This work was supported by NIH grants R01DK119183 (to G.A. and M.D.C.) and S10OD028596 (to G.A.) and by the Cell Physiology and Model Organisms Kidney Imaging Cores of the Pittsburgh Center for Kidney Research (P30DK079307).

Care and handling of the animals were carried out in accordance with the University of Pittsburgh Institutional Animal Care and Use Committee. Female, 2-4-month-old C57Bl/6J mice were used for the present study. The mice were obtained commercially (see Table of Materials).
1. Equipment assembly and setup
<ol>
	<li>Perform Ca2+ imaging with an upright microscope equipped with a high-resolution camera and a stable light source (see Table of Materials).
	<ol>
		<li>Acquire the images with a microscope compatible software that permits direct control of the camera, light source, and piezo actuator via a USB digital I/O device (see Table of Materials). 
		NOTE: Figure 1 represents a schematic of the setup. GCaMP5G has a peak excitation wavelength of 470 nm and a peak emission wavelength of 497 nm28. Use a filter cube suitable for GCaMP5G imaging.</li>
	</ol>
	</li>
	<li>For the stimulation of individual urothelial cells in the bladder mucosa preparation (see step 2), use a piezoelectric actuator controlled by a single channel open-loop piezo controller (see Table of Materials). Mount the piezoelectric actuator in a micromanipulator.
	<ol>
		<li>Mount the glass micropipettes in a pipette holder affixed to the piezoelectric actuator (Figure 1). Remotely operate the piezo controller by an analog/digital converter controlled by an electrophysiology data acquisition and analysis program (see Table of Materials). 
		NOTE: An external trigger, in the form of a transistor-transistor logic (TTL) signal initiated by the imaging software, is used to trigger the stimulation protocol in the electrophysiology data acquisition and analysis software that moves the piezoelectric actuator (Figure 1).</li>
	</ol>
	</li>
	<li>Ensure that the imaging software (see Table of Materials) interfaces with the digital I/O device. Configure a digital output channel in the USB digital I/O device to deliver the TTL signal to the analog/digital converter, which will initiate the protocol in the electrophysiology software that moves the piezoelectric actuator.
	<ol>
		<li>Use a cable to connect the Start BNC port in the analog/digital converter to the ground (GND) and a screw terminal in the USB digital I/O device.</li>
	</ol>
	</li>
	<li>Set up the recording protocol in the imaging software. 
	NOTE: The following steps describe how to generate a protocol that will send a TTL signal and start collecting images at specified time intervals.
	<ol>
		<li>Set up the acquisition protocol in the imaging software by clicking on the Experiment Manager and selecting New Experiment. A new window will open.</li>
		<li>Select the icon Time Lapse Loop and drag it to the newly opened window; set the number of cycles to 2 and the interval to the fastest setting allowed in the setup.</li>
		<li>From the icon Transmitted Shutter/Manual Shutter, select the icon NI USB-6501 and drag it into the Time Lapse Loop window, and then set the NI USB-6501 as closed. Drag an additional NI USB-6501 from the Transmitted Shutter/Manual Shutter into the Time Lapse Loop window and set it as open. To connect the two NI USB-6501 icons, drag the arrowhead on the side of the NI USB-6501 closed icon and pull until it touches the NI USB-6501 open icon. A line that connects both icons will appear.</li>
		<li>Drag another Time Lapse Loop into the Experiment Manager window and connect it to the first one by pulling the arrowhead. Set the parameters for recording. Set the number of cycles of the new Time Lapse Loop to 2400.</li>
		<li>From the Image Acquisition icon, drag the GFP filter into the recently opened Time Lapse Loop and set the exposure time to 100 ms.</li>
		<li>Set the camera image type to 8-bit, resolution to 576 x 576 (Binning 4 x 4), pixel clock to 480 MHz, and Hot pixel correction to standard. 
		NOTE: The parameters can be adjusted depending on the level of expression of the fluorescent sensor, camera sensitivity, and setup configuration.</li>
	</ol>
	</li>
	<li>Set up the Lab Bench in the electrophysiology software (see Table of Materials). 
	NOTE: This will define the output signal in mV of the analog/digital converter.
	<ol>
		<li>Go to the Configure menu in the electrophysiology software and select Lab Bench. In the resulting Output Signals window, select Analog Out #1, press the Add signal, and name it (e.g., &#34;Piezo&#34;). Set the Signal Unit to mV and the Scale Factor (mV/V) to 1.</li>
	</ol>
	</li>
	<li>Generate a stimulation protocol in the electrophysiology software following the steps below.
	<ol>
		<li>To generate a new stimulation protocol in the electrophysiology software, go to the menu item Acquire and select New Protocol.</li>
		<li>Set Mode/Rate to Episodic Stimulation, Run/Trial to 1, Sweep/Run to 1, and Sweep Duration (s) to 150.</li>
		<li>In the Outputs menu, select channel #1 Piezo as Analog out.</li>
		<li>In the Trigger menu, set the trigger to Digitizer Start Input and Internal Timer.</li>
		<li>In the Waveform menu, select channel #1 and Analog Waveform with Epochs. Set Step A to Step, First Level to 0, Delta level to 0, First duration to 10000, and Delta duration to 0.</li>
		<li>Set Step B to Step, First level to 10, Delta Level to 0, First Duration to 1000, and Delta Duration to 0. Set Step C to Step, First Level to 0, Delta Level to 0, First Duration to 130000, and Delta Duration to 0.</li>
		<li>Save the protocol and name it Stimulation protocol.</li>
	</ol>
	</li>
	<li>Use a BNC cable to connect Analog Output #1 in the analog/digital converter to the EXT INPUT in the front of the single-channel open-loop piezo controller (see Table of Materials).</li>
	<li>Fabricate glass micropipettes for the mechanical stimulation of umbrella cells from capillary glass tubing following the steps below.
	<ol>
		<li>Place the capillary glass in a puller (see Table of Materials) and adjust it with the capillary retaining knobs.</li>
		<li>Position the heater unit at its full height. Adjust the first pull terminating position slider of the puller to 5.</li>
		<li>Set the first heater knob to 76.7 and the second knob to 52.7.</li>
		<li>Pull the glass capillary in two steps by pressing the start button.</li>
		<li>Close the tip of the micropipette with a microforge (see Table of Materials) with the heater adjustment knob set at 60. 
		NOTE: For the present protocol, the final diameter of the micropipette tip used for poking individual cells is ~1-3 &#181;m.</li>
	</ol>
	</li>
	<li>Verify that the stimulating micropipette moves the distance specify in the stimulation protocol. 
	NOTE: The following steps are to ensure that 12.5 s after initiating the stimulation protocol in the electrophysiology software, a voltage pulse with a duration of 1 s is generated, making the piezoelectric actuator move 20 &#956;m.&#160;
	<ol>
		<li>Mount a micropipette in the holder and attach it to the piezoelectric actuator.</li>
		<li>Place the piezoelectric actuator and the attached micropipette parallel to the center of the microscope stage and within the area of view.</li>
		<li>Focus on the pipette&#39;s tip and immobilize the piezoelectric mounting rod (see Table of Materials) with tape. Under bright field illumination, adjust the camera parameters to achieve a clear image of the pipette tip.</li>
		<li>Open the Stimulation protocol in the electrophysiology software and set it to play. 
		NOTE: The electrophysiology protocol will not start until the TTL signal sent from the imaging software is received.</li>
		<li>From Experiment Manager in the imaging software, press Start to initiate data acquisition. 
		NOTE: This will trigger the protocol in the electrophysiology software, which will drive the piezoelectric actuator. The protocol in the imaging software will generate a file with the images of the experiment.</li>
	</ol>
	</li>
	<li>Verify the distance traveled by the pipette following the steps below.
	<ol>
		<li>To measure the distance traveled by the pipette during the stimulation protocol, select the Count and Measure item in the imaging software.</li>
		<li>From the Measure menu, select the Measurement and ROI option, and a new window will appear below the movie window.</li>
		<li>From the Measure menu, choose the Arbitrary Line.</li>
		<li>In the movie window, draw an arbitrary line starting at the pipette&#39;s tip. Note that the end of the arbitrary line will be adjusted later.</li>
		<li>Right click on the arbitrary line to convert the line into a region of interest (ROI), which will be visible in all movie frames.</li>
		<li>Inspect the movie and adjust the end of the arbitrary line (ROI) to the final position traveled by the pipette in response to the stimulus. 
		&#8203;NOTE: The distance traveled by the pipette in &#181;m will appear in the Measurement and ROI window. The stimulation protocol can be modified according to user needs.</li>
	</ol>
	</li>
</ol>
2. In situ transduction and isolation of the bladder mucosa
<ol>
	<li>Transduce female mouse bladders in situ with an adenovirus encoding the cDNA of CGaMP5G according to the procedure described in the virus transduction protocol31.
	<ol>
		<li>When transducing urothelial cells for Ca2+ imaging experiments, instill the bladders with 50 &#181;L of the solution containing 2 x 107 infectious viral particles (IVP). 
		NOTE: This technique tends to restrict&#160;the expression of CGaMP5G to the umbrella cell layer. Alternatively, experiments could be conducted with urinary bladders harvested from transgenic mice expressing CGaMP5G in urothelial cells (i.e., GCaMP5G expression driven by a uroplakin-2 promotor) or any other cell type of interest.</li>
	</ol>
	</li>
	<li>24-72 h after transduction, euthanize the mice by CO2 asphyxiation and perform a thoracotomy by opening the chest cavity with scissors to cause the lungs to collapse.</li>
	<li>Canulate the urethra with a 24 G catheter. Expose the bladder by an abdominal incision of ~1.5 cm through the skin and muscle, and use a 6.0 suture (see Table of Materials) to secure the catheter to the urethra.</li>
	<li>Harvest the bladder and urethra32 and affix to a silicone rubber holder pad (see Table of Materials) bathed in recording solution containing (in mM): 135 NaCl, 5.0 KCl, 1 MgCl2, 2.5 CaCl2, 10 glucose, 10 HEPES, pH 7.4, and bubbled up with 100% O2 (Figure 2A).</li>
	<li>Separate the bladder mucosa from the underlying muscular layer with fine forceps following previously published reports32 (Figure 2B).</li>
	<li>Cut open the bladder mucosa and pin it down with the urothelium facing up to a silicone elastomer insert in the bottom of a 35 mm diameter tissue cultured dish (Figure 2C).</li>
</ol>
3. Mechanical stimulation of individual urothelial cells and Ca2+ imaging
<ol>
	<li>Mount the tissue cultured dish with the pinned bladder mucosa in the microscope stage equipped with a culture dish incubator with resistive heating elements (Figure 2E). Perfuse (following the manufacturer&#39;s instructions, see Table of Materials) the cell culture dish continuously at a rate of 1.7 mL/min with recording solution warmed at ~37 &#176;C with an in-line heater.</li>
	<li>Maintain the temperature of the tissue dish incubator and solutions at ~37 &#176;C with a dual channel bipolar temperature controller model (see Table of Materials). Equilibrate the tissue in the chamber with continuous perfusion for at least 15 min before conducting further experimental procedures.</li>
	<li>To record mechanically induced Ca2+ transients, submerge the micropipette in the solution bathing the pinned bladder mucosa in the tissue dish. Move the micropipette&#160;to the center of the field with the help of a low magnification scanning objective (4x) using bright field illumination.
	<ol>
		<li>Move the micropipette near the surface of the urothelial tissue by coordinately moving the micromanipulator in the vertical plane and adjusting the focus.</li>
		<li>Switch the objective to one with a higher magnification (20x) suitable for immunofluorescence with a high numerical aperture (NA).</li>
		<li>Set the micromanipulator to Fine and move the micropipette&#160;near the top of the target cell.</li>
		<li>Change the field of view to camera and press Live in the imaging software. 
		NOTE: This must turn on the reflected shutter and allow the observation of the fluorescent signal emitting from the tissue in the computer.</li>
		<li>Adjust the focus to visualize the top of the cell and adjust the pipette&#39;s position if necessary.</li>
		<li>Open the Stimulation protocol in the electrophysiology software and set it to play. 
		NOTE: The protocol in the electrophysiology software will not start until the TTL signal sent from the imaging software is received.</li>
		<li>From Experiment Manager in the imaging software, press Start to initiate data acquisition. This will trigger the protocol in the electrophysiology software, which will drive the piezoelectric actuator and generate a file with the images of the experiment. The stimulation protocol can be modified according to the user&#39;s needs.</li>
	</ol>
	</li>
</ol>
4. Data analysis
<ol>
	<li>Quantify the fluorescence intensity over time following the steps below.
	<ol>
		<li>Open the image file in the imaging software and select the Count and Measure window. Select the polygon tool and draw a ROI on the boundaries of the cell that was poked.</li>
		<li>Go to the Measure window, select Intensity Profile, set the measurement to Over time, Results to Average, Background Subtraction to none, and then press Execute. The mean fluorescence intensity will be computed over time (Figure 3).</li>
		<li>To export the intensity profile data, click on the Excel icon in the Intensity Profile Results window. This will let the user choose the destination folder and file name and save the data in a .xlsx format.</li>
	</ol>
	</li>
	<li>Perform data analysis using scientific graphing and data analysis software (see Table of Materials). 
	NOTE: The amplitude of the Ca2+ peak evoked by poking is expressed as the change in fluorescence intensity (&#916;F/F), where F is the fluorescence intensity of GCaMP5G at time 0, and &#916;F is the difference between the fluorescence intensity maxima and the basal at time 0. The decay of the Ca2+ response can also be calculated (not shown).</li>
</ol>

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G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+roles+for+PIEZO1+and+PIEZO2+in+urothelial+mechanotransduction+and+lower+urinary+tract+interoception.">Functional roles for PIEZO1 and PIEZO2 in urothelial mechanotransduction and lower urinary tract interoception.</a> JCI Insight. 6 (19), 152984 (2021).</li><li>Durnin, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+ex+vivo+bladder+model+with+detrusor+smooth+muscle+removed+to+analyse+biologically+active+mediators+released+from+the+suburothelium.">An ex vivo bladder model with detrusor smooth muscle removed to analyse biologically active mediators released from the suburothelium.</a> The Journal of Physiology. 597 (6), 1467-1485 (2019).</li><li>Delmas, P., Coste, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechano-gated+ion+channels+in+sensory+systems.">Mechano-gated ion channels in sensory systems.</a> Cell. 155 (2), 278-284 (2013).</li><li>Tavernarakis, N., Driscoll, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Degenerins.+At+the+core+of+the+metazoan+mechanotransducer.">Degenerins. At the core of the metazoan mechanotransducer.</a> Annals of the New York Academy of Sciences. 940 (1), 28-41 (2001).</li><li>Peyronnet, R., Tran, D., Girault, T., Frachisse, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanosensitive+channels:+feeling+tension+in+a+world+under+pressure.">Mechanosensitive channels: feeling tension in a world under pressure.</a> Frontiers in Plant Science. 5, 558 (2014).</li><li>Blount, P., Iscla, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Life+with+bacterial+mechanosensitive+channels,+from+discovery+to+physiology+to+pharmacological+target.">Life with bacterial mechanosensitive channels, from discovery to physiology to pharmacological target.</a> Microbiology and Molecular Biology Reviews. 84 (1), 00055 (2020).</li><li>Booth, I. R., Miller, S., M&#252;ller, A., Lehtovirta-Morley, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+evolution+of+bacterial+mechanosensitive+channels.">The evolution of bacterial mechanosensitive channels.</a> Cell Calcium. 57 (3), 140-150 (2015).</li></ol>

The authors have nothing to disclose.

All organisms, and seemingly most cell types, express ion channels that respond to mechanical stimuli20,33,34,35,36,37. The function of these mechanically activated channels has been predominantly assessed with the patch-clamp technique. However, due to accessibility issues, patch-clamp studies of mechanically activated ion c...

Epithelial cells in the urinary tract are subjected to mechanical forces as the urinary filtrate travels through the nephrons, and urine is pumped out of the renal pelvis and travels through the ureters to be stored in the urinary bladder. It has been long recognized that mechanical forces (e.g., shear stress and stretch) exerted by fluids on the epithelial cells that line the urinary tract regulate the reabsorption of protein in the proximal tubule and of solutes in the distal nephron1,2,3,4,5,6,7,8,9,10,11,12,13, as well as the storage of urine in the urinary bladder and micturition14,15,16,17.
The conversion of mechanical stimuli into electrical and biochemical signals, a process referred to as mechanotransduction, is mediated by proteins that respond to the deformation of cellular structures or the associated extracellular matrix18,19,20,21. Mechanically activated ion channels are unique in the sense that they transition from a closed state to an open permeable state in response to changes in membrane tension, pressure, or shear stress18,19,20,21,22. In addition, Ca2+ transients can be initiated by&#160;integrin-mediated mechanotransduction or by activation of mechanoresponsive adhesion systems at cell-cell junctions23,24,25,26. Ion channel function is usually assessed with the patch-clamp technique, which involves the formation of a gigaohm seal between the cell membrane and the patch pipette27. However, cells located in deep tissue layers with a dense extracellular matrix (e.g., kidney tubules) or surrounded by a physical barrier (e.g., glycocalyx) are difficult to access with a glass micropipette. Likewise, cells embedded or that are integral parts of tissues with poor mechanical stability (e.g., the urothelium) can not be readily studied with the patch-clamp technique. Because many mechanically activated ion channels are permeable to Ca2+, an alternative approach is to assess their activity by fluorescent microscopy using a Ca2+-sensitive dye or genetically encoded calcium indicators (GECIs) such as GCaMP. Recent efforts in protein engineering have significantly increased the dynamic range, sensitivity, and response of GECIs28,29,30, and advances in genetics have allowed their expression in specific cell populations, making them ideally suited to study mechanotransduction.
The urothelium, the stratified epithelium that covers the interior of the urinary bladder, functions as a barrier, preventing the diffusion of urinary solutes into the bladder interstitium, but also functions as a transducer, sensing bladder fullness and communicating these events to the underlying nerves and musculature16. Previous studies have shown that the communication between the urothelium and underlying tissues requires the mechanically activated ion channels Piezo1 and Piezo231. To assess mechanically induced Ca2+ transients in urothelial cells, a new technique described that uses adenoviral gene transfer to express the Ca2+ sensor GCaMP5G in urothelial cells was developed. This technique employs a mucosal sheet preparation that provides easy access to the outermost umbrella cell layer and a computer-assisted system for the simultaneous mechanical stimulation of individual cells with a closed glass micropipette and recording of changes in fluorescence over time.

<table><tbody><tr><td>20x Objective</td><td>Olympus</td><td>UMPlanFL N</td><td></td></tr><tr><td>24 G &amp;frac34;&amp;rdquo; catheter</td><td>Medline&amp;nbsp;</td><td>Suresite IV slide&amp;nbsp;</td><td></td></tr><tr><td>4x Objective</td><td>Olympus</td><td>UPlanFL N</td><td></td></tr><tr><td>Analog/digital converter</td><td>Molecular Devices</td><td>Digidata 1440A</td><td></td></tr><tr><td>Anti-GFP antibody</td><td>Abcam&amp;nbsp;</td><td>Ab6556</td><td></td></tr><tr><td>Beam splitter</td><td>Chroma</td><td>T495lpxr</td><td></td></tr><tr><td>Bipolar temperature controller&amp;nbsp;</td><td>Warner Instruments</td><td>TC-344B</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Fluka</td><td>21114-1L</td><td>1 M solution</td></tr><tr><td>cellSens software</td><td>Olympus</td><td></td><td>Imaging software</td></tr><tr><td>CMOS camera</td><td>Hamamatsu</td><td>ORCA fusion</td><td></td></tr><tr><td>Donkey anti-rabbit conjugated to Alexa Fluor 488&amp;nbsp;</td><td>Jackson ImmunoResearch</td><td>711-545-152</td><td></td></tr><tr><td>Excel</td><td>Microsoft Corporation</td><td></td><td></td></tr><tr><td>Filter&amp;nbsp;</td><td>Chroma&amp;nbsp;</td><td>ET470/40X</td><td></td></tr><tr><td>Glass capillaries Corning 8250 glass</td><td>Warner Instruments&amp;nbsp;</td><td>G85150T-4</td><td></td></tr><tr><td>Glucose</td><td>Sigma</td><td>G8270</td><td></td></tr><tr><td>HEPES&amp;nbsp;</td><td>Sigma</td><td>H4034</td><td></td></tr><tr><td>Inline heater&amp;nbsp;</td><td>Warner Instruments</td><td>SH-27B</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>793590</td><td></td></tr><tr><td>Light source</td><td>Sutter Instruments</td><td>Lambda XL&amp;nbsp;</td><td></td></tr><tr><td>Manifold pump tubing</td><td>Fisherbrand</td><td>14-190-510</td><td>ID 1.52 mm</td></tr><tr><td>Manifold pump tubing</td><td>Fisherbrand</td><td>14-190-533</td><td>ID 2.79 mm</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>M9272</td><td></td></tr><tr><td>Mice&amp;nbsp;</td><td>Jackson Lab</td><td>664</td><td>2-4 months old female C57BL/6J</td></tr><tr><td>Microforge</td><td>Narishige&amp;nbsp;</td><td>MF-830</td><td></td></tr><tr><td>Micromanipulator</td><td>Sutter Instruments</td><td>MP-285</td><td></td></tr><tr><td>Microscope</td><td>Olympus</td><td>BX51W</td><td></td></tr><tr><td>Mounting media with DAPI</td><td>Invitrogen</td><td>S36964&amp;nbsp;</td><td>Slowfade Diamond Antifade with DAPI</td></tr><tr><td>NaCl&amp;nbsp;</td><td>Sigma</td><td>S7653</td><td></td></tr><tr><td>pClamp software</td><td>Molecular Devices</td><td>Version 10.4</td><td>Patch-clamp electrophysiology data acquisition and analysis software</td></tr><tr><td>Peristaltic pump</td><td>Gilson</td><td>Minipuls 3</td><td></td></tr><tr><td>Piezoelectric actuator</td><td>Thorlabs</td><td>PAS005</td><td></td></tr><tr><td>Pipette holder</td><td>World Precision Instruments</td><td></td><td></td></tr><tr><td>Pipette puller</td><td>Narishige</td><td>PP-830</td><td></td></tr><tr><td>Quick exchange heated base with perfusion and adapter ring kit</td><td>Warner Instruments</td><td>QE-1</td><td>Quick exchange platform fits 35 mm dish&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Rhodamine-phalloidin&amp;nbsp;</td><td>Invitrogen</td><td>R415</td><td></td></tr><tr><td>Sigma-Plot</td><td>Systat Software Inc</td><td>Version 14.0</td><td>Scientific graphing and data analysis software&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Silicone elastomer</td><td>Dow</td><td>Sylgard 184</td><td></td></tr><tr><td>Single channel open-loop piezo controller</td><td>Thorlabs</td><td>MDT694B</td><td></td></tr><tr><td>Square grid holder pad</td><td>Ted Pella</td><td>10520</td><td></td></tr><tr><td>Suture</td><td>AD Surgical</td><td>S-S618R13</td><td>6-0 Sylk</td></tr><tr><td>Teflon mounting rod</td><td>Custom made</td><td></td><td>Use to mount the piezoelectric actuator in the micromanipulator</td></tr><tr><td>Tubing</td><td>Fisher Scientific</td><td>14171129</td><td>Tygon S3 ID 1/16 IN, OD 1/8 IN</td></tr><tr><td>USB Digital I/O device&amp;nbsp;</td><td>National Instruments</td><td>NI USB-6501</td><td></td></tr></tbody></table>

ex vivo analysis of mechanically activated ca2+ transients in urothelial cells

Mechanically activated ion channels are biological transducers that convert mechanical stimuli such as stretch or shear forces into electrical and biochemical signals. In mammals, mechanically activated channels are essential for the detection of external and internal stimuli in processes as diverse as touch sensation, hearing, red blood cell volume regulation, basal blood pressure regulation, and the sensation of urinary bladder fullness. While the function of mechanically activated ion channels has been extensively studied in the in vitro setting using the patch-clamp technique, assessing their function in their native environment remains a difficult task, often because of limited access to the sites of expression of these channels (e.g., afferent terminals, Merkel cells, baroreceptors, and kidney tubules) or difficulties applying the patch-clamp technique (e.g., the apical surfaces of urothelial umbrella cells). This protocol describes a procedure to assess mechanically evoked Ca2+ transients using the fluorescent sensor GCaMP5G in an ex vivo urothelial preparation, a technique that could be readily adapted for the study of mechanically evoked Ca2+ events in other native tissue preparations.

The present protocol describes a technique to assess mechanically evoked Ca2+ transients in umbrella cells using the fluorescent Ca2+ sensor GCaMP5G. Adenoviral transduction was employed to express GCaMP5G in urothelial cells due to its high efficiency and because it produces an elevated level of expression. Fluorescent images of stained cryosections from a transduced bladder are shown in Figure 2D. For these experiments, GCaMP5G expression is highest in the umbrella ce...

Watch this Scientific Journal Video about Ex Vivo Analysis of Mechanically Activated Ca2+ Transients in Urothelial Cells at JoVE.com

Ex Vivo Analysis of Mechanically Activated Ca2+ Transients in Urothelial Cells

This protocol describes a methodology to assess the function of mechanically activated ion channels in native urothelial cells using the fluorescent Ca2+ sensor GCaMP5G.

Ex Vivo Analysis of Mechanically Activated Ca2+ Transients in Urothelial Cells

Mechanically activated ion channels are biological transducers that convert mechanical stimuli such as stretch or shear forces into electrical and ...

ex-vivo-analysis-mechanically-activated-ca2-transients-urothelial

Research

JoVE Journal

Medicine

1.0K Views.  University of Pittsburgh. Dit protocol beschrijft een techniek om mechanisch opgewekte calciumgebeurtenis te bestuderen in een ex vivo urotheelpreparaat. De techniek is relatief eenvoudig uit te voeren en kan worden aangepast om mechanisch opgewekte calciumgebeurtenissen in andere inheemse weefselpreparaten te bestuderen. Fabriceer de micropipettes door het capillaire glas in de trekker te plaatsen en het aan te passen met de capillairhoudende knoppen.Trek het glazen capillair in twee stappen volgens de instruc...

Video: Ex Vivo Analysis of Mechanically Activated Ca2+ Transients in Urothelial Cells

Focal Ca2+ Transient Detection in Smooth Muscle

Ca2+ imaging of smooth muscle provides insight into cellular mechanisms that may not result in changes of membrane potential, such as the release of Ca2+ from internal stores, and allows multiple cells to be monitored simultaneously to assess, for example, coupling in syncytial tissue. Subcellular Ca2+ transients are common in smooth muscle, yet are difficult to measure accurately because of the problems caused by their stochastic occurrence, over an often wide field of view, in an organ that it prone to contract. To overcome this problem, we've developed a series of imaging protocols and analysis routines to acquire and then analyse, in an automated fashion, the frequency, location and amplitude of such events. While this approach may be applied in other contexts, our own work involves the detection of local purinergic Ca2+ transients for locating transmitter release with submicron resolution. 
ATP is released as a cotransmitter from autonomic nerves, where it binds to P2X1 receptors on the smooth muscle of the detrusor and vas deferens. Ca2+ enters the smooth muscle, resulting in purinergic neuroeffector Ca2+ transients (NCTs). The focal Ca2+ transients allow the optical monitoring of neurotransmitter release in a manner that has many advantages over electrophysiology. Apart from the greatly improved spatial resolution, optical recording has the additional advantage of allowing the recording of transmitter release from many distinguishable sites simultaneously. Furthermore, the optical plane of focus is easier to maintain or correct during long recording series than is the repositioning of an intracellular sharp microelectrode. 
In summary, a method for imaging of Ca2+ fluorescence is outlined which details the preparation of tissue, and the acquisition and analysis of data. We outline the use of several scripts for the analysis of such Ca2+ transients.

Focal Ca2+ Transient Detection in Smooth Muscle

Details methods for high-resolution Ca2+ imaging of smooth muscle within isolated organs, including: preparation of the tissue, image acquisition and data analysis.

Ca2+ imaging of smooth muscle provides insight into cellular mechanisms that may not result in changes of membrane potential, such as the release of Ca2+ ...

Biology

Mechanical Stimulation-induced Calcium Wave Propagation in Cell Monolayers: The Example of Bovine Corneal Endothelial Cells

Intercellular communication is essential for the coordination of physiological processes between cells in a variety of organs and tissues, including the brain, liver, retina, cochlea and vasculature. In experimental settings, intercellular Ca2+-waves can be elicited by applying a mechanical stimulus to a single cell. This leads to the release of the intracellular signaling molecules IP3 and Ca2+ that initiate the propagation of the Ca2+-wave concentrically from the mechanically stimulated cell to the neighboring cells. The main molecular pathways that control intercellular Ca2+-wave propagation are provided by gap junction channels through the direct transfer of IP3 and by hemichannels through the release of ATP. Identification and characterization of the properties and regulation of different connexin and pannexin isoforms as gap junction channels and hemichannels are allowed by the quantification of the spread of the intercellular Ca2+-wave, siRNA, and the use of inhibitors of gap junction channels and hemichannels. Here, we describe a method to measure intercellular Ca2+-wave in monolayers of primary corneal endothelial cells loaded with Fluo4-AM in response to a controlled and localized mechanical stimulus provoked by an acute, short-lasting deformation of the cell as a result of touching the cell membrane with a micromanipulator-controlled glass micropipette with a tip diameter of less than 1 &mu;m. We also describe the isolation of primary bovine corneal endothelial cells and its use as model system to assess Cx43-hemichannel activity as the driven force for intercellular Ca2+-waves through the release of ATP. Finally, we discuss the use, advantages, limitations and alternatives of this method in the context of gap junction channel and hemichannel research.

Intercellular Ca2+-waves are driven by gap junction channels and hemichannels. Here, we describe a method to measure intercellular Ca2+-waves in cell monolayers in response to a local single-cell mechanical stimulus and its application to investigate the properties and regulation of gap junction channels and hemichannels.

Intercellular communication is essential for the coordination of physiological processes between cells in a variety of organs and tissues, including the ...

A Decentralized (Ex Vivo) Murine Bladder Model with the Detrusor Muscle Removed for Direct Access to the Suburothelium during Bladder Filling

Previous studies have established the release of chemical substances from flat bladder mucosa sheets affixed in Ussing chambers and exposed to changes in hydrostatic pressure or mechanical stretch and from cultured urothelial cells upon hydrostatic pressure changes, stretch, cell swelling, or drag forces, and in bladder lumen at end of filling. Such findings led to the assumption that these mediators are also released in suburothelium (SubU)/lamina propria (LP) during bladder filling, where they affect cells deep in the bladder wall to ultimately regulate bladder excitability. There are at least two obvious limitations in such studies: 1) none of these approaches provide direct information about the presence of mediators in SubU/LP, and 2) the stimuli used are not physiological and do not recapitulate authentic filling of the bladder. Here, we discuss a procedure that enables direct access to the suburothelial surface of the bladder mucosa in the course of bladder filling. The murine detrusor-free preparation we created closely resembles filling of the intact bladder and allows pressure-volume studies to be performed on the bladder in the absence of confounding signaling from spinal reflexes and detrusor smooth muscle. Using the novel detrusor-free bladder model, we recently demonstrated that intravesical measurements of mediators cannot be used as a proxy to what has been released or present in the SubU/LP during bladder filling. The model enables examination of urothelium-derived signaling molecules that are released, generated by metabolism and/or transported into the SubU/LP during the course of bladder filling to transmit information to neurons and smooth muscle of the bladder and regulate its excitability during continence and micturition.

A Decentralized Ex Vivo Murine Bladder Model with the Detrusor Muscle Removed for Direct Access to the Suburothelium during Bladder Filling

The detrusor-free bladder model enables direct access to the suburothelium to study local mechanisms for regulation of biologically active mediator availability in suburothelium/lamina propria during storage and voiding of urine. The preparation closely resembles filling of an intact bladder and allows pressure-volume studies to be performed without systemic influences.

Previous studies have established the release of chemical substances from flat bladder mucosa sheets affixed in Ussing chambers and exposed to changes in ...

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 ...

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 ...

Analysis of Cardiac Contractile Dysfunction and Ca2+ Transients in Rodent Myocytes

Contractile dysfunction and Ca2+ transients are often analyzed at the cellular level as part of a comprehensive assessment of cardiac-induced injury and/or remodeling. One approach for assessing these functional alterations utilizes unloaded shortening and Ca2+ transient analyses in primary adult cardiac myocytes. For this approach, adult myocytes are isolated by collagenase digestion, made Ca2+ tolerant, and then adhered to laminin-coated coverslips, followed by electrical pacing in serum-free media. The general protocol utilizes adult rat cardiac myocytes but can be readily adjusted for primary myocytes from other species. Functional alterations in myocytes from injured hearts can be compared to sham myocytes and/or to in vitro therapeutic treatments. The methodology includes the essential elements needed for myocyte pacing, along with the cell chamber and platform components. The detailed protocol for this approach incorporates the steps for measuring unloaded shortening by sarcomere length detection and cellular Ca2+ transients measured with the ratiometric indicator Fura-2 AM, as well as for raw data analysis.

Analysis of Cardiac Contractile Dysfunction and Ca2+ Transients in Rodent Myocytes

A set of protocols are presented that describe the measurement of contractile function via sarcomere length detection along with calcium (Ca2+) transient measurement in isolated rat myocytes. The application of this approach for studies in animal models of heart failure is also included.

Contractile dysfunction and Ca2+ transients are often analyzed at the cellular level as part of a comprehensive assessment of cardiac-induced injury ...

In Vivo Luminal Measurement of Distension-Evoked Urothelial ATP Release in Rodents

ATP, released from the urothelium in response to bladder distension, is thought to play a significant sensory role in the control of micturition. Therefore, accurate measurement of urothelial ATP release in a physiological setting is an important first step in studying the mechanisms that control purinergic signaling in the urinary bladder. Existing techniques to study mechanically evoked urothelial ATP release utilize cultured cells plated on flexible supports or bladder tissue pinned into Ussing chambers; however, each of these techniques does not fully emulate conditions in the intact bladder. Therefore, an experimental setup was developed to directly measure ATP concentrations in the lumen of the rodent urinary bladder.
In this setup, the bladders of anesthetized rodents are perfused through catheters in both the dome of the bladder and via the external urethral orifice. Pressure in the bladder is increased by capping the urethral catheter while perfusing sterile fluid into the bladder through the dome. Measurement of intravesical pressure is achieved using a pressure transducer attached to the bladder dome catheter, akin to the setup used for cystometry. Once the desired pressure is reached, the urethral catheter's cap is removed, and fluid collected for ATP quantification by luciferin-luciferase assay. Through this experimental setup, the mechanisms controlling both mechanical and chemical stimulation of urothelial ATP release can be interrogated by including various agonists or antagonists into the perfusate or by comparing results between wildtype and genetically modified animals.

In Vivo Luminal Measurement of Distension-Evoked Urothelial ATP Release in Rodents

This protocol describes the procedure for measuring ATP concentrations in the lumen of the bladder in an anesthetized rodent.

ATP, released from the urothelium in response to bladder distension, is thought to play a significant sensory role in the control of micturition. ...

Applications of Spatio-temporal Mapping and Particle Analysis Techniques to Quantify Intracellular Ca2+ Signaling In Situ

Ca2+ imaging of isolated cells or specific types of cells within intact tissues often reveals complex patterns of Ca2+ signaling. This activity requires careful and in-depth analyses and quantification to capture as much information about the underlying events as possible. Spatial, temporal and intensity parameters intrinsic to Ca2+ signals such as frequency, duration, propagation, velocity and amplitude may provide some biological information required for intracellular signalling. High-resolution Ca2+ imaging typically results in the acquisition of large data files that are time consuming to process in terms of translating the imaging information into quantifiable data, and this process can be susceptible to human error and bias. Analysis of Ca2+ signals from cells in situ typically relies on simple intensity measurements from arbitrarily selected regions of interest (ROI) within a field of view (FOV). This approach ignores much of the important signaling information contained in the FOV. Thus, in order to maximize recovery of information from such high-resolution recordings obtained with Ca2+dyes or optogenetic Ca2+ imaging, appropriate spatial and temporal analysis of the Ca2+ signals is required. The protocols outlined in this paper will describe how a high volume of data can be obtained from Ca2+ imaging recordings to facilitate more complete analysis and quantification of Ca2+ signals recorded from cells using a combination of spatiotemporal map (STM)-based analysis and particle-based analysis. The protocols also describe how different patterns of Ca2+ signaling observed in different cell populations in situ can be analyzed appropriately. For illustration, the method will examine Ca2+ signaling in a specialized population of cells in the small intestine, interstitial cells of Cajal (ICC), using GECIs.

Applications of Spatio-temporal Mapping and Particle Analysis Techniques to Quantify Intracellular Ca2+ Signaling In Situ

Genetically encoded Ca2+ indicators (GECIs) have radically changed how in situ Ca2+ imaging is performed. To maximize data recovery from such recordings, appropriate analysis of Ca2+ signals is required. The protocols in this paper facilitate the quantification of Ca2+ signals recorded in situ using spatiotemporal mapping and particle-based analysis.

Ca2+ imaging of isolated cells or specific types of cells within intact tissues often reveals complex patterns of Ca2+ signaling. This activity requires ...

Preparation and Utilization of Freshly Isolated Human Detrusor Smooth Muscle Cells for Characterization of 9-Phenanthrol-Sensitive Cation Currents

Detrusor smooth muscle (DSM) cells present within the urinary bladder wall ultimately facilitate urine storage and voiding. Preparation of the viable, fresh, and isolated DSM cells presents an important technical challenge whose achievement provides optimal cells for subsequent functional and molecular studies. The method developed and elaborated herein, successfully used by our group for over a decade, describes dissection of human urinary bladder specimens obtained from open bladder surgeries followed by an enzymatic two-step treatment of DSM pieces and mechanical trituration to obtain freshly isolated DSM cells. The initial step involves dissection to separate the DSM layer (also known as muscularis propria) from mucosa (urothelium, lamina propria, and muscularis mucosa) and the adjacent connective, vascular, and adipose tissues present. The DSM is then cut into pieces (2-3 mm x 4-6 mm) in nominal Ca2+-containing dissection/digestion solution (DS). DSM pieces are next transferred to and sequentially treated separately with DS containing papain and collagenase at ~37 &#176;C for 30-45 min per step. Following washes with DS containing enzyme-free bovine serum and trituration with a fire-polished pipette, the pieces release single DSM cells. Freshly isolated DSM cells are ideally suited for patch-clamp electrophysiological and pharmacological characterizations of ion channels. Specifically, we show that the TRPM4 channel blocker 9-phenanthrol reduces voltage-step evoked cation currents recorded with the amphotericin-B perforated patch-clamp approach. DSM cells can also be studied by other techniques such as single cell RT-PCR, microarray analysis, immunocytochemistry, in situ proximity ligation assay, and Ca2+ imaging. The main advantage of utilizing single DSM cells is that the observations made relate directly to single cell characteristics revealed. Studies of freshly isolated human DSM cells have provided important insights characterizing the properties of various ion channels including cation-permeable in the urinary bladder and will continue as a gold standard in elucidating DSM cellular properties and regulatory mechanisms.

We describe a method for preparation of the single freshly isolated detrusor smooth muscle cells from human urinary bladder specimens employing a two-step enzymatic procedure. The obtained viable DSM cells can be studied by various single cell techniques including the described amphotericin-B patch-clamp electrophysiology to reveal physiological and pharmacological properties.

Detrusor smooth muscle (DSM) cells present within the urinary bladder wall ultimately facilitate urine storage and voiding. Preparation of the viable, ...

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 ...

Ex Vivo Analysis of Mechanically Activated Ca²⁺ Transients in Urothelial Cells

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