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

Marcelo D. Carattino; Wily G. Ruiz; Gerard Apodaca

doi:10.3791/64532

Medicine

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

Published: September 28, 2022 doi: 10.3791/64532

Marcelo D. Carattino^1,2, Wily G. Ruiz¹, Gerard Apodaca^1,2

¹Renal-Electrolyte Division, Department of Medicine, University of Pittsburgh, ²Department of Cell Biology, University of Pittsburgh

Summary

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

Abstract

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.

Introduction

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 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.

Subscription Required. Please recommend JoVE to your librarian.

Protocol

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

Perform Ca²⁺ imaging with an upright microscope equipped with a high-resolution camera and a stable light source (see Table of Materials).
1. 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.
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.
1. 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).
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.
1. 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.
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.
1. Set up the acquisition protocol in the imaging software by clicking on the Experiment Manager and selecting New Experiment. A new window will open.
2. 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.
3. 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.
4. 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.
5. From the Image Acquisition icon, drag the GFP filter into the recently opened Time Lapse Loop and set the exposure time to 100 ms.
6. 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.
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.
1. 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., "Piezo"). Set the Signal Unit to mV and the Scale Factor (mV/V) to 1.
Generate a stimulation protocol in the electrophysiology software following the steps below.
1. To generate a new stimulation protocol in the electrophysiology software, go to the menu item Acquire and select New Protocol.
2. Set Mode/Rate to Episodic Stimulation, Run/Trial to 1, Sweep/Run to 1, and Sweep Duration (s) to 150.
3. In the Outputs menu, select channel #1 Piezo as Analog out.
4. In the Trigger menu, set the trigger to Digitizer Start Input and Internal Timer.
5. 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.
6. 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.
7. Save the protocol and name it Stimulation protocol.
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).
Fabricate glass micropipettes for the mechanical stimulation of umbrella cells from capillary glass tubing following the steps below.
1. Place the capillary glass in a puller (see Table of Materials) and adjust it with the capillary retaining knobs.
2. Position the heater unit at its full height. Adjust the first pull terminating position slider of the puller to 5.
3. Set the first heater knob to 76.7 and the second knob to 52.7.
4. Pull the glass capillary in two steps by pressing the start button.
5. 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 µm.
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 μm.
1. Mount a micropipette in the holder and attach it to the piezoelectric actuator.
2. Place the piezoelectric actuator and the attached micropipette parallel to the center of the microscope stage and within the area of view.
3. Focus on the pipette'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.
4. 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.
5. 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.
Verify the distance traveled by the pipette following the steps below.
1. To measure the distance traveled by the pipette during the stimulation protocol, select the Count and Measure item in the imaging software.
2. From the Measure menu, select the Measurement and ROI option, and a new window will appear below the movie window.
3. From the Measure menu, choose the Arbitrary Line.
4. In the movie window, draw an arbitrary line starting at the pipette's tip. Note that the end of the arbitrary line will be adjusted later.
5. Right click on the arbitrary line to convert the line into a region of interest (ROI), which will be visible in all movie frames.
6. 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.
  NOTE: The distance traveled by the pipette in µm will appear in the Measurement and ROI window. The stimulation protocol can be modified according to user needs.

2. In situ transduction and isolation of the bladder mucosa

Transduce female mouse bladders in situ with an adenovirus encoding the cDNA of CGaMP5G according to the procedure described in the virus transduction protocol³¹.
1. When transducing urothelial cells for Ca²⁺ imaging experiments, instill the bladders with 50 µL of the solution containing 2 x 10⁷ infectious viral particles (IVP).
  NOTE: This technique tends to restrict 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.
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.
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.
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).
Separate the bladder mucosa from the underlying muscular layer with fine forceps following previously published reports³² (Figure 2B).
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).

3. Mechanical stimulation of individual urothelial cells and Ca²⁺ imaging

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'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 °C with an in-line heater.
Maintain the temperature of the tissue dish incubator and solutions at ~37 °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.
To record mechanically induced Ca²⁺ transients, submerge the micropipette in the solution bathing the pinned bladder mucosa in the tissue dish. Move the micropipette to the center of the field with the help of a low magnification scanning objective (4x) using bright field illumination.
1. Move the micropipette near the surface of the urothelial tissue by coordinately moving the micromanipulator in the vertical plane and adjusting the focus.
2. Switch the objective to one with a higher magnification (20x) suitable for immunofluorescence with a high numerical aperture (NA).
3. Set the micromanipulator to Fine and move the micropipette near the top of the target cell.
4. 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.
5. Adjust the focus to visualize the top of the cell and adjust the pipette's position if necessary.
6. 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.
7. 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's needs.

4. Data analysis

Quantify the fluorescence intensity over time following the steps below.
1. 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.
2. 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).
3. 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.
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 (ΔF/F), where F is the fluorescence intensity of GCaMP5G at time 0, and ΔF 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).

Subscription Required. Please recommend JoVE to your librarian.

Representative Results

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 cell layer. A sequence of representative images captured during an experiment where an umbrella cell was poked are shown in Figure 3A. The first image in Figure 3A shows a fluorescent view of the stimulating pipette positioned on the top of an umbrella cell of a bladder mucosa expressing GCaMP5G. Mechanical stimulation of the umbrella cell expressing GCaMP5G causes a deformation (see image Figure 3A at 12.5 s), followed by a rapid increase in the fluorescence emission (see image Figure 3A at 13.5 s). The change in fluorescence was plotted as a function of time (Figure 3B). As reported previously³¹, poking evokes a Ca²⁺ response in most umbrella cells tested in bladders from control and wild-type mice transduced with GCaMP5G. Deleting Piezo1 and Piezo2 from umbrella cells reduced the peak amplitude of the Ca²⁺ response³¹. When comparing the effect of different treatments or genetic backgrounds on mechanically evoked Ca²⁺ responses, it is essential to standardize the experimental conditions, including transduction of the bladder, mounting of the tissue, size of the tip of the pipette used for indentation, mechanical stimulus duration and amplitude, and imaging conditions.

Figure 1: Experimental setup to record mechano-activated Ca²⁺ transients in urothelial cells. The rig consists of an upright microscope, fluorescent light source, excitation and emission filters, and a CMOS camera. The imaging software controls the system. A piezoelectric actuator controlled by a single channel open-loop piezo controller is used to move the poking micropipette. Glass micropipettes are mounted in a pipette holder affixed to the piezoelectric actuator. The piezoelectric actuator and associated micropipette are mounted on a micromanipulator (not shown). The piezo open-loop controller is remotely operated by an analog/digital converter and electrophysiology software. A recording protocol in the imaging software delivers a TTL signal via a digital I/O device, and this initiates the stimulation protocol in the electrophysiology software that controls the piezoelectric and, subsequently, image capture. Please click here to view a larger version of this figure.

Figure 2: Isolation and mounting of the bladder mucosa for Ca²⁺ imaging. (A) Mucosal sheet preparation with an attached catheter. The bladder mucosa was stripped from the underlying muscular layer with fine forceps. (B) Magnified image of the stripped bladder mucosa shown in panel (A). (C) The mucosa was pinned down with the urothelium facing up with 0.15 mm insect pins to a silicone elastomer insert. (D) Confocal immunofluorescence cross-section images of the mucosal sheet preparation stained with an antibody against GFP and a secondary donkey anti-rabbit conjugated antibody (green), rhodamine-phalloidin (red), and DAPI (blue). The bladder mucosal preparation includes part of the lamina propria (LP). Arrows indicate the location of the umbrella cell layer (Ub). Scale bars = 50 µm. (E) Photograph of the experimental setup. a, in-line heater; b, piezoelectric actuator; c, heater elements and controller; d, pipette holder and micropipette; e, mucosal sheet attached to the silicone insert in a tissue cultured dish and dish incubator. Please click here to view a larger version of this figure.

Figure 3. Recording and analysis of mechano-activated Ca²⁺ transients in urothelial cells. (A) Sequence of fluorescent images captured at different time points during the course of an experiment. Note that mechanical stimulation of the umbrella cell causes a deformation followed by a rapid increase in the emission of GCaMP5G (arrow). The borders of the stimulated cell (ROI) are marked in red. (B) Change in fluorescence intensity (ΔF/F) over time for eight independent mechanically stimulated cells in a bladder mucosal preparation from a control mouse. The time of indentation (12.5 s) is marked with a blue arrow. The data in (B) have been modified from Dalghi et al.³¹. Please click here to view a larger version of this figure.

Subscription Required. Please recommend JoVE to your librarian.

Discussion

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 channels have been largely restricted to dissociated cells and cell lines. Since many mechanically activated ion channels are permeable to Ca²⁺, we took advantage of recent advances in microscopy, biosensing, and micromanipulation to develop an imaging method to assess the function of mechanically activated channels in an ex vivo urothelial preparation. In this protocol, urothelial cells are transduced with an adenovirus encoding for GCaMP5G. Adenoviral vectors are ideally suited for studying Ca²⁺ signaling in the urothelium as they provide a high rate of transduction and high levels of expression of GCaMP5G in urothelial cells (Figure 2D). Under these conditions, background fluorescence is relatively low. If adenoviral transduction is used to express a genetically encoded Ca²⁺ sensor, special emphasis should be given to optimizing the conditions, so that only the cells of interest express the sensor. As an alternative, expression of genetically encoded Ca²⁺ indicators (GECIs) could be achieved by crossing mice carrying a floxed transgene encoding for the protein of interest (e.g., GCaMP5G) and mice expressing Cre recombinase under the control of a suitable promotor. This approach does not require surgical procedures and has the advantage of providing targeted expression and even levels of expression of proteins in cells expressing Cre recombinase.

The protocol described here uses an upright wide-field microscope to assess mechanically activated Ca²⁺ transients in umbrella cells stimulated with a closed fire-polished micropipette. The method could be adapted to study mechanotransduction in deeper layers of the urothelium or even cells in the lamina propria with an upright confocal or two-photon microscope. Using a confocal microscope may provide the necessary resolution to define subcellular changes in intracellular Ca²⁺ that occur in response to mechanical stimulation. This protocol employs a closed fire-polished micropipette to poke individual umbrella cells. The main advantage of using a micropipette for stimulation is that the mechanical perturbation exerted on the tissue is transitory and limited to the cell of interest and the surrounding area. Whereas cell stretching systems can be used to mechanically stimulate cells grown on PDMS membranes, inherent limitations impede the use of such devices for imaging experiments with ex vivo preparations. These limitations include challenges with mounting and imaging a mouse bladder and, given the folded nature of the bladder mucosa, difficulties in maintaining focus on the cells of interest while stretching the tissue.

The protocol to measure mechanically activated Ca²⁺ transients in urothelial preparations has some limitations. First, the rig includes relatively expensive components, and its assembly and setup require expertise with microscopes, micromanipulators, analog-digital converters, and relatively specialized software. Like patch-clamping, this technique requires the investigator to learn how to use a micromanipulator and make glass micropipettes for mechanical stimulation. However, unlike the patch-clamp technique, which involves the formation of a high resistance seal between the pipette and the membrane, the procedure described here to measure mechanically activated Ca²⁺ transients is relatively simple and only requires the investigator to position the stimulatory pipette close to the surface of the cell to be stimulated. Therefore, a trained investigator can potentially assess the response to the poking of 8-10 cells in 1 hour, which is inconceivable with patch-clamp technique.

Given the importance that mechanically activated ion channels have in normal body function and disease states, new methods are needed to study these channels in their native milieu. As described in this protocol, imaging methods can provide unique insight into how these channels sense changes in their environment and generate physiological responses. Given the accessibility of the mucosal surface of the tube- or sac-shaped organs, the method described here could be adapted to study mechanotransduction in other settings, including the gut, urogenital tract, blood vessels, etc. Thus, studying mechanical responses in the body may be broadly useful.

Subscription Required. Please recommend JoVE to your librarian.

Disclosures

The authors have nothing to disclose.

Acknowledgments

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).

Materials

Name	Company	Catalog Number	Comments
20x Objective	Olympus	UMPlanFL N
24 G ¾” catheter	Medline	Suresite IV slide
4x Objective	Olympus	UPlanFL N
Analog/digital converter	Molecular Devices	Digidata 1440A
Anti-GFP antibody	Abcam	Ab6556
Beam splitter	Chroma	T495lpxr
Bipolar temperature controller	Warner Instruments	TC-344B
CaCl₂	Fluka	21114-1L	1 M solution
cellSens software	Olympus		Imaging software
CMOS camera	Hamamatsu	ORCA fusion
Donkey anti-rabbit conjugated to Alexa Fluor 488	Jackson ImmunoResearch	711-545-152
Excel	Microsoft Corporation
Filter	Chroma	ET470/40X
Glass capillaries Corning 8250 glass	Warner Instruments	G85150T-4
Glucose	Sigma	G8270
HEPES	Sigma	H4034
Inline heater	Warner Instruments	SH-27B
KCl	Sigma	793590
Light source	Sutter Instruments	Lambda XL
Manifold pump tubing	Fisherbrand	14-190-510	ID 1.52 mm
Manifold pump tubing	Fisherbrand	14-190-533	ID 2.79 mm
MgCl₂	Sigma	M9272
Mice	Jackson Lab	664	2-4 months old female C57BL/6J
Microforge	Narishige	MF-830
Micromanipulator	Sutter Instruments	MP-285
Microscope	Olympus	BX51W
Mounting media with DAPI	Invitrogen	S36964	Slowfade Diamond Antifade with DAPI
NaCl	Sigma	S7653
pClamp software	Molecular Devices	Version 10.4	Patch-clamp electrophysiology data acquisition and analysis software
Peristaltic pump	Gilson	Minipuls 3
Piezoelectric actuator	Thorlabs	PAS005
Pipette holder	World Precision Instruments
Pipette puller	Narishige	PP-830
Quick exchange heated base with perfusion and adapter ring kit	Warner Instruments	QE-1	Quick exchange platform fits 35 mm dish
Rhodamine-phalloidin	Invitrogen	R415
Sigma-Plot	Systat Software Inc	Version 14.0	Scientific graphing and data analysis software
Silicone elastomer	Dow	Sylgard 184
Single channel open-loop piezo controller	Thorlabs	MDT694B
Square grid holder pad	Ted Pella	10520
Suture	AD Surgical	S-S618R13	6-0 Sylk
Teflon mounting rod	Custom made		Use to mount the piezoelectric actuator in the micromanipulator
Tubing	Fisher Scientific	14171129	Tygon S3 ID 1/16 IN, OD 1/8 IN
USB Digital I/O device	National Instruments	NI USB-6501

DOWNLOAD MATERIALS LIST

References

Kunau, R. T., Webb, H. L., Borman, S. C. Characteristics of the relationship between the flow rate of tubular fluid and potassium transport in the distal tubule of the rat. Journal of Clinical Investigation. 54 (6), 1488-1495 (1974).
Engbretson, B. G., Stoner, L. C. Flow-dependent potassium secretion by rabbit cortical collecting tubule in vitro. American Journal of Physiology. 253 (5), 896-903 (1987).
Satlin, L. M., Sheng, S., Woda, C. B., Kleyman, T. R. Epithelial Na(+) channels are regulated by flow. American Journal of Physiology Renal Physiology. 280 (6), 1010-1018 (2001).
Woda, C. B., et al. Ontogeny of flow-stimulated potassium secretion in rabbit cortical collecting duct: functional and molecular aspects. American Journal of Physiology Renal Physiology. 285 (4), 629-639 (2003).
Malnic, G., Berliner, R. W., Giebisch, G. Flow dependence of K+ secretion in cortical distal tubules of the rat. American Journal of Physiology. 256 (5), 932-941 (1989).
Khuri, R. N., Strieder, W. N., Giebisch, G. Effects of flow rate and potassium intake on distal tubular potassium transfer. American Journal of Physiology. 228 (4), 1249-1261 (1975).
Good, D. W., Wright, F. S. Luminal influences on potassium secretion: sodium concentration and fluid flow rate. American Journal of Physiology. 236 (2), 192-205 (1979).
Wong, K. R., Berry, C. A., Cogan, M. G. Flow dependence of chloride transport in rat S1 proximal tubules. American Journal of Physiology. 269 (6), 870-875 (1995).
Garvin, J. L. Glucose absorption by isolated perfused rat proximal straight tubules. American Journal of Physiology. 259 (4), 580-586 (1990).
Malnic, G., Klose, R. M., Giebisch, G. Micropuncture study of renal potassium excretion in the rat. American Journal of Physiology. 206 (4), 674-686 (1964).
Malnic, G., Klose, R. M., Giebisch, G. Micropuncture study of distal tubular potassium and sodium transport in rat nephron. American Journal of Physiology. 211 (3), 529-547 (1966).
Cabral, P. D., Garvin, J. L. Luminal flow regulates NO and O2(-) along the nephron. American Journal of Physiology. 300 (5), 1047-1053 (2011).
Raghavan, V., Rbaibi, Y., Pastor-Soler, N. M., Carattino, M. D., Weisz, O. A. Shear stress-dependent regulation of apical endocytosis in renal proximal tubule cells mediated by primary cilia. Proceedings of the National Academy of Sciences. 111 (23), 8506-8511 (2014).
Lewis, S. A., de Moura, J. L. Apical membrane area of rabbit urinary bladder increases by fusion of intracellular vesicles: an electrophysiological study. The Journal of Membrane Biology. 82 (2), 123-136 (1984).
Fowler, C. J., Griffiths, D., de Groat, W. C. The neural control of micturition. Nature Reviews Neuroscience. 9 (6), 453-466 (2008).
Dalghi, M. G., Montalbetti, N., Carattino, M. D., Apodaca, G. The urothelium: life in a liquid environment. Physiological Reviews. 100 (4), 1621-1705 (2020).
Khandelwal, P., Abraham, S. N., Apodaca, G. Cell biology and physiology of the uroepithelium. American Journal of Physiology Renal Physiology. 297 (6), 1477-1501 (2009).
Sachs, F. Stretch-activated ion channels: what are they. Physiology. 25 (1), 50-56 (2010).
Martinac, B. Mechanosensitive ion channels: molecules of mechanotransduction. Journal of Cell Science. 117 (12), 2449-2460 (2004).
Ranade, S. S., Syeda, R., Patapoutian, A. Mechanically activated ion channels. Neuron. 87 (6), 1162-1179 (2015).
Cox, C. D., Bavi, N., Martinac, B. Biophysical principles of ion-channel-mediated mechanosensory transduction. Cell Reports. 29 (1), 1-12 (2019).
Carattino, M. D., Sheng, S., Kleyman, T. R. Epithelial Na+ channels are activated by laminar shear stress. Journal of Biological Chemistry. 279 (6), 4120-4126 (2004).
Ross, T. D., et al. Integrins in mechanotransduction. Current Opinion in Cell Biology. 25 (5), 613-618 (2013).
Dieterle, M. P., Husari, A., Rolauffs, B., Steinberg, T., Tomakidi, P. Integrins, cadherins and channels in cartilage mechanotransduction: perspectives for future regeneration strategies. Expert Reviews in Molecular Medicine. 23, 14 (2021).
Huveneers, S., de Rooij, J. Mechanosensitive systems at the cadherin-F-actin interface. Journal of Cell Science. 126, 403-413 (2013).
Sun, Z., Guo, S. S., Fässler, R. Integrin-mediated mechanotransduction. Journal of Cell Biology. 215 (4), 445-456 (2016).
Hamill, O. P., Marty, A., Neher, E., Sakmann, B., Sigworth, F. J. Improved patch-clamp techniques for high-resolution current recording from cells and cell-free membrane patches. Pflügers Archiv. European Journal of Physiology. 391 (2), 85-100 (1981).
Akerboom, J., et al. Optimization of a GCaMP calcium indicator for neural activity imaging. The Journal of Neuroscience. 32 (40), 13819-13840 (2012).
Akerboom, J., et al. Genetically encoded calcium indicators for multi-color neural activity imaging and combination with optogenetics. Frontiers in Molecular Neuroscience. 6, 2 (2013).
Sun, X. R., et al. Fast GCaMPs for improved tracking of neuronal activity. Nature Communications. 4, 2170 (2013).
Dalghi, M. G., et al. Functional roles for PIEZO1 and PIEZO2 in urothelial mechanotransduction and lower urinary tract interoception. JCI Insight. 6 (19), 152984 (2021).
Durnin, L., et al. An ex vivo bladder model with detrusor smooth muscle removed to analyse biologically active mediators released from the suburothelium. The Journal of Physiology. 597 (6), 1467-1485 (2019).
Delmas, P., Coste, B. Mechano-gated ion channels in sensory systems. Cell. 155 (2), 278-284 (2013).
Tavernarakis, N., Driscoll, M. Degenerins. At the core of the metazoan mechanotransducer. Annals of the New York Academy of Sciences. 940 (1), 28-41 (2001).
Peyronnet, R., Tran, D., Girault, T., Frachisse, J. M. Mechanosensitive channels: feeling tension in a world under pressure. Frontiers in Plant Science. 5, 558 (2014).
Blount, P., Iscla, I. Life with bacterial mechanosensitive channels, from discovery to physiology to pharmacological target. Microbiology and Molecular Biology Reviews. 84 (1), 00055 (2020).
Booth, I. R., Miller, S., Müller, A., Lehtovirta-Morley, L. The evolution of bacterial mechanosensitive channels. Cell Calcium. 57 (3), 140-150 (2015).

Medicine