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. Gregory Blass is acknowledged for critical proofreading of the manuscript. This research was supported by the National Institutes of Health grant HL108880 and American Diabetes Association grant 1-15-BS-172 (to AS), and the Ben J. Lipps Research Fellowship from the American Society of Nephrology (to DVI).

Animal use and welfare should adhere to the NIH Guide for the Care and Use of Laboratory Animals following protocols reviewed and approved by the Institutional Animal Care and Use Committee (IACUC).
1. Kidney Flush
<ol>
	<li>Use 8 to 12 week old male rat (suggested is a Sprague Dawley strain, however other strains of different age and gender can be used with appropriate changes).</li>
	<li>Anesthetize the animal according to the procedure allowed by IACUC protocol; monitor depth of anesthesia and inspect the animal. The detailed description of the surgery to be performed in 1.3 - 1.8 can be found in Ilatovskaya et al.7</li>
	<li>After proper anesthesia, place the animal on a temperature-controlled surgical table, make a midline incision of the abdomen (up to 3 inches in length), and uncover the vena cava and the aorta.</li>
	<li>Insert ligature around the celiac and superior mesenteric arteries and the abdominal aorta above those; do not ligate.</li>
	<li>Blunt dissect the abdominal aorta below the renal arteries, and place two ligatures around it, but do not ligate, then clamp the aorta above the ligatures and tie the lower thread.</li>
	<li>Catheterize the aorta with a polyethylene PE50 tubing (attached to a syringe pump filled with PBS) below the clamp and fix the catheter with the second ligature; remove the clamp, turn on the pump, and ligate the aorta and mesenteric with celiac arteries. Quickly make incision in renal vein to relieve the pressure.</li>
	<li>Infuse the aorta with the pre-chilled PBS for 2 or 3 min at a rate of 6 ml/min.</li>
	<li>Stop perfusion, excise and decapsulate7 the kidneys, and put them on ice in the PBS solution. Euthanize the animal according to protocol approved by IACUC.</li>
</ol>
2. Isolation of the Rat Glomeruli
<ol>
	<li>Prepare 30 ml of fresh solution of 5% BSA in RPMI 1640.</li>
	<li>Using a razor blade and scissors, isolate the cortex of both kidneys, and then mince until homogenous. This procedure has been described earlier7.</li>
	<li>Push the tissue minced during the previous step through a 100 mesh stainless steel sieve (pre-soaked in 5% BSA/RPMI solution) using a spatula. Collect the flow-through and by gravity force allow the flow-through to pass through a 140 mesh sieve.</li>
	<li>Filter the flow-through collected from the 140 mesh sieve using a pre-soaked 200 mesh sieve, discard the filtrate, and wash the top of the sieve with 10 - 15 ml of the prepared BSA/RPMI solution to collect the glomeruli that sediment on the sieve.</li>
	<li>Put the BSA/RPMI solution containing glomeruli on ice in a 15 ml tube and let the glomeruli sediment at the bottom of the tube for up to 20 min. The glomeruli concentrate on the bottom of the tube will be clearly seen. Remove the excess solution, leaving approximately 2 ml in the tube.</li>
</ol>
3. Single-channel Patch-clamp Electrophysiology
<ol>
	<li>Prepare 5 x 5 mm cover glass chips by coating them with MW 70,000 - 150,000 poly-&#671;-lysine, and let dry. Use approximately 30 &#181;l of 0.01% sterile filtered solution in water per cover glass.</li>
	<li>Warm up the experimental solutions to RT and fill the patch-clamp chamber and pipette. For TRPC channels monitoring, use a bath solution, in mM: 126 NaCl, 1 CaCl2, 10 HEPES, 2 MgCl2, 10 glucose, pH 7.4; pipette: 126 NaCl, 1.5 CaCl2, 10 HEPES, 10 glucose; pH 7.4.
	<ol>
		<li>Add inhibitors to the pipette solution to block activity of endogenous channels, which are not relevant for the studies (recommended are: 100 &#181;M niflumic acid or DIDS (to block Ca2+-activated Cl- channels), 10 mM TEA (to inhibit the large-conductance Ca2+-dependent K+ channel), 10 nM iberiotoxin (to block the Ca2+-activated K+ channels), 10 &#181;M nicardipine (to block N-type Ca2+ channels)) directly before the patch-clamp experiment.</li>
	</ol>
	</li>
	<li>Gently mix the glomeruli-containing solution, and then apply approximately 50 &#181;l&#160;of it to the poly-&#671;-lysine coated cover glass chips. Let the glomeruli attach for approximately 5 min.</li>
	<li>Move the glass chips with glomeruli to the patch-clamp chamber pre-filled with the bath solution; perfuse the chamber at a rate of 3 ml/min for 1 min to ensure the removal of the unattached glomeruli.</li>
	<li>Conduct a conventional patch-clamp experiment in a cell-attached mode7. With a glass pipette (7 - 10 M&#937; pipette resistance) form a high-resistance seal between a pipette and a podocyte membrane by applying gentle suction (a pipette attached to a podocyte on the surface of the isolated glomerulus is shown in Figure 2, on the left).</li>
	<li>For the cell-attached measurements, low-pass the currents at 300 Hz by an eight-pole Bessel filter.</li>
	<li>Use isolated glomeruli in patch-clamp experiments for up to 4 - 6 hr. Keep the stock glomeruli fraction on ice.</li>
</ol>
4. Ratiometric Confocal Fluorescence Measurements of Intracellular Calcium Concentration in the Podocytes
<ol>
	<li>Place 500 &#181;l of the glomeruli fraction (described at 2.5) in 0.5 ml conical tube and add calcium dyes Fura Red, AM and Fluo-4, AM. Use 2 mM and 1 mM stock concentrations of Fura Red, AM and Fluo-4, AM, respectively (store at -20 &#176;C, dissolved in DMSO) and use 2.5 &#181;l of each dye for a 500 &#181;l of glomeruli fraction. Immediately after addition of the dyes cover the tube with aluminum foil.</li>
	<li>Place tubes on a rotating shaker for at least 20 min up to 1 hr at RT. 
	Note: Pharmacological agents can be added during this step.</li>
	<li>Prepare glass coverslips, cover them with poly-&#671;-lysine and allow drying using heated plate set to 70 &#176;C.</li>
	<li>Once the loading of calcium dyes is complete, apply 100 &#181;l of the glomeruli containing solution to the poly-&#671;-lysine coated coverslips and let them stick to the surface for about 5 min. Mount the glomeruli-attached coverslips into an imaging chamber, and perfuse with the bath solution (containing (in mM): 145 NaCl, 2 CaCl2, 4.5 KCl, 2 MgCl2, 10 HEPES, pH 7.35) at a rate of 3 ml/min to remove the unattached glomeruli and the remaining dyes.</li>
	<li>Set the confocal laser scanning microscope to a 488 nm excitation wavelength and emission filters (525/25 and 650/25 nm for Fluo-4 and Fura Red, respectively). Set the imaging software to a desired frequency and resolution.</li>
	<li>Find the glomeruli in brightfield and then turn on the detection of the fluorescence signal. Adjust the intensity of the laser for each dye to avoid saturation of the signal. Choose the focal plane with the podocytes which are directly attached to the glass. This minimizes the effect caused by contraction of a glomerulus in response to drug application. Double-check that the glomerulus of choice is well attached to the glass;</li>
	<li>Start imaging the focal plane of choice (take 512 x 512 images with the frequency set at 4 sec to visualize fast Ca2+ transient changes. Use a 60X/NA 1.4 or similar objective lens for high resolution image), apply drugs of interest, and record the response.
	<ol>
		<li>Select a desired focal plane (as close to the surface of the glass as possible, to ensure imaging of podocytes on the surface of the glomerulus). Check the intensity of fluorescence on the Fluo4 and FuraRed channels, and make sure that the glomerulus is clearly seen in brightfield.</li>
		<li>Start imaging. Before application of any drugs, record at least 1 min of baseline fluorescence to make sure that the signal is stable (there are no sudden spikes or fading of the signal).</li>
		<li>Apply desired drugs with the help of a micropipette; be careful and ensure the drug was able to diffuse well and reach the glomerulus. Mix the bath solution gently if needed, monitor the selected focal plane and check that it did not move out of focus because of the drug application.</li>
		<li>Record the changes in fluorescence intensity for the Fluo4 and FuraRed signals. Make sure the recording is long enough, by waiting until the signal reaches the plateau level or reaches a peak and then returns to baseline. Perform solution change or addition of any other drugs if needed.</li>
		<li>Stop the recording, and save the file in the native format of the software.</li>
	</ol>
	</li>
</ol>
5. Image Analysis for the Calcium Measurements
<ol>
	<li>Perform image analysis with the ImageJ open software equipped with the ND Utility plugin that allows importing images in the native ND2 format.</li>
	<li>Import the image sequence; make sure to split the channels and use a hyperstack grayscale mode.</li>
	<li>Follow the Analyze &#8594; Tools&#160;&#8594; ROI manager path in the ImageJ software to open a ROI manager window. Select several regions of interest (podocytes) using an oval selection tool and the &#8220;Add (t)&#8221; function in the ROI Manager. As the last ROI, select the area in the background; save the selected ROIs (More&#160;&#8594; Save).</li>
	<li>Highlight the window containing the channel selected for analyses. Use the More&#160;&#8594; Multi Measure function in the ROI Manager, mark the &#8220;one row per slice&#8221; box in the dialogue that pops out, and then click OK. The results will be shown in the format that can be set up in the Results window: enter into the menu option Results&#160;&#8594; Set Measurements, select &#8220;Mean gray value&#8221; and calculate the pixel intensity values for each ROI, which will be displayed in the Results window in separate columns for each ROI.</li>
	<li>Copy the measured ROI intensity values for each channel (Fluo-4 and Fura Red) 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 Fluo-4 to Fura Red channels. Plot scatter/line point-time changes of Ca2+ transient for each ROI. Calculate Mean/SE values for selected glomeruli. 
	Note: Time column should be set according to imaging frequency selected at 4.5.</li>
</ol>
6. Intracellular Calcium Concentration Calculations Using Fluo-4 Fluorescence Signal
<ol>
	<li>Take a sample of the glomeruli and perform the experimental protocol up to step 4.7. After recording of the background fluorescence add Ionomycin (final concentration in the bath chamber should be 10 &#181;M) to the bath solution, and record fluorescence intensity increase. Once the intensity reaches its maximum and the decay starts, add MnCl2 (final concentration should be 5 mM) to quench the fluorescence8 .</li>
	<li>Analyze the data obtained in 6.1. Copy the Fluo-4 signal ROI intensity values obtained according to the protocol described at 5.1 - 5.5 by preferred analysis software.</li>
	<li>Calculate intracellular calcium concentration (in nM) in the podocyte corresponding to the ROI using a formula: 
	<img alt="Equation 1" src="/files/ftp_upload/52850/52850equation1.jpg" /> 
	where Kd is a pre-determined dissociation constant for Fluo-4 (345 nM), F is the intensity at the timepoint that you are calculating the calcium concentration for (baseline), and Fmin and Fmax are the intensity values at the point of maximum calcium load (after ionomycin application) and after quenching of the fluorescence (with MnCl2), respectively (see Figure 3).</li>
</ol>

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The authors have nothing to disclose.

The approach described here allows for the analysis of calcium handling by the podocytes of the rodent glomeruli. This technique allows application of patch-clamp single channel electrophysiology and fluorescence ratiometric confocal imaging. However, both approaches can be used separately, on their own. The proposed protocol has several relatively simple steps, including 1) kidney flush; 2) isolation of the glomeruli by differential sieving; 3) performing patch-clamp electrophysiological experiments, or incubation of th...

Kidneys maintain homeostatic balance for various substances and regulate blood volume in a way that determines total blood pressure. Disturbances in the renal filtration, reabsorption or secretion lead to or accompany pathological states, ranging from hyper- or hypotension to end stage renal disease that eventually requires kidney transplantation. The renal filtering unit (glomerulus) consists of three layers &#8211; the capillary endothelium, basement membrane and a single-cell layer of epithelial cells &#8211; podocytes, which play a major role in the maintenance of the slit-diaphragm integrity and function1. Dysfunction in the permselective glomerular filter causes urinary loss of macromolecules, such as proteinuria. Various agents may affect the structure of the podocytes and their foot processes, which determine the integrity of the glomeruli filtration barrier.
The podocytes are involved in the maintenance of the glomeruli filtration function. It has been established that improper calcium handling by the podocyte leads to cell injury and plays an important role in the progression of various forms of nephropathies2,3. Therefore, development of a model which allows for direct measuring of intracellular calcium concentration changes will be instrumental for studies of podocyte function. Isolated glomeruli were previously used in a numerous studies including measurement of albumin reflection coefficient changes4 and assessment of integral cellular currents in the whole-cell electrophysiological patch-clamp measurements5,6. In the present paper we describe the protocol that allows the researcher to measure intracellular calcium concentration changes in response to applications of pharmacological agents, estimate basal levels of calcium within the cells, and assess individual calcium channels activity. Ratometric calcium concentration measurements and patch-clamp electrophysiology were used to determine changes in the intracellular calcium concentration within the podocyte and channel activity, respectively.

<table><tbody><tr><td>Fluo4 AM</td><td>Life Technologies</td><td>F14217</td><td>500 &amp;micro;l in DMSO</td></tr><tr><td>FuraRed AM</td><td>Life Technologies</td><td>F-3020</td><td></td></tr><tr><td>Poly-&amp;#671;-lysine</td><td>Sigma-Aldrich</td><td>P4707</td><td></td></tr><tr><td>Pluronic acid</td><td>Sigma-Aldrich</td><td>F-68</td><td>solution</td></tr><tr><td>Ionomycin</td><td>Sigma-Aldrich</td><td>I3909-1ML</td><td></td></tr><tr><td>Tube rotator</td><td>Miltenyi Biotec GmbH</td><td>130-090-753</td><td>Germany</td></tr><tr><td>Nikon confocal microscope (inverted)</td><td>Nikon</td><td>Nikon A1R</td><td>Laser exitation 488 nm. Emission filters 500-550 nm and 570-620 nm</td></tr><tr><td>Objective</td><td>Nikon</td><td>Plan Apo 60x/NA 1.4 Oil</td><td></td></tr><tr><td>Cover Glass</td><td>Thermo Scientific</td><td>6661B52</td><td></td></tr><tr><td>High vacuum grease</td><td>Dow Corning</td><td></td><td>Silicone Compound</td></tr><tr><td>Software</td><td>Nikon</td><td>Nikon NIS-Elements</td><td></td></tr><tr><td>Recording/perfusion chamber</td><td>Warner Instruments</td><td>RC-26</td><td></td></tr><tr><td>Patch clamp amplifier</td><td>Molecular Devices</td><td>MultiClamp 700B</td><td></td></tr><tr><td>Data acquisition system</td><td>Molecular Devices</td><td>Digidata 1440A</td><td>Axon Digidata&amp;reg; System</td></tr><tr><td>Low pass filter</td><td>Warner Instruments</td><td>LPF-8</td><td>8 pole Bessel</td></tr><tr><td>Borosilicate glass capillaries</td><td>World Precision Instruments</td><td>1B150F-4</td><td></td></tr><tr><td>Micropipette puller</td><td>Sutter Instrument Co</td><td>P-97</td><td>Flaming/Brown type micropipette puller</td></tr><tr><td>Microforge</td><td>Narishige</td><td>MF-830</td><td>Japan</td></tr><tr><td>Motorized micromanipulator</td><td>Sutter Instrument Co</td><td>MP-225</td><td></td></tr><tr><td>Inverted microscope</td><td>Nikon</td><td>Eclipse Ti</td><td></td></tr><tr><td>Microvibration isolation table</td><td>TMC</td><td></td><td>equipped with Faraday cage</td></tr><tr><td>Multichannel valve perfusion system</td><td>AutoMake Scientific</td><td>Valve Bank II</td><td></td></tr><tr><td>Recording/perfusion chamber</td><td>Warner Instruments</td><td>RC-26</td><td></td></tr><tr><td>Software</td><td>Molecular Devices</td><td>pClamp 10.2</td><td></td></tr><tr><td>Nicardipine</td><td>Sigma-Aldrich</td><td>N7510</td><td></td></tr><tr><td>Iberiotoxin</td><td>Sigma</td><td>I5904-5UG</td><td></td></tr><tr><td>Niflumic acid</td><td>Sigma-Aldrich</td><td>N0630</td><td></td></tr><tr><td>DIDS</td><td>Sigma-Aldrich</td><td>D3514-25MG</td><td></td></tr><tr><td>TEA chloride</td><td>Tocris</td><td>T2265</td><td></td></tr><tr><td>RPMI 1640</td><td>Life Technologies</td><td>11835030</td><td>without antibiotics</td></tr><tr><td>BSA</td><td>Sigma-Aldrich</td><td>A8327</td><td>30% albumin solution</td></tr><tr><td>Temperature controlled surgical table</td><td>MCW core</td><td></td><td>for rodents</td></tr><tr><td>Steel sieves:</td><td></td><td></td><td>#100 (150 &amp;mu;m), 140 (106 &amp;mu;m)</td></tr><tr><td>Gilson, Inc SIEVE 3 SS FH NO200</td><td>Fisher Sci</td><td>50-871-316</td><td></td></tr><tr><td>Gilson, Inc SIEVE 3 SS FH NO270</td><td>Fisher Sci</td><td>50-871-318</td><td></td></tr><tr><td>Gilson, Inc SIEVE 3 SS FH NO400</td><td>Fisher Sci</td><td>50-871-320</td><td></td></tr><tr><td>mesh 200</td><td>Sigma-Aldrich</td><td>s4145</td><td>screen for CD-1</td></tr><tr><td>Binocular microscope</td><td>Nikon</td><td>Eclipse TS100</td><td></td></tr><tr><td>Binocular microscope</td><td>Nikon</td><td>SMZ745</td><td></td></tr><tr><td>Syringe pump-based perfusion system</td><td>Harvard Apparatus</td><td></td><td></td></tr><tr><td>Polyethylene tubing</td><td>Sigma-Aldrich</td><td>PE50</td><td></td></tr><tr><td>Isofluorane anesthesia</td><td><a href="http://www.vetequip.com/" target="_blank"><img src="/img/new_window.png" /> VetEquip, Inc.</a></td><td>911103</td><td></td></tr><tr><td>Other basic reagents</td><td>Sigma-Aldrich</td><td></td><td></td></tr></tbody></table>

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.

Here we addressed the problem of measuring acute changes in the calcium levels in the podocytes. Figure 1 shows a schematic representation of the experimental protocol designed in order to perform high resolution live fluorescence confocal imaging and single ion channel activity recordings in the podocytes of the freshly isolated rodent glomeruli. Briefly, after the rat is anaesthetized, the kidneys should be flushed with PBS to clear them of blood. Then, the kidneys are excised and decapsulated, and glo...

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

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 ...

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Research

JoVE Journal

Biology

10.9K Views.  Medical College of Wisconsin. 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. 

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

Immunohistochemical and Calcium Imaging Methods in Wholemount Rat Retina

In this paper we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of wholemount retinas for immunohistochemistry and, 2) calcium imaging for the study of voltage gated calcium channel (VGCC) mediated calcium signaling in retinal ganglion cells. The calcium imaging method we describe circumvents issues concerning non-specific loading of displaced amacrine cells in the ganglion cell layer.

Immunohistochemistry protocols are used to study the localization of a specific protein in the retina. Calcium imaging techniques are employed to study calcium dynamics in retinal ganglion cells and their axons.

In this paper we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of wholemount retinas for ...

Neuroscience

An Efficient Sieving Method to Isolate Intact Glomeruli from Adult Rat Kidney

Preservation of glomerular structure and function is pivotal in the prevention of glomerulonephritis, a category of kidney disease characterized by proteinuria which can eventually lead to chronic and end-stage renal disease. The glomerulus is a complex apparatus responsible for the filtration of plasma from the body. In disease, structural integrity is lost and allows for the abnormal leakage of plasma contents into the urine. A method to isolate and examine glomeruli in culture is critical for the study of these diseases. In this protocol, an efficient method of retrieving intact glomeruli from adult rat kidneys while conserving structural and morphological characteristics is described. This process is capable of generating high yields of glomeruli per kidney with minimal contamination from other nephron segments. With these glomeruli, injury conditions can be mimicked by incubating them with a variety of chemical toxins, including protamine sulfate, which causes foot process effacement and proteinuria in animal models. Degree of injury can be assessed using transmission electron microscopy, immunofluorescence staining, and western blotting. Nephrin and Wilms Tumor 1 (WT1) levels can also be assessed from these cultures. Due to the ease and flexibility of this protocol, the isolated glomeruli can be utilized as described or in a way that best suits the needs of the researcher to help better study glomerular health and structure in diseased states.

The main focus of this protocol is to efficiently isolate viable primary glomeruli cultures with minimal contaminants for use in a variety of downstream applications. The isolated glomeruli retain structural relationships between component cell types and can be cultured ex vivo for a short time.

Preservation of glomerular structure and function is pivotal in the prevention of glomerulonephritis, a category of kidney disease characterized by ...

Medicine

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 ...

Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial function, cell death, cell metabolism and cell migration to name but a few. Cytosolic calcium is normally maintained at low nanomolar concentrations; rather it is found in high micromolar to millimolar concentrations in the endoplasmic reticulum, mitochondrial matrix and the extracellular compartment. Upon stimulation, a transient increase in cytosolic calcium serves to signal downstream events. Detecting changes in cytosolic calcium is normally performed using a live cell imaging set up with calcium binding dyes that exhibit either an increase in fluorescence intensity or a shift in the emission wavelength upon calcium binding. However, a live cell imaging set up is not freely accessible to all researchers. Alternative detection methods have been optimized for immunological cells with flow cytometry and for non-immunological adherent cells with a fluorescence microplate reader. Here, we describe an optimized, simple method for detecting changes in epithelial cells with flow cytometry using a single wavelength calcium binding dye. Adherent renal proximal tubule epithelial cells, which are normally difficult to load with dyes, were loaded with a fluorescent cell permeable calcium binding dye in the presence of probenecid, brought into suspension and calcium signals were monitored before and after addition of thapsigargin, tunicamycin and ionomycin.

Calcium is involved in numerous physiological and pathophysiological signaling pathways. Live cell imaging requires specialized equipment and can be time consuming. A quick, simple method using a flow cytometer to determine relative changes in cytosolic calcium in adherent epithelial cells brought into suspension was optimized.

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial ...

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, electrical excitability and cell proliferation. The diversity and specificity of Ca2+ signaling derives from mechanisms by which Ca2+ signals are generated to act over different time and spatial scales, ranging from cell-wide oscillations and waves occurring over the periods of minutes to local transient Ca2+ microdomains (Ca2+ puffs) lasting milliseconds. Recent advances in electron multiplied CCD (EMCCD) cameras now allow for imaging of local Ca2+ signals with a 128 x 128 pixel spatial resolution at rates of &#62;500 frames sec-1 (fps). This approach is highly parallel and enables the simultaneous monitoring of hundreds of channels or puff sites in a single experiment. However, the vast amounts of data generated (ca. 1 Gb per min) render visual identification and analysis of local Ca2+ events impracticable. Here we describe and demonstrate the procedures for the acquisition, detection, and analysis of local IP3-mediated Ca2+ signals in intact mammalian cells loaded with Ca2+ indicators using both wide-field epi-fluorescence (WF) and total internal reflection fluorescence (TIRF) microscopy. Furthermore, we describe an algorithm developed within the open-source software environment Python that automates the identification and analysis of these local Ca2+ signals. The algorithm localizes sites of Ca2+ release with sub-pixel resolution; allows user review of data; and outputs time sequences of fluorescence ratio signals together with amplitude and kinetic data in an Excel-compatible table.

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Here we present techniques for imaging local IP3-mediated Ca2+ events using fluorescence microscopy in intact mammalian cells loaded with Ca2+ indicators together with an algorithm that automates identification and analysis of these events.

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, ...

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.

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 ...

In Vivo Calcium Imaging of Neuronal Ensembles in Networks of Primary Sensory Neurons in Intact Dorsal Root Ganglia

Ca2+ imaging can be used as a proxy for cellular activity, including action potentials and various signaling mechanisms involving Ca2+ entry into the cytoplasm or the release of intracellular Ca2+ stores. Pirt-GCaMP3-based Ca2+ imaging of primary sensory neurons of the dorsal root ganglion (DRG) in mice offers the advantage of simultaneous measurement of a large number of cells. Up to 1,800 neurons can be monitored, allowing neuronal networks and somatosensory processes to be studied as an ensemble in their normal physiological context at a populational level in vivo. The large number of neurons monitored allows the detection of activity patterns that would be challenging to detect using other methods. Stimuli can be applied to the mouse hindpaw, allowing the direct effects of stimuli on the DRG neuron ensemble to be studied. The number of neurons producing Ca2+ transients as well as the amplitude of Ca2+ transients indicates sensitivity to specific sensory modalities. The diameter of neurons provides evidence of activated fiber types (non-noxious mechano vs. noxious pain fibers, A&#946;, A&#948;, and C fibers). Neurons expressing specific receptors can be genetically labeled with td-Tomato and specific Cre recombinases together with Pirt-GCaMP. Therefore, Pirt-GCaMP3 Ca2+ imaging of DRG provides a powerful tool and model for the analysis of specific sensory modalities and neuron subtypes acting as an ensemble at the populational level to study pain, itch, touch, and other somatosensory signals.

In Vivo Calcium Imaging of Neuronal Ensembles in Networks of Primary Sensory Neurons in Intact Dorsal Root Ganglia

This protocol describes the surgical exposure of the dorsal root ganglion (DRG) followed by GCaMP3 (genetically-encoded Ca2+ indicator; Green Fluorescent Protein-Calmodulin-M13 Protein 3) Ca2+ imaging of the neuronal ensembles using Pirt-GCaMP3 mice while applying a variety of stimuli to the ipsilateral hind paw.

Ca2+ imaging can be used as a proxy for cellular activity, including action potentials and various signaling mechanisms involving Ca2+ entry into the ...

Imaging Calcium Dynamics in Subpopulations of Mouse Pancreatic Islet Cells

Pancreatic islet hormones regulate blood glucose homeostasis. Changes in blood glucose induce oscillations of cytosolic calcium in pancreatic islet cells that trigger secretion of three main hormones: insulin (from &#946;-cells), glucagon (&#945;-cells) and somatostatin (&#948;-cells). &#946;-Cells, which make up the majority of islet cells and are electrically coupled to each other, respond to the glucose stimulus as one single entity. The excitability of the minor subpopulations, &#945;-cells and &#948;-cells (making up around 20% (30%) and 4% (10%) of the total rodent1 (human2) islet cell numbers, respectively) is less predictable and is therefore of special interest.
Calcium sensors are delivered into the peripheral layer of cells within the isolated islet. The islet or a group of islets is then immobilized and imaged using a fluorescence microscope. The choice of the imaging mode is between higher throughput (wide-field) and better spatial resolution (confocal). Conventionally, laser scanning confocal microscopy is used for imaging tissue, as it provides the best separation of the signal between the neighboring cells. A wide-field system can be utilized too, if the contaminating signal from the dominating population of &#946;-cells is minimized.
Once calcium dynamics in response to specific stimuli have been recorded, data are expressed in numerical form as fluorescence intensity vs. time, normalized to the initial fluorescence and baseline-corrected, to remove the effects linked to bleaching of the fluorophore. Changes in the spike frequency or partial area under the curve (pAUC) are computed vs. time, to quantify the observed effects. pAUC is more sensitive and quite robust whereas spiking frequency provides more information on the mechanism of calcium increase.
Minor cell subpopulations can be identified using functional responses to marker compounds, such as adrenaline and ghrelin, that induce changes in cytosolic calcium in a specific populations of islet cells.

Here, we present a protocol for imaging and quantifying calcium dynamics in heterogeneous cell populations, such as pancreatic islet cells. Fluorescent reporters are delivered into the peripheral layer of cells within the islet, which is then immobilized and imaged, and per-cell analysis of the dynamics of fluorescence intensity is performed.

Pancreatic islet hormones regulate blood glucose homeostasis. Changes in blood glucose induce oscillations of cytosolic calcium in pancreatic islet cells ...

Single Cell Calcium Imaging for Channel Activation Studies

Single-Cell Calcium Imaging for Studying the Activation of Calcium Ion Channels

Single cell Ca2+ imaging is essential for the study of Ca2+ channels activated by various stimulations like temperature, voltage, native compound and chemicals et al. It primarily relies on microscopy imaging technology and the related Ca2+ indicator Fura-2/AM (AM is the abbreviation for Acetoxymethyl ester). Inside the cells, Fura-2/AM is hydrolyzed by esterases into Fura-2, which can reversibly bind with free cytoplasmic Ca2+. The maximum excitation wavelength shifts from 380nm to 340nm (when saturated with Ca2+) upon binding. The emitted fluorescence intensity is quantitatively related to the concentration of bound Ca2+. By measuring the 340/380 ratio, the Ca2+ concentration in the cytoplasm can be determined, eliminating errors caused by variations in the loading efficiency of the fluorescent probe among different samples. This technology allows for real-time, quantitative, and simultaneous monitoring of Ca2+ changes in multiple cells. The results are stored in &#8220;.XLSX&#8221; format for subsequent analysis, which is fast and generates intuitive change curves, greatly improving the detection efficiency. From different experimental perspectives, this article lists the use of this technology to detect Ca2+ signals in cells with endogenous or overexpressed channel proteins. Meantime, different methods for activating cells were also showed and compared. The aim is to provide readers with a clearer understanding of the usage and applications of single cell Ca2+ imaging.

This article presents a method for real-time, quantitative monitoring of calcium ion (Ca2+) concentrations in cells using single-cell Ca2+ imaging with the Fura-2/AM dye. This technique enables efficient dye loading and accurate calculation of Ca2+ levels through fluorescence intensity ratios, making it a simple and rapid approach for research applications.

Single cell Ca2+ imaging is essential for the study of Ca2+ channels activated by various stimulations like temperature, voltage, native compound and ...

Isolating and Imaging Live, Intact Pacemaker Regions of Mouse Renal Pelvis by Vibratome Sectioning

The renal pelvis (RP) is a funnel-shaped, smooth muscle structure that facilitates normal urine transport from the kidney to the ureter by regular, propulsive contractions. Regular RP contractions rely on pacemaker activity, which originates from the most proximal region of the RP at the pelvis-kidney junction (PKJ). Due to the difficulty in accessing and preserving intact preparations of the PKJ, most investigations on RP pacemaking have focused on single-cell electrophysiology and Ca2+ imaging experiments. Although important revelations on RP pacemaking have emerged from such work, these experiments have several intrinsic limitations, including the inability to accurately determine cellular identity in mixed suspensions and the lack of in situ imaging of RP pacemaker activity. These factors have resulted in a limited understanding of the mechanisms that underlie normal rhythmic RP contractions. In this paper, a protocol is described to prepare intact segments of mouse PKJ using a vibratome sectioning technique. By combining this approach with mice expressing cell-specific reporters and genetically encoded Ca2+ indicators, investigators may be able to more accurately study the specific cell types and mechanisms responsible for peristaltic RP contractions that are vital for normal urine transport.

The goal of this protocol is to isolate intact pacemaker regions of the mouse renal pelvis using vibratome sectioning. These sections can then be used for in situ Ca2+ imaging to elucidate Ca2+ transient properties of pacemaker cells and other interstitial cells in vibratome slices.

The renal pelvis (RP) is a funnel-shaped, smooth muscle structure that facilitates normal urine transport from the kidney to the ureter by regular, ...

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

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