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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
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			<td>20x Objective</td>
			<td>Olympus</td>
			<td>UMPlanFL N</td>
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			<td>24 G &#190;&#8221; catheter</td>
			<td>Medline&#160;</td>
			<td>Suresite IV slide&#160;</td>
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			<td>4x Objective</td>
			<td>Olympus</td>
			<td>UPlanFL N</td>
			<td></td>
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			<td>Analog/digital converter</td>
			<td>Molecular Devices</td>
			<td>Digidata 1440A</td>
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			<td>Anti-GFP antibody</td>
			<td>Abcam&#160;</td>
			<td>Ab6556</td>
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			<td>Beam splitter</td>
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			<td>T495lpxr</td>
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			<td>Bipolar temperature controller&#160;</td>
			<td>Warner Instruments</td>
			<td>TC-344B</td>
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			<td>CaCl2</td>
			<td>Fluka</td>
			<td>21114-1L</td>
			<td>1 M solution</td>
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			<td>cellSens software</td>
			<td>Olympus</td>
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			<td>Imaging software</td>
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			<td>CMOS camera</td>
			<td>Hamamatsu</td>
			<td>ORCA fusion</td>
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			<td>Donkey anti-rabbit conjugated to Alexa Fluor 488&#160;</td>
			<td>Jackson ImmunoResearch</td>
			<td>711-545-152</td>
			<td></td>
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			<td>Excel</td>
			<td>Microsoft Corporation</td>
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			<td>Filter&#160;</td>
			<td>Chroma&#160;</td>
			<td>ET470/40X</td>
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			<td>Glass capillaries Corning 8250 glass</td>
			<td>Warner Instruments&#160;</td>
			<td>G85150T-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma</td>
			<td>G8270</td>
			<td></td>
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			<td>HEPES&#160;</td>
			<td>Sigma</td>
			<td>H4034</td>
			<td></td>
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			<td>Inline heater&#160;</td>
			<td>Warner Instruments</td>
			<td>SH-27B</td>
			<td></td>
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			<td>KCl</td>
			<td>Sigma</td>
			<td>793590</td>
			<td></td>
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		<tr>
			<td>Light source</td>
			<td>Sutter Instruments</td>
			<td>Lambda XL&#160;</td>
			<td></td>
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			<td>Manifold pump tubing</td>
			<td>Fisherbrand</td>
			<td>14-190-510</td>
			<td>ID 1.52 mm</td>
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			<td>Manifold pump tubing</td>
			<td>Fisherbrand</td>
			<td>14-190-533</td>
			<td>ID 2.79 mm</td>
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			<td>MgCl2</td>
			<td>Sigma</td>
			<td>M9272</td>
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			<td>Mice&#160;</td>
			<td>Jackson Lab</td>
			<td>664</td>
			<td>2-4 months old female C57BL/6J</td>
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			<td>Microforge</td>
			<td>Narishige&#160;</td>
			<td>MF-830</td>
			<td></td>
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			<td>Micromanipulator</td>
			<td>Sutter Instruments</td>
			<td>MP-285</td>
			<td></td>
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		<tr>
			<td>Microscope</td>
			<td>Olympus</td>
			<td>BX51W</td>
			<td></td>
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			<td>Mounting media with DAPI</td>
			<td>Invitrogen</td>
			<td>S36964&#160;</td>
			<td>Slowfade Diamond Antifade with DAPI</td>
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		<tr>
			<td>NaCl&#160;</td>
			<td>Sigma</td>
			<td>S7653</td>
			<td></td>
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		<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>
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		<tr>
			<td>Pipette holder</td>
			<td>World Precision Instruments</td>
			<td></td>
			<td></td>
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			<td>Pipette puller</td>
			<td>Narishige</td>
			<td>PP-830</td>
			<td></td>
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			<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&#160;&#160;</td>
		</tr>
		<tr>
			<td>Rhodamine-phalloidin&#160;</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&#160;&#160;</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&#160;</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 ...

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Universidad de Cartagena UDEC

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JoVE Journal

Medicine

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

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

Ex Vivo Infection of Live Tissue with Oncolytic Viruses

Oncolytic Viruses (OVs) are novel therapeutics that selectively replicate in and kill tumor cells1. Several clinical trials evaluating the effectiveness of a variety of oncolytic platforms including HSV, Reovirus, and Vaccinia OVs as treatment for cancer are currently underway2-5. One key characteristic of oncolytic viruses is that they can be genetically modified to express reporter transgenes which makes it possible to visualize the infection of tissues by microscopy or bio-luminescence imaging6,7. This offers a unique advantage since it is possible to infect tissues from patients ex vivo prior to therapy in order to ascertain the likelihood of successful oncolytic virotherapy8. To this end, it is critical to appropriately sample tissue to compensate for tissue heterogeneity and assess tissue viability, particularly prior to infection9. It is also important to follow viral replication using reporter transgenes if expressed by the oncolytic platform as well as by direct titration of tissues following homogenization in order to discriminate between abortive and productive infection. The object of this protocol is to address these issues and herein describes 1. The sampling and preparation of tumor tissue for cell culture 2. The assessment of tissue viability using the metabolic dye alamar blue 3. Ex vivo infection of cultured tissues with vaccinia virus expressing either GFP or firefly luciferase 4. Detection of transgene expression by fluorescence microscopy or using an In Vivo Imaging System (IVIS) 5. Quantification of virus by plaque assay. This comprehensive method presents several advantages including ease of tissue processing, compensation for tissue heterogeneity, control of tissue viability, and discrimination between abortive infection and bone fide viral replication.

Ex Vivo Infection of Live Tissue with Oncolytic Viruses

Oncolytic viruses are promising for cancer therapeutics. The ability to ascertain the infectability of live tissue specimens obtained from patients prior to treatment is a unique advantage of this therapeutic approach. This protocol describes how to process tissues for ex vivo infection with oncolytic virus and subsequent viral quantification.

Oncolytic Viruses (OVs) are novel therapeutics that selectively replicate in and kill tumor cells1. Several clinical trials evaluating the effectiveness ...

In vivo 19F MRI for Cell Tracking

In vivo 19F MRI allows quantitative cell tracking without the use of ionizing radiation. It is a noninvasive technique that can be applied to humans. Here, we describe a general protocol for cell labeling, imaging, and image processing. The technique is applicable to various cell types and animal models, although here we focus on a typical mouse model for tracking murine immune cells. The most important issues for cell labeling are described, as these are relevant to all models. Similarly, key imaging parameters are listed, although the details will vary depending on the MRI system and the individual setup. Finally, we include an image processing protocol for quantification. Variations for this, and other parts of the protocol, are assessed in the Discussion section. Based on the detailed procedure described here, the user will need to adapt the protocol for each specific cell type, cell label, animal model, and imaging setup. Note that the protocol can also be adapted for human use, as long as clinical restrictions are met.

In vivo 19F MRI for Cell Tracking

We describe a general protocol for in vivo cell tracking using MRI in a mouse model with ex vivo labeled cells. A typical protocol for cell labeling, image acquisition processing and quantification is included.

In vivo 19F MRI allows quantitative cell tracking without the use of ionizing radiation. It is a noninvasive technique that can be applied to humans. ...

Molecular Profiling of the Invasive Tumor Microenvironment in a 3-Dimensional Model of Colorectal Cancer Cells and Ex vivo Fibroblasts

Invading colorectal cancer (CRC) cells have acquired the capacity to break free from their sister cells, infiltrate the stroma, and remodel the extracellular matrix (ECM). Characterizing the biology of this phenotypically distinct group of cells could substantially improve our understanding of early events during the metastatic cascade.

Tumor invasion is a dynamic process facilitated by bidirectional interactions between malignant epithelium and the cancer associated stroma. In order to examine cell-specific responses at the tumor stroma-interface we have combined organotypic co-culture and laser micro-dissection techniques.
Organotypic models, in which key stromal constituents such as fibroblasts are 3-dimentioanally co-cultured with cancer epithelial cells, are highly manipulatable experimental tools which enable invasion and cancer-stroma interactions to be studied in near-physiological conditions.

Laser microdissection (LMD) is a technique which entails the surgical dissection and extraction of the various strata within tumor tissue, with micron level precision.
By combining these techniques with genomic, transcriptomic and epigenetic profiling we aim to develop a deeper understanding of the molecular characteristics of invading tumor cells and surrounding stromal tissue, and in doing so potentially reveal novel biomarkers and opportunities for drug development in CRC.&#160;&#160;&#160;

Molecular Profiling of the Invasive Tumor Microenvironment in a 3-Dimensional Model of Colorectal Cancer Cells and Ex vivo Fibroblasts

Molecular profiling of laser microdissected cells and stroma from synthetic, tissue-engineered colorectal cancer models represents a novel and manipulatable approach to characterize and dissect the distinctive biology at the interface between tumor cells at the invasive front and cancer associated stromal cells. &#160;

Invading colorectal cancer (CRC) cells have acquired the capacity to break free from their sister cells, infiltrate the stroma, and remodel the ...

An Ex vivo Model to Study Hormone Action in the Human Breast

The study of hormone action in the human breast has been hampered by lack of adequate model systems. Upon in vitro culture, primary mammary epithelial cells tend to lose hormone receptor expression. Widely used hormone receptor positive breast cancer cell lines are of limited relevance to the in vivo situation. Here, we describe an ex vivo model to study hormone action in the human breast. Fresh human breast tissue specimens from surgical discard material such as reduction mammoplasties or mammectomies are mechanically and enzymatically digested to obtain tissue fragments containing ducts and lobules and multiple stromal cell types. These tissue microstructures kept in basal medium without growth factors preserve their intercellular contacts, the tissue architecture, and remain hormone responsive for several days. They are readily processed for RNA and protein extraction, histological analysis or stored in freezing medium. Fluorescence activated cell sorting (FACS) can be used to enrich for specific cell populations. This protocol provides a straightforward, standard approach for translational studies with highly complex, varied human specimens.

An Ex vivo Model to Study Hormone Action in the Human Breast

We have developed a novel ex vivo model to study hormone action in the human breast. It is based on tissue microstructures isolated from surgical breast tissue specimens which preserve tissue architecture, intercellular interactions, and paracrine signaling.

The study of hormone action in the human breast has been hampered by lack of adequate model systems. Upon in vitro culture, primary mammary epithelial ...

Studying Pancreatic Cancer Stem Cell Characteristics for Developing New Treatment Strategies

Pancreatic ductal adenocarcinoma (PDAC) contains a subset of exclusively tumorigenic cancer stem cells (CSCs) which have been shown to drive tumor initiation, metastasis and resistance to radio- and chemotherapy. Here we describe a specific methodology for culturing primary human pancreatic CSCs as tumor spheres in anchorage-independent conditions. Cells are grown in serum-free, non-adherent conditions in order to enrich for CSCs while their more differentiated progenies do not survive and proliferate during the initial phase following seeding of single cells. This assay can be used to estimate the percentage of CSCs present in a population of tumor cells. Both size (which can range from 35 to 250 micrometers) and number of tumor spheres formed represents CSC activity harbored in either bulk populations of cultured cancer cells or freshly harvested and digested tumors 1,2. Using this assay, we recently found that metformin selectively ablates pancreatic CSCs; a finding that was subsequently further corroborated by demonstrating diminished expression of pluripotency-associated genes/surface markers and reduced in vivo tumorigenicity of metformin-treated cells. As the final step for preclinical development we treated mice bearing established tumors with metformin and found significantly prolonged survival. Clinical studies testing the use of metformin in patients with PDAC are currently underway (e.g., NCT01210911, NCT01167738, and NCT01488552). Mechanistically, we found that metformin induces a fatal energy crisis in CSCs by enhancing reactive oxygen species (ROS) production and reducing mitochondrial transmembrane potential. In contrast, non-CSCs were not eliminated by metformin treatment, but rather underwent reversible cell cycle arrest. Therefore, our study serves as a successful example for the potential of in vitro sphere formation as a screening tool to identify compounds that potentially target CSCs, but this technique will require further in vitro and in vivo validation to eliminate false discoveries.

Pancreatic cancer stem cells (CSCs) can be expanded in vitro using the anchorage-independent sphere culture technique, which represents a powerful tool to study CSC biology and can serve as the first step to develop novel CSC-targeting therapies. Here the methodology for expanding, analyzing and targeting of pancreatic CSCs is provided.

Pancreatic ductal adenocarcinoma (PDAC) contains a subset of exclusively tumorigenic cancer stem cells (CSCs) which have been shown to drive tumor ...

Depletion of Mouse Cells from Human Tumor Xenografts Significantly Improves Downstream Analysis of Target Cells

The use of in vitro cell line models for cancer research has been a useful tool. However, it has been shown that these models fail to reliably mimic patient tumors in different assays1. Human tumor xenografts represent the gold standard with respect to tumor biology, drug discovery, and metastasis research 2-4. Tumor xenografts can be derived from different types of material like tumor cell lines, tumor tissue from primary patient tumors4 or serially transplanted tumors. When propagated in vivo, xenografted tissue is infiltrated and vascularized by cells of mouse origin. Multiple factors such as the tumor entity, the origin of xenografted material, growth rate and region of transplantation influence the composition and the amount of mouse cells present in tumor xenografts. However, even when these factors are kept constant, the degree of mouse cell contamination is highly variable.
Contaminating mouse cells significantly impair downstream analyses of human tumor xenografts. As mouse fibroblasts show high plating efficacies and proliferation rates, they tend to overgrow cultures of human tumor cells, especially slowly proliferating subpopulations. Mouse cell derived DNA, mRNA, and protein components can bias downstream gene expression analysis, next-generation sequencing, as well as proteome analysis 5. To overcome these limitations, we have developed a fast and easy method to isolate untouched human tumor cells from xenografted tumor tissue. This procedure is based on the comprehensive depletion of cells of mouse origin by combining automated tissue dissociation with the benchtop tissue dissociator and magnetic cell sorting. Here, we demonstrate that human target cells can be can be obtained with purities higher than 96% within less than 20 min independent of the tumor type.

Human tumor xenografts are vascularized and infiltrated by cells of mouse origin during the growth phase in vivo. To circumvent the bias caused by these contaminating cells during downstream analysis, we developed a method allowing for comprehensive depletion of all mouse cells by magnetic cell sorting.

The use of in vitro cell line models for cancer research has been a useful tool. However, it has been shown that these models fail to reliably mimic ...

Optimized Analysis of In Vivo and In Vitro Hepatic Steatosis

Establishing a system of procedures to qualitatively and quantitatively characterize in vivo and in vitro hepatic steatosis is important for metabolic study in the liver. Here, numerous assays are described to comprehensively measure the phenotype and parameters of hepatic steatosis in mouse and hepatocyte models. 
Combining the physiological, histological, and biochemical methods, this system can be used to assess the progress of hepatic steatosis. In vivo, the measurements of body weight and nuclear magnetic resonance (NMR) provide a general understanding of mice in a non-invasive manner. Hematoxylin and Eosin (H&amp;E) and Oil Red O staining determine the histological morphology and lipid deposition of liver tissue under nutrient overload conditions, such as high-fat diet feeding. Next, the total lipid contents are isolated by chloroform/methanol extraction, which are followed by a biochemical analysis for triglyceride and cholesterol. Moreover, mouse primary hepatocytes are treated with high glucose plus insulin to stimulate lipid accumulation, an efficient in vitro model to mimic diet-induced hyperglycemia and hyperinsulinemia in vivo. Then, the lipid deposition is measured by Oil Red O staining and chloroform/methanol extraction. Oil Red O staining determines intuitive hepatic steatotic phenotypes, while the lipid extraction analysis determines the parameters that can be analyzed statistically. The present protocols are of interest to scientists in the fields of fatty liver diseases, insulin resistance, and type 2 diabetes.

Optimized Analysis of In Vivo and In Vitro Hepatic Steatosis

Here, optimized methods to generate in vivo and in vitro models of hepatic steatosis and to analyze the steatotic phenotypes and related physiological parameters are described.

Establishing a system of procedures to qualitatively and quantitatively characterize in vivo and in vitro hepatic steatosis is important for metabolic ...

Preparation Steps for Measurement of Reactivity in Mouse Retinal Arterioles Ex Vivo

Vascular insufficiency and alterations in normal retinal perfusion are among the major factors for the pathogenesis of various sight-threatening ocular diseases, such as diabetic retinopathy, hypertensive retinopathy, and possibly glaucoma. Therefore, retinal microvascular preparations are pivotal tools for physiological and pharmacological studies to delineate the underlying pathophysiological mechanisms and to design therapies for the diseases. Despite the wide use of mouse models in ophthalmic research, studies on retinal vascular reactivity are scarce in this species. A major reason for this discrepancy is the challenging isolation procedures owing to the small size of these retinal blood vessels, which is ~ &#8804; 30 &#181;m in luminal diameter. To circumvent the problem of direct isolation of these retinal microvessels for functional studies, we established an isolation and preparation technique that enables ex vivo studies of mouse retinal vasoactivity under near-physiological conditions. Although the present experimental preparations will specifically refer to the mouse retinal arterioles, this methodology can readily be employed to microvessels from rats.

Preparation Steps for Measurement of Reactivity in Mouse Retinal Arterioles Ex Vivo

Many vision-threatening ocular diseases are associated with dysfunctional retinal microvessels. Therefore, the measurement of retinal arteriole responses is important to investigate the underlying pathophysiological mechanisms. This article describes a detailed protocol for mouse retinal arteriole isolation and preparation to assess the effects of vasoactive substances on vascular diameter.

Vascular insufficiency and alterations in normal retinal perfusion are among the major factors for the pathogenesis of various sight-threatening ocular ...

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, normothermic ex vivo liver perfusion (NEVLP) has been introduced as a method to evaluate and modify organ function. NEVLP has many advantages in comparison to hypothermic and subnormothermic perfusion including reduced preservation injury, restoration of normal organ function under physiologic conditions, assessment of organ performance, and as a platform for organ repair, remodeling, and modification. Both murine and porcine NEVLP models have been described. We demonstrate a rat model of NEVLP and use this model to show one of its important applications &#8212; the use of a therapeutic molecule added to liver perfusate. Catalase is an endogenous reactive oxygen species (ROS) scavenger and has been demonstrated to decrease ischemia-reperfusion in the eye, brain, and lung. Pegylation has been shown to target catalase to the endothelium. Here, we added pegylated-catalase (PEG-CAT) to the base perfusate and demonstrated its ability to mitigate liver preservation injury. An advantage of our rodent NEVLP model is that it is inexpensive in comparison to larger animal models. A limitation of this study is that it does not currently include post-perfusion liver transplantation. Therefore, prediction of the function of the organ post-transplantation cannot be made with certainty. However, the rat liver transplant model is well established and certainly could be used in conjunction with this model. In conclusion, we have demonstrated an inexpensive, simple, easily replicable NEVLP model using rats. Applications of this model can include testing novel perfusates and perfusate additives, testing software designed for organ evaluation, and experiments designed to repair organs.

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant liver donor shortage, and criteria for liver donors have been expanded. Normothermic ex vivo liver perfusion (NEVLP) has been developed to evaluate and modify organ function. This study demonstrates a rat model of NEVLP and tests the ability of pegylated-catalase, to mitigate liver preservation injury.

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, ...

Ex Vivo Expansion of Hematopoietic Stem Cells from Human Umbilical Cord Blood-derived CD34+ Cells Using Valproic Acid

Umbilical cord blood (UCB) units provide an alternative source of human hematopoietic stem cells (HSCs) for patients who require allogeneic bone marrow transplantation. While UCB has several unique advantages, the limited numbers of HSCs within each UCB unit limits their use in regenerative medicine and HSC transplantation in adults. Efficient expansion of functional human HSCs can be achieved by ex vivo culturing of CD34+ cells isolated from UCBs and treated with a deacetylase inhibitor, valproic acid (VPA). The protocol detailed here describes the culture conditions and methodology to rapidly isolate CD34+ cells and expand to a high degree a pool of primitive HSCs. The expanded HSCs are capable of establishing both short-term and long-term engraftment and are able to give rise to all types of differentiated hematopoietic cells. This method also holds potential for clinical application in autologous HSC gene therapy and provides an attractive approach to overcome the loss of functional HSCs associated with gene editing.

Ex Vivo Expansion of Hematopoietic Stem Cells from Human Umbilical Cord Blood-derived CD34+ Cells Using Valproic Acid

Here, we describe the ex vivo expansion of hematopoietic stem cells from CD34+ cells derived from umbilical cord blood treated with a combination of a cytokine cocktail and VPA. This method leads to a significant degree of ex vivo expansion of primitive HSCs for either clinical or laboratory applications.

Umbilical cord blood (UCB) units provide an alternative source of human hematopoietic stem cells (HSCs) for patients who require allogeneic bone marrow ...

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

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Optimized Analysis of In Vivo and In Vitro Hepatic Steatosis

Preparation Steps for Measurement of Reactivity in Mouse Retinal Arterioles Ex Vivo

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

Ex Vivo Expansion of Hematopoietic Stem Cells from Human Umbilical Cord Blood-derived CD34⁺ Cells Using Valproic Acid