and air pressure the taper is widened without increasing the tip opening, leading to lower pipette resistance. This is critical to obtain low noise recordings of small cells but is useful in many applications.

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording.  We first developed this method for fabricating pipettes suitable for ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes &gt;10 &mu;m in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions. Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.

Reconstituting functional membrane proteins into giant liposomes of defined composition is a powerful approach when combined with patch-clamp electrophysiology. However, conventional giant liposome production may be incompatible with protein stability. We describe protocols for producing giant liposomes from pure lipids or small liposomes containing ion channels.

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify ...

One-channel Cell-attached Patch-clamp Recording

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate depends on properties intrinsic to each channel protein as well as the mechanism by which it is generated and controlled and represents an important area of current research. Information about the operational dynamics of ion channel proteins can be obtained by observing long stretches of current produced by a single molecule. Described here is a protocol for obtaining one-channel cell-attached patch-clamp current recordings for a ligand gated ion channel, the NMDA receptor, expressed heterologously in HEK293 cells or natively in cortical neurons. Also provided are instructions on how to adapt the method to other ion channels of interest by presenting the example of the mechano-sensitive channel PIEZO1. This method can provide data regarding the channel&#8217;s conductance properties and the temporal sequence of open-closed conformations that make up the channel&#8217;s activation mechanism, thus helping to understand their functions in health and disease.

Described here is a procedure for obtaining long stretches of current recording from one ion channel with the cell-attached patch-clamp technique. This method allows for observing, in real time, the pattern of open-close channel conformations that underlie the biological signal. These data inform about channel properties in undisturbed biological membranes.

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate ...

Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow GUVs must be modified in order to form GUVs containing functional transmembrane proteins. This article describes two dehydration-rehydration methods &#8212; electroformation and gel-assisted swelling &#8212; to form GUVs containing the voltage-gated potassium channel, KvAP. In both methods, a solution of protein-containing small unilamellar vesicles is partially dehydrated to form a stack of membranes, which is then allowed to swell in a rehydration buffer. For the electroformation method, the film is deposited on platinum electrodes so that an AC field can be applied during film rehydration. In contrast, the gel-assisted swelling method uses an agarose gel substrate to enhance film rehydration. Both methods can produce GUVs in low (e.g., 5 mM) and physiological (e.g., 100 mM) salt concentrations. The resulting GUVs are characterized via fluorescence microscopy, and the function of reconstituted channels measured using the inside-out patch-clamp configuration. While swelling in the presence of an alternating electric field (electroformation) gives a high yield of defect-free GUVs, the gel-assisted swelling method produces a more homogeneous protein distribution and requires no special equipment.

The reconstitution of the transmembrane protein, KvAP, into giant unilamellar vesicles (GUVs) is demonstrated for two dehydration-rehydration methods &#8212; electroformation, and gel-assisted swelling. In both methods, small unilamellar vesicles containing the protein are fused together to form GUVs that can then be studied by fluorescence microscopy and patch-clamp electrophysiology.

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow ...

Single-channel Analysis and Calcium Imaging in the Podocytes of the Freshly Isolated Glomeruli

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells in the pathogenesis of various renal diseases has been established since podocyte injury leads to proteinuria and foot process effacement. It was previously reported that various endogenous agents may cause a dramatic overload in intracellular Ca2+ concentration in podocytes, presumably leading to albuminuria, and this likely occurs via calcium-conducting ion channels. Therefore, it appeared important to study calcium handling in the podocytes both under normal conditions and in various pathological states. However, available experimental approaches have remained somewhat limited to cultured and transfected cells. Although they represent a good basic model for such studies, they are essentially extracted from the native environment of the glomerulus. Here we describe the methodology of studying podocytes as a part of the freshly isolated whole glomerulus. This preparation retains the functional potential of the podocytes, which are still attached to the capillaries; therefore, podocytes remain in the environment that conserves the major parts of the glomeruli filtration apparatus. The present manuscript elaborates on two experimental approaches that allow 1) real-time detection of calcium concentration changes with the help of ratiometric confocal fluorescence microscopy, and 2) the recording of the single ion channels activity in the podocytes of the freshly isolated glomeruli. These methodologies utilize the advantages of the native environment of the glomerulus that enable researchers to resolve acute changes in the intracellular calcium handling in response to applications of various agents, measure basal concentration of calcium within the cells (for instance, to evaluate disease progression), and assess and manipulate calcium conductance at the level of single ion channels.

Changes in the intracellular calcium levels in the podocytes are one of the most important means to control the filtration function of glomeruli. Here we explain a high-throughput approach that allows detection of real-time calcium handling and single ion channels activity in the podocytes of the freshly isolated glomeruli.

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells ...